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Plant organelle positioning
Plant organelle positioning Masamitsu Wada1,2 and Noriyuki Suetsugu2 Correct positioning and active movement of organelles within cells are essential for cellular homeostasis and adaptation to external stresses. Unlike animal and fungal systems, plant organelle positioning has not yet been revealed at the molecular level. The recent development of organelle-targeting green fluorescent protein (GFP) constructs and genetic analyses using Arabidopsis thaliana have shed new light on the field of plant organelle positioning, which has been found to be regulated by mechanisms that are similar to and/or distinct from those used by animals and fungi. Addresses 1 Department of Biological Sciences, Graduate School of Science, Tokyo Metropolitan University, Tokyo 192-0397, Japan e-mail: [email protected] 2 Division of Biological Regulation and Photobiology, National Institute for Basic Biology, Okazaki 444-8585, Japan
Current Opinion in Plant Biology 2004, 7:626–631 This review comes from a themed issue on Cell biology Edited by Martin Hu¨skamp and Yasunori Machida Available online 25th September 2004 1369-5266/$ – see front matter # 2004 Elsevier Ltd. All rights reserved. DOI 10.1016/j.pbi.2004.09.005
Abbreviations AF actin filament chup1 chloroplast unusual positioning1 cluA clustered mitochondriaA ER endoplasmic reticulum fmt friendly mitochondrial GFP green fluorescent protein LAT-B latrunculin-B MT microtubule NEM N-ethylmaleimide PTS peroxisome-targeting signal
Introduction In general, plants are sessile organisms; even when they are exposed to stress conditions such as strong light, high temperature, desiccation, physical stress, and pathogen attack, they cannot escape and have to overcome the stresses. Hence, plants evolved sophisticated adaptive responses at the organelle level as well as at the cell, tissue, and organ levels to avoid stresses [1,2]. Intracellular organelle positioning (i.e. distributing organelles appropriately within cells) plays an important role in allowing plants to overcome environmental stresses and to proliferate life efficiently. Each organelle species must Current Opinion in Plant Biology 2004, 7:626–631
have its own efficient mechanism for intracellular positioning in response to environmental changes, but harmonic coordination of the positions of the various organelles is also indispensable in allowing each organelle to accomplish its function. To gain the proper position in a cell, organelles should move to the preferable sites and fix themselves in the position. Analyses of organelle positioning in various plant species have revealed that two types of cytoskeleton, microtubules (MTs) and/or actin filaments (AFs), mediate organelle movement and its anchoring, although AFs are predominantly utilized for organelle movements in higher plants [3]. The movement of nuclei and chloroplasts has been well studied for some time because these organelles are easily distinguished by their size and color, respectively, under microscopy. Chloroplast movement has been especially well studied, not only because the movement of these organelles is easily observed but also because their movement can be induced by environmental factors, light and mechanical stress (Figure 1; [4]). Recent technical advances in utilizing green fluorescent protein (GFP) genes that are targeted to various organelles, such as mitochondria [5,6], peroxisomes [7–10], and Golgi stacks [11,12], have opened new areas of research into the positioning of plant organelles (Figure 1). In this review, we discuss recent analyses of plant organelle movement, including studies of the movement of chloroplasts, mitochondria and peroxisomes. We also discuss the movement of Golgi stacks in mature cells but not in young growing cells such as tip-growing root-hair cells, pollen tubes, rhizoids and the protonemal cells of ferns and mosses [13,14]. The movement and positioning of nuclei are not reviewed here because no new results are available. We also omit the inheritance of organelles during cell division because organelle distribution during cell division has been well studied and documented elsewhere [15], as has organelle distribution within tip-growing cells [13,14]. Instead, we focus on the movement and positioning of organelles in fully developed cells to maximize their function.
CHUP1, a possible link between chloroplasts and AFs during chloroplast movement The photorelocation of chloroplasts is the most-characterized organelle movement in plants. Light of low fluence rate induces the movement of chloroplasts towards light, whereas light of high fluence rate causes chloroplasts to move away from the strong light. Most plant www.sciencedirect.com
Plant organelle positioning Wada and Suetsugu 627
Figure 1
Wildtype
(a)
Chloroplasts
(b)
Mitochondria
(c)
Peroxisomes
Chup1
Distribution of chloroplasts, mitochondria and peroxisomes in wildtype and chup1 mutants. (a) Cross-section of a light-adapted leaf. In the wildtype, chloroplasts are positioned on the upper and lower cell surfaces, whereas in the cells of chup1 mutants, chloroplasts are aggregated on cell bottom. Bar represents 30 mm. (b) Mitochondrial distribution viewed by transient expression of mitochondria-targeting GFP. The positioning of mitochondria is similar in wildtype and chup1 mutants. Bar represents 10 mm. (c) Peroxisomal positioning viewed by transient expression of peroxisome-targeting GFP. Peroxisomes are localized close to chloroplasts. In chup1 mutant cells, peroxisomes are closely associated with aggregated chloroplasts. Bar represents 10 mm.
species utilize AFs for chloroplast movement, but the MT system is also involved in chloroplast movement in a few lower plant species [4]. Recently, chloroplast unusual positioning1 (chup1) mutants were isolated as Arabidopsis thaliana mutants whose chloroplasts have a defective light-avoidance response [16]. In chup1 mutants, chloroplasts aggregated aberrantly on the cell bottom regardless of the light conditions (Figure 1), a distribution pattern resembling chloroplast aggregation in mesophyll cells of A. thaliana treated with the anti-AF drug latrunculin-B (LAT-B) [17]. In spite of the severe chloroplast-positioning phenotype, the movement or positioning of other organelles (i.e. nuclei, mitochondria, and peroxisomes) in chup1 mutants was similar to that in wildtype plants (Figure 1). The CHUP1 gene encodes a protein that comprises multiple domains, including a coiled-coil domain, an actin-binding www.sciencedirect.com
domain, a proline-rich motif (PRM) and two leucinezipper domains. A fusion of glutathione S-transferase with the actin-binding domain of CHUP1 is capable of binding to F-actin in vitro, indicating that CHUP1 is an actin-binding protein [16]. Moreover, CHUP1 contains a PRM, a domain that is known to recruit profilactin (a complex of profilin and G-actin) during F-actin polymerization [18]. Immunolabeling with actin monoclonal antibody showed that chloroplasts aligned along the thick AFs but not along MTs, and were enclosed within fine AFs [17]. Visualization of in-vivo AFs using GFP–mouse talin fusions revealed, however, that the pattern of AFs in the cells of chup1 mutants is similar to that in wildtype cells. Importantly, when the GFP fused to the carboxy-terminus end of the amino-terminal hydrophobic segment of CHUP1 (amino acid 1–25) was transiently expressed in onion epidermal cells, and in Arabidopsis epidermal and mesophyll cells, the fusion proteins were localized on chloroplast peripheries [16]. This localization pattern was similar to that of A. thaliana OUTER ENVELOPE PROTEIN7 (AtOEP7)–GFP, which is known to be targeted to the chloroplast outer membrane [19]. Together, the chup1 mutant phenotype, and the localization and actin-binding of the CHUP1 protein strongly suggest that CHUP1 could regulate actin polymerization on chloroplasts (Figure 2). Although chup1 mutants are defective in chloroplast photorelocation movement, their chloroplast distribution is markedly different from that of phot1 phot2 double mutants, which are defective in two phototropins (the blue light receptors for chloroplast photorelocation movement) [20]. The chloroplasts of chup1 mutants are aggregated on bottom of cells and often do not attach to the cell surface, whereas in phot1 phot2 double mutants, chloroplasts are distributed randomly and are associated with the cell surface. In the wildtype cells, all of the chloroplasts that show photorelocation movement attach to the cell surface in just one layer. Hence, CHUP1 proteins are likely to be necessary for the correct positioning of chloroplasts on the cell surface but not for chloroplast motility itself (Figure 2). For chloroplasts to move, some kind of connection between the chloroplast and the cell surface must be necessary. The pattern of chloroplast aggregation that occurs in chup1 mutants is detrimental to survival under natural conditions. Kasahara et al. [21] showed that the chloroplast-avoidance response is essential to allow escape from photodamage under strong light conditions. When continuously irradiated with white light at 1400 mmol m 2 s 1 (the typical intensity of sunlight in summer), chup1 mutants were photo-bleached and eventually died. Hence, the correct positioning or movement of chloroplasts is an essential adaptation for the survival of plants in natural environments. Current Opinion in Plant Biology 2004, 7:626–631
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Figure 2
Chloroplast
CHUP1 Plasma membrane G-actin
CHUP1 domain
Profilin
Hydrophobic-region
Profilactin
Actin-binding
Myosin
Proline-rich Current Opinion in Plant Biology
Working model of chloroplast positioning. The amino-terminal hydrophobic region of CHUP1 can target to the chloroplast outer membrane. The proline-rich motif of CHUP1 may serve in actin polymerization to recruit profilactin. CHUP1 binds polymerized F-actin through its actin-binding domain. These functions of CHUP1 may be important in anchoring chloroplasts to the plasma membrane. Myosin motor protein(s) may be necessary for chloroplast motility along actin filaments.
A conserved mechanism regulates mitochondrial positioning in higher plants and fungi The basic mechanism of mitochondrial morphology and positioning was elucidated from extensive research using yeast or animals [22]. However, higher plant mitochondria are different from those of yeast or animals in their morphology and motility system [5,6]. For example, mitochondrial transport is dependent on MTs in yeast and animals [22], whereas it depends on AFs in higher plants [6]. Therefore, because some of the mechanisms that control mitochondrial transport or distribution may be plant-specific, molecular genetic analysis in plants is indispensable to an improved understanding of plant mitochondrial positioning. By screening mutagenized Arabidopsis seedlings expressing GFP targeted to mitochondria, Logan et al. [23] isolated five mutants that are defective in mitochondrial morphology or distribution [23]. Among them, the friendly mitochondrial (fmt) mutant has some clusters consisting of tens of mitochondria. Many other nonclustered mitochondria in fmt mutants behaved like those in wildtype cells. Positional cloning of the FMT gene showed that it encodes an Arabidopsis homolog of the Dictyostelium discoideum clustered mitochondriaA (cluA) gene, whose disruption in D. discoideum causes the aggregation of mitochondria near the cell center [24]. The CLU1 gene, a Saccharomyces cerevisiae homolog of the FMT and cluA genes, is also involved in mitochondrial distribution [25], indicating that the function of FMTlike genes is conserved in higher plants and fungi. HowCurrent Opinion in Plant Biology 2004, 7:626–631
ever, as the mitochondria of D. discoideum cluA disruptants are interconnected by membraneous strands [26] but those of A. thaliana fmt mutants are not [23], the cluA and FMT genes may function differently in mitochondrial positioning. The FMT protein has tetratricopeptide repeat domain, which is known as a protein–protein interaction domain. At present, the in-vivo function of the FMT gene during mitochondrial positioning remained to be determined; the sub-cellular localization and interacting partner of the FMT protein are unknown. In D. discoideum, drug treatment that disrupts AFs or MTs cannot mimic the cluA disruptant phenotype, suggesting that the cluA gene may function independently of the cytoskeleton [26]. Treatment of tobacco cultured cells with the anti-actin drug LAT-B, the myosin ATPase inhibitor 2,3-butanedione and the sulfhydryl-modifying agent N-ethylmaleimide (NEM) (but not with the MT-depolymerizing drug oryzaline) induced clustered mitochondria and inhibited mitochondrial movement [6], causing a morphology that partially resembled the fmt mutant phenotype. Further analyses of fmt and other mutants will shed light on mechanism of the involvement of AFs and FMT protein in mitochondrial positioning.
Acto-myosin-dependent peroxisomal movement Peroxisomes have multiple important functions in plants, such as fatty-acid b-oxidation, photorespiration, and peroxisomal biogenesis, and these functions have been studied extensively using A. thaliana [27]. Moreover, the www.sciencedirect.com
Plant organelle positioning Wada and Suetsugu 629
peroxisome-targeting signals (PTS), the carboxy-terminal Ser-Lys-Leu tripeptide (PTS-1) and the amino-terminal (R/K)(L/V/I)(X)5(H/Q)(L/A) nonapeptide (PTS-2), are well-known among eukaryotes. Using PTS-fused GFP, four groups independently discovered that peroxisomes move actively in various cell types of A. thaliana [8–10] and in the epidermal cells of onion [10] or leek [7]. The velocity and motility of peroxisomes differ according to the plant cell-types examined: movement can involve slow or rapid directional movement, random oscillations or stop-and-go movements [7–10]. AF-depolymerizing drugs (e.g. LAT-B etc.) inhibit peroxisome movement but MT-depolymerizing drugs do not, indicating that peroxisome movement is dependent on AF but not on MT [7–10]. Co-localization studies have revealed that peroxisomes are closely associated with AFs but not with MTs [7,8,10]. As the myosin inhibitors 2,3-butanedione and NEM also inhibit peroxisome movement, the actomyosin system may be involved in peroxisome movement. Leaf peroxisomes often localize near chloroplasts, probably to maximize photorespiration [9]. It is worth noting that peroxisomes are located at the bottom of the cells of chup1 mutants, alongside the aggregated chloroplasts (Figure 1; [16]). This evidence suggests that the peroxisome–chloroplast association is strong, and that the distribution of peroxisomes is partially dependent on chloroplast movement, at least in leaf cells. Although many mutants that have defective peroxisome functions have been isolated, mutants that are deficient in peroxisome movement and positioning have not yet been isolated [27]. Recently, mutants that are defective in peroxisome morphology were screened by visual inspection of mutagenized GFP–PTS1-transgenic seedlings [28]. A screen using the same mutagenized GFP– PTS1 seedlings has the potential to isolate mutants that are defective in peroxisome movement and positioning.
Golgi movement associated with ER networks Bidirectional coordinate protein transport between Golgi and endoplasmic reticulum (ER) is essential for cell functions, suggesting the importance of the relative positions of the Golgi and ER. When GFP constructs targeted to both Golgi and ER were expressed in leaves of Nicotiana clevelandii, Golgi stacks were shown to move actively in close association with the ER tubules [11]. The application of the anti-actin drug cytochalasin D and the myosin inhibitor NEM inhibited Golgi movement. Importantly, Golgi bodies colocalize with the AF network near ER, suggesting that Golgi movement along ER depends on the acto-myosin system [11]. Nebenfu¨ hr et al. [12] further investigated Golgi motility in tobacco cultured cells and found that Golgi stacks repeated rapid vectorial movements and had a wiggling state, i.e. ‘stopand-go’ movement, which was also shown to be dependent on acto-myosin [12]. When Golgi stacks moved along the same track, they often stopped at the same www.sciencedirect.com
point. Therefore, Nebenfu¨ hr et al. [12] hypothesized that specific sites, such as active ER export sites or areas of cell-wall growth, generate a ‘stop signal’. Interestingly, the expression of a dominant-negative mutation of the Arabidopsis Rab GTPase AtRab1b, AtRab1b (N121I), disrupted the normal movement of Golgi stacks on the ER networks in tobacco [29]. Although the expression of wildtype AtRab1b did not affect Golgi movement, coexpression of wildtype AtRab1b with AtRab1b (N121I) rescued the dominant-negative effect of AtRab1b (N121I), suggesting that AtRab1b activity is necessary for plant Golgi movement. Members of this subclass of Rab GTPases have been implicated in vesicular trafficking in the secretary pathway in yeast, mammals and plants [30,31]. This novel function of AtRab1b suggests an exciting relationship between Golgi movement and vesicular trafficking. A recent paper demonstrated that aggregation of ER networks and Golgi accumulation occurred at the infection site of oomycete pathogen in A. thaliana [32]. At oomycete penetration sites, AFs but not MTs dynamically re-organized and formed large bundles. Takemoto et al. [32] suggested that the accumulation of ER at the penetration site serves as the ‘stop signal’ for Golgi movement, resulting in Golgi accumulation at the penetration site. ER–Golgi accumulation at the pathogen infection site may serve as secretion site for pathogen resistance.
Conclusions Organelle movements in seed plants are mediated mostly by AFs rather than by MTs, although most of the work carried out to date has involved inhibitor studies that have not provided detailed information about mechanisms. Precise studies on CHUP1 proteins that are localized on the chloroplast outer envelope may provide a strong clue to shed light on the mechanism of organelle movements, at least on chloroplast movement (Figure 2; [16]). The mechanism of movements may not be so difficult to determine because the involvement of AFs is clear. The most important questions are how organelles recognize where to go and stay, and how organelles are fixed at their appropriate positions. During chloroplast positioning within the cells of fern gametophytes [33,34] and Arabidopsis mesophyll cells [17], ring-like structures made of AFs appear and seem to connect chloroplasts to the cell surface when they have reached their position. We do not know, however, whether the ring-like AFs bind chloroplasts to the cell surface to anchor them. Recently, western blot and immunofluorescence studies using the maize myosin XI antibody have revealed that maize myosin XI co-localizes with mitochondria and plastids but not with nuclei, Golgi, ER, and peroxisomes [35]. Therefore, chloroplast or mitochondrial movement may be mediated by the acto-myosin system (Figure 2). Although no information is available yet about the posCurrent Opinion in Plant Biology 2004, 7:626–631
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sible association of intermediate filaments or cytoskeletal proteins with the cytoplasmic face in plants, the recent identification of a bacterial intermediate-filament-like protein opened the possibility that plants also have intermediate filaments [36,37]. The fine structure underneath the plasma membrane and the genes that are involved in constructing this structure must be studied if we are to understand organelle movement and positioning.
Acknowledgements This work was partly supported by Grand-in-Aid for Scientific Research (on Priority Areas, no.s 13139203 and 13304061) from the Ministry of Education, Sports, Science and Technology (MEXT) of Japan, and Grant-in-Aid for Scientific Research (no. 16107002) from the Japan Society for the Promotion of Science to MW, and by a grant from the Research Fellowships of the Japan Society for the Promotion of Science for Young Scientists to NS.
References and recommended reading Papers of particular interest, published within the annual period of review, have been highlighted as: of special interest of outstanding interest 1.
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4. Wada M, Kagawa T, Sato Y: Chloroplast movement. Annu Rev Plant Biol 2003, 54:455-468. An excellent recent review on chloroplast movement. 5.
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Logan DC, Leaver CJ: Mitochondria-targeted GFP highlights the heterogeneity of mitochondrial shape, size and movement within living plant cells. J Exp Bot 2000, 51:865-871. Van Gestel K, Ko¨ hler RH, Verbelen J-P: Plant mitochondria move on F-actin, but their positioning in the cortical cytoplasm depends on both F-actin and microtubules. J Exp Bot 2002, 53:659-667.
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Collings DA, Harper JDI, Marc J, Overall RL, Mullen RT: Life in the fast lane: actin-based motility of plant peroxisomes. Can J Bot 2002, 80:430-441.
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Jedd G, Chua N-H: Visualization of peroxisomes in living plant cells reveals acto-myosin-dependent cytoplasmic streaming and peroxisome budding. Plant Cell Physiol 2002, 43:384-392.
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Mano S, Nakamori C, Hayashi M, Kato A, Kondo M, Nishimura M: Distribution and characterization of peroxisomes in Arabidopsis by visualization with GFP: dynamic morphology and actin-dependent movement. Plant Cell Physiol 2002, 43:331-341.
10. Mathur J, Mathur N, Hu¨ lskamp M: Simultaneous visualization of peroxisomes and cytoskeletal elements reveals actin-based and not microtubule-based peroxisome motility in plants. Plant Physiol 2002, 128:1031-1045. 11. Boevink P, Oparka K, Santa Cruz S, Martin B, Betteridge A, Hawes C: Stacks on tracks: the plant Golgi apparatus traffics on an actin/ER network. Plant J 1998, 15:441-447.
14. Wasteneys GO, Galway ME: Remodeling the cytoskeleton for growth and form: an overview with some new views. Annu Rev Plant Biol 2003, 54:691-722. An extensive recent review of the role of the cytoskeleton in plant cell growth. 15. Sheahan MB, Rose RJ, McCurdy DW: Organelle inheritance in plant cell division: the actin cytoskeleton is required for unbiased inheritance of chloroplasts, mitochondria and endoplasmic reticulum in dividing protoplasts. Plant J 2004, 37:379-390. The authors analyzed organelle movement during the cell division of tobacco protoplasts in detail. They showed that actin filaments but not microtubules were necessary for the dynamic re-distribution of chloroplasts, mitochondria and ER. 16. Oikawa K, Kasahara M, Kiyosue T, Kagawa T, Suetsugu N, Takahashi F, Kanegae T, Niwa Y, Kadota A, Wada M: CHLOROPLAST UNUSUAL POSITIONING1 is essential for proper chloroplast positioning. Plant Cell 2003, 15:2805-2815. The authors isolated a chloroplast-positioning mutant, chup1. The CHUP1 gene encodes a novel protein that has a coiled-coil domain, an actin-binding domain and two-leucine zipper domains. The actinbinding domain of CHUP1 can bind to F-actin in vitro. A fusion of the amino-terminal hydrophobic region of CHUP1 and GFP targeted to the chloroplast outer membrane. These results suggest that CHUP1 may function as a chloroplast-anchoring protein that links with actin filaments. 17. Kandasamy MK, Meagher RB: Actin-organelle interaction: association with chloroplast in Arabidopsis leaf mesophyll cells. Cell Motil Cytoskeleton 1999, 44:110-118. 18. Holt MR, Koffer A: Cell motility: proline-rich proteins promote protrusions. Trends Cell Biol 2001, 11:38-46. 19. Lee YJ, Kim DH, Kim Y-W, Hwang I: Identification of a signal that distinguishes between the chloroplast outer envelope membrane and the endomembrane system in vivo. Plant Cell 2001, 13:2175-2190. 20. Sakai T, Kagawa T, Kasahara M, Swartz TE, Christie JM, Briggs WR, Wada M, Okada K: Arabidopsis nph1 and npl1: blue light receptors that mediate both phototropism and chloroplast relocation. Proc Natl Acad Sci USA 2001, 98:6969-6974. 21. Kasahara M, Kagawa T, Oikawa K, Suetsugu N, Miyao M, Wada M: Chloroplast avoidance movement reduces photodamage in plants. Nature 2002, 420:829-832. 22. Yaffe MP: The machinery of mitochondrial inheritance and behaviour. Science 1999, 283:1493-1497. 23. Logan DC, Scott I, Tobin AK: The genetic control of plant mitochondrial morphology and dynamics. Plant J 2003, 36:500-509. The authors isolated mutants that are defective in mitochondrial morphology and distribution by screening mutagenized seedlings whose mitochondria were labeled with GFP. The fmt mutant showed partially aggregated mitochondria. The FMT gene is a homologue of the D. discoideum cluA and S. cerevisiae CLU1 genes, whose disruptants also showed mitochondrial aggregation. Therefore, this result indicates that the mechanism of mitochondrial positioning is conserved between higher plants and fungi. 24. Zhu Q, Hulen D, Liu T, Clarke M: The cluAS mutant of Dictyostelium identifies a novel class of proteins required for dispersion of mitochondria. Proc Natl Acad Sci USA 1997, 94:7308-7313. 25. Fields SD, Conrad MN, Clarke M: The S. cerevisiae CLU1 and D. discoideum cluA genes are functional homologues that influence mitochondrial morphology and distribution. J Cell Sci 1998, 111:1717-1727. 26. Fields SD, Arana Q, Heuser J, Clarke M: Mitochondrial membrane dynamics are altered in cluAS mutants of Dictyostelium. J Muscle Res Cell Motil 2002, 23:829-838.
12. Nebenfu¨ hr A, Gallagher LA, Dunahay TG, Frohlick JA, Mazurkiewicz AM, Meehl JB, Staehelin LA: Stop-and-go movements of plant Golgi stacks are mediated by the acto-myosin system. Plant Physiol 1999, 121:1127-1141.
27. Hayashi M, Nishimura M: Entering a new era of research on plant peroxisomes. Curr Opin Plant Biol 2003, 6:577-582.
13. Hepler PK, Vidali L, Cheung AY: Polarized cell growth in higher plants. Annu Rev Cell Dev Biol 2001, 17:159-187.
28. Mano S, Nakamori C, Kondo M, Hayashi M, Nishimura M: An Arabidopsis dynamin-related protein, DRP3A, controls
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both peroxisomal and mitochondrial division. Plant J 2004, 38:487-498. The authors screened mutagenized GFP–PTS1 seedlings under fluorescent microscopy to isolate peroxisomal morphology mutants. This screening method will allows mutants that are defective in peroxisomal movement or positioning to be isolated visually. 29. Batoko H, Zheng H-Q, Hawes C, Moore I: A Rab1 GTPase is required for transport between the endoplasmic reticulum and Golgi apparatus and for normal Golgi movements in plants. Plant Cell 2000, 12:2201-2217. 30. Zerial M, McBride H: Rab proteins as membrane organizers. Nat Rev Mol Cell Biol 2001, 2:107-117. 31. Rutherford S, Moore I: The Arabidopsis Rab GTPase family: another enigma variation. Curr Opin Plant Biol 2002, 5:518-528. 32. Takemoto D, Jones DA, Hardham AR: GFP-tagging of cell components reveals the dynamics of subcellular reorganization in response to infection of Arabidopsis by oomycete pathogens. Plant J 2003, 33:775-792. The authors showed that actin filaments, ER and Golgi were dynamically re-organized during oomycete infection, resulting in the accumulation of ER and Golgi at the penetration site of oomycete pathogens. They show, therefore, that Golgi movement occurs not only under the normal conditions but also in response to pathogen attack, indicating that Golgi movement is active and adaptive.
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33. Kadota A, Wada M: Photoinduction of circular F-actin on chloroplast in a fern protonemal cell. Protoplasma 1989, 151:171-174. 34. Kadota A, Wada M: Photoinduction of formation of circular structures by microfilaments on chloroplasts during intracellular orientation in protonemal cells of the fern Adiantum capillus-veneris. Protoplasma 1992, 167:97-107. 35. Wang Z, Pesacreta TC: A subclass of myosin XI is associated with mitochondria, plastids, and the molecular chaperone subunit TCP-1a in maize. Cell Motil Cytoskeleton 2004, 57:218-232. The authors investigated the sub-cellular localization of myosin XI in maize cells. Using a maize myosin XI antibody, they showed that myosin XI was present in plastids and mitochondria but not in nuclei, Golgi, ER, and peroxisomes. 36. Ausmees N, Kuhn JR, Jacobs-Wagner C: The bacterial cytoskeleton: an intermediate filament-like function in cell shape. Cell 2003, 115:705-713. The authors showed that some bacteria have an intermediate filamentlike protein, crescentin. Although crescentin is not similar to metazoan intermediate filament proteins, it has an extended coiled-coil signature. This result suggests that some Arabidopsis proteins that have the long coiled-coil domain may function as the intermediate filaments in plants. 37. Gindullis F, Rose A, Patel S, Meier I: Four signature motifs define the first class of structurally related large coiled-coil proteins in plants. BMC Genomics 2002, 3:9.
Current Opinion in Plant Biology 2004, 7:626–631
Current Opinion in Plant Biology 2004, 7:626–631 This review comes from a themed issue on Cell biology Edited by Martin Hu¨skamp and Yasunori Machida Available online 25th September 2004 1369-5266/$ – see front matter # 2004 Elsevier Ltd. All rights reserved. DOI 10.1016/j.pbi.2004.09.005
Abbreviations AF actin filament chup1 chloroplast unusual positioning1 cluA clustered mitochondriaA ER endoplasmic reticulum fmt friendly mitochondrial GFP green fluorescent protein LAT-B latrunculin-B MT microtubule NEM N-ethylmaleimide PTS peroxisome-targeting signal
Introduction In general, plants are sessile organisms; even when they are exposed to stress conditions such as strong light, high temperature, desiccation, physical stress, and pathogen attack, they cannot escape and have to overcome the stresses. Hence, plants evolved sophisticated adaptive responses at the organelle level as well as at the cell, tissue, and organ levels to avoid stresses [1,2]. Intracellular organelle positioning (i.e. distributing organelles appropriately within cells) plays an important role in allowing plants to overcome environmental stresses and to proliferate life efficiently. Each organelle species must Current Opinion in Plant Biology 2004, 7:626–631
have its own efficient mechanism for intracellular positioning in response to environmental changes, but harmonic coordination of the positions of the various organelles is also indispensable in allowing each organelle to accomplish its function. To gain the proper position in a cell, organelles should move to the preferable sites and fix themselves in the position. Analyses of organelle positioning in various plant species have revealed that two types of cytoskeleton, microtubules (MTs) and/or actin filaments (AFs), mediate organelle movement and its anchoring, although AFs are predominantly utilized for organelle movements in higher plants [3]. The movement of nuclei and chloroplasts has been well studied for some time because these organelles are easily distinguished by their size and color, respectively, under microscopy. Chloroplast movement has been especially well studied, not only because the movement of these organelles is easily observed but also because their movement can be induced by environmental factors, light and mechanical stress (Figure 1; [4]). Recent technical advances in utilizing green fluorescent protein (GFP) genes that are targeted to various organelles, such as mitochondria [5,6], peroxisomes [7–10], and Golgi stacks [11,12], have opened new areas of research into the positioning of plant organelles (Figure 1). In this review, we discuss recent analyses of plant organelle movement, including studies of the movement of chloroplasts, mitochondria and peroxisomes. We also discuss the movement of Golgi stacks in mature cells but not in young growing cells such as tip-growing root-hair cells, pollen tubes, rhizoids and the protonemal cells of ferns and mosses [13,14]. The movement and positioning of nuclei are not reviewed here because no new results are available. We also omit the inheritance of organelles during cell division because organelle distribution during cell division has been well studied and documented elsewhere [15], as has organelle distribution within tip-growing cells [13,14]. Instead, we focus on the movement and positioning of organelles in fully developed cells to maximize their function.
CHUP1, a possible link between chloroplasts and AFs during chloroplast movement The photorelocation of chloroplasts is the most-characterized organelle movement in plants. Light of low fluence rate induces the movement of chloroplasts towards light, whereas light of high fluence rate causes chloroplasts to move away from the strong light. Most plant www.sciencedirect.com
Plant organelle positioning Wada and Suetsugu 627
Figure 1
Wildtype
(a)
Chloroplasts
(b)
Mitochondria
(c)
Peroxisomes
Chup1
Distribution of chloroplasts, mitochondria and peroxisomes in wildtype and chup1 mutants. (a) Cross-section of a light-adapted leaf. In the wildtype, chloroplasts are positioned on the upper and lower cell surfaces, whereas in the cells of chup1 mutants, chloroplasts are aggregated on cell bottom. Bar represents 30 mm. (b) Mitochondrial distribution viewed by transient expression of mitochondria-targeting GFP. The positioning of mitochondria is similar in wildtype and chup1 mutants. Bar represents 10 mm. (c) Peroxisomal positioning viewed by transient expression of peroxisome-targeting GFP. Peroxisomes are localized close to chloroplasts. In chup1 mutant cells, peroxisomes are closely associated with aggregated chloroplasts. Bar represents 10 mm.
species utilize AFs for chloroplast movement, but the MT system is also involved in chloroplast movement in a few lower plant species [4]. Recently, chloroplast unusual positioning1 (chup1) mutants were isolated as Arabidopsis thaliana mutants whose chloroplasts have a defective light-avoidance response [16]. In chup1 mutants, chloroplasts aggregated aberrantly on the cell bottom regardless of the light conditions (Figure 1), a distribution pattern resembling chloroplast aggregation in mesophyll cells of A. thaliana treated with the anti-AF drug latrunculin-B (LAT-B) [17]. In spite of the severe chloroplast-positioning phenotype, the movement or positioning of other organelles (i.e. nuclei, mitochondria, and peroxisomes) in chup1 mutants was similar to that in wildtype plants (Figure 1). The CHUP1 gene encodes a protein that comprises multiple domains, including a coiled-coil domain, an actin-binding www.sciencedirect.com
domain, a proline-rich motif (PRM) and two leucinezipper domains. A fusion of glutathione S-transferase with the actin-binding domain of CHUP1 is capable of binding to F-actin in vitro, indicating that CHUP1 is an actin-binding protein [16]. Moreover, CHUP1 contains a PRM, a domain that is known to recruit profilactin (a complex of profilin and G-actin) during F-actin polymerization [18]. Immunolabeling with actin monoclonal antibody showed that chloroplasts aligned along the thick AFs but not along MTs, and were enclosed within fine AFs [17]. Visualization of in-vivo AFs using GFP–mouse talin fusions revealed, however, that the pattern of AFs in the cells of chup1 mutants is similar to that in wildtype cells. Importantly, when the GFP fused to the carboxy-terminus end of the amino-terminal hydrophobic segment of CHUP1 (amino acid 1–25) was transiently expressed in onion epidermal cells, and in Arabidopsis epidermal and mesophyll cells, the fusion proteins were localized on chloroplast peripheries [16]. This localization pattern was similar to that of A. thaliana OUTER ENVELOPE PROTEIN7 (AtOEP7)–GFP, which is known to be targeted to the chloroplast outer membrane [19]. Together, the chup1 mutant phenotype, and the localization and actin-binding of the CHUP1 protein strongly suggest that CHUP1 could regulate actin polymerization on chloroplasts (Figure 2). Although chup1 mutants are defective in chloroplast photorelocation movement, their chloroplast distribution is markedly different from that of phot1 phot2 double mutants, which are defective in two phototropins (the blue light receptors for chloroplast photorelocation movement) [20]. The chloroplasts of chup1 mutants are aggregated on bottom of cells and often do not attach to the cell surface, whereas in phot1 phot2 double mutants, chloroplasts are distributed randomly and are associated with the cell surface. In the wildtype cells, all of the chloroplasts that show photorelocation movement attach to the cell surface in just one layer. Hence, CHUP1 proteins are likely to be necessary for the correct positioning of chloroplasts on the cell surface but not for chloroplast motility itself (Figure 2). For chloroplasts to move, some kind of connection between the chloroplast and the cell surface must be necessary. The pattern of chloroplast aggregation that occurs in chup1 mutants is detrimental to survival under natural conditions. Kasahara et al. [21] showed that the chloroplast-avoidance response is essential to allow escape from photodamage under strong light conditions. When continuously irradiated with white light at 1400 mmol m 2 s 1 (the typical intensity of sunlight in summer), chup1 mutants were photo-bleached and eventually died. Hence, the correct positioning or movement of chloroplasts is an essential adaptation for the survival of plants in natural environments. Current Opinion in Plant Biology 2004, 7:626–631
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Figure 2
Chloroplast
CHUP1 Plasma membrane G-actin
CHUP1 domain
Profilin
Hydrophobic-region
Profilactin
Actin-binding
Myosin
Proline-rich Current Opinion in Plant Biology
Working model of chloroplast positioning. The amino-terminal hydrophobic region of CHUP1 can target to the chloroplast outer membrane. The proline-rich motif of CHUP1 may serve in actin polymerization to recruit profilactin. CHUP1 binds polymerized F-actin through its actin-binding domain. These functions of CHUP1 may be important in anchoring chloroplasts to the plasma membrane. Myosin motor protein(s) may be necessary for chloroplast motility along actin filaments.
A conserved mechanism regulates mitochondrial positioning in higher plants and fungi The basic mechanism of mitochondrial morphology and positioning was elucidated from extensive research using yeast or animals [22]. However, higher plant mitochondria are different from those of yeast or animals in their morphology and motility system [5,6]. For example, mitochondrial transport is dependent on MTs in yeast and animals [22], whereas it depends on AFs in higher plants [6]. Therefore, because some of the mechanisms that control mitochondrial transport or distribution may be plant-specific, molecular genetic analysis in plants is indispensable to an improved understanding of plant mitochondrial positioning. By screening mutagenized Arabidopsis seedlings expressing GFP targeted to mitochondria, Logan et al. [23] isolated five mutants that are defective in mitochondrial morphology or distribution [23]. Among them, the friendly mitochondrial (fmt) mutant has some clusters consisting of tens of mitochondria. Many other nonclustered mitochondria in fmt mutants behaved like those in wildtype cells. Positional cloning of the FMT gene showed that it encodes an Arabidopsis homolog of the Dictyostelium discoideum clustered mitochondriaA (cluA) gene, whose disruption in D. discoideum causes the aggregation of mitochondria near the cell center [24]. The CLU1 gene, a Saccharomyces cerevisiae homolog of the FMT and cluA genes, is also involved in mitochondrial distribution [25], indicating that the function of FMTlike genes is conserved in higher plants and fungi. HowCurrent Opinion in Plant Biology 2004, 7:626–631
ever, as the mitochondria of D. discoideum cluA disruptants are interconnected by membraneous strands [26] but those of A. thaliana fmt mutants are not [23], the cluA and FMT genes may function differently in mitochondrial positioning. The FMT protein has tetratricopeptide repeat domain, which is known as a protein–protein interaction domain. At present, the in-vivo function of the FMT gene during mitochondrial positioning remained to be determined; the sub-cellular localization and interacting partner of the FMT protein are unknown. In D. discoideum, drug treatment that disrupts AFs or MTs cannot mimic the cluA disruptant phenotype, suggesting that the cluA gene may function independently of the cytoskeleton [26]. Treatment of tobacco cultured cells with the anti-actin drug LAT-B, the myosin ATPase inhibitor 2,3-butanedione and the sulfhydryl-modifying agent N-ethylmaleimide (NEM) (but not with the MT-depolymerizing drug oryzaline) induced clustered mitochondria and inhibited mitochondrial movement [6], causing a morphology that partially resembled the fmt mutant phenotype. Further analyses of fmt and other mutants will shed light on mechanism of the involvement of AFs and FMT protein in mitochondrial positioning.
Acto-myosin-dependent peroxisomal movement Peroxisomes have multiple important functions in plants, such as fatty-acid b-oxidation, photorespiration, and peroxisomal biogenesis, and these functions have been studied extensively using A. thaliana [27]. Moreover, the www.sciencedirect.com
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peroxisome-targeting signals (PTS), the carboxy-terminal Ser-Lys-Leu tripeptide (PTS-1) and the amino-terminal (R/K)(L/V/I)(X)5(H/Q)(L/A) nonapeptide (PTS-2), are well-known among eukaryotes. Using PTS-fused GFP, four groups independently discovered that peroxisomes move actively in various cell types of A. thaliana [8–10] and in the epidermal cells of onion [10] or leek [7]. The velocity and motility of peroxisomes differ according to the plant cell-types examined: movement can involve slow or rapid directional movement, random oscillations or stop-and-go movements [7–10]. AF-depolymerizing drugs (e.g. LAT-B etc.) inhibit peroxisome movement but MT-depolymerizing drugs do not, indicating that peroxisome movement is dependent on AF but not on MT [7–10]. Co-localization studies have revealed that peroxisomes are closely associated with AFs but not with MTs [7,8,10]. As the myosin inhibitors 2,3-butanedione and NEM also inhibit peroxisome movement, the actomyosin system may be involved in peroxisome movement. Leaf peroxisomes often localize near chloroplasts, probably to maximize photorespiration [9]. It is worth noting that peroxisomes are located at the bottom of the cells of chup1 mutants, alongside the aggregated chloroplasts (Figure 1; [16]). This evidence suggests that the peroxisome–chloroplast association is strong, and that the distribution of peroxisomes is partially dependent on chloroplast movement, at least in leaf cells. Although many mutants that have defective peroxisome functions have been isolated, mutants that are deficient in peroxisome movement and positioning have not yet been isolated [27]. Recently, mutants that are defective in peroxisome morphology were screened by visual inspection of mutagenized GFP–PTS1-transgenic seedlings [28]. A screen using the same mutagenized GFP– PTS1 seedlings has the potential to isolate mutants that are defective in peroxisome movement and positioning.
Golgi movement associated with ER networks Bidirectional coordinate protein transport between Golgi and endoplasmic reticulum (ER) is essential for cell functions, suggesting the importance of the relative positions of the Golgi and ER. When GFP constructs targeted to both Golgi and ER were expressed in leaves of Nicotiana clevelandii, Golgi stacks were shown to move actively in close association with the ER tubules [11]. The application of the anti-actin drug cytochalasin D and the myosin inhibitor NEM inhibited Golgi movement. Importantly, Golgi bodies colocalize with the AF network near ER, suggesting that Golgi movement along ER depends on the acto-myosin system [11]. Nebenfu¨ hr et al. [12] further investigated Golgi motility in tobacco cultured cells and found that Golgi stacks repeated rapid vectorial movements and had a wiggling state, i.e. ‘stopand-go’ movement, which was also shown to be dependent on acto-myosin [12]. When Golgi stacks moved along the same track, they often stopped at the same www.sciencedirect.com
point. Therefore, Nebenfu¨ hr et al. [12] hypothesized that specific sites, such as active ER export sites or areas of cell-wall growth, generate a ‘stop signal’. Interestingly, the expression of a dominant-negative mutation of the Arabidopsis Rab GTPase AtRab1b, AtRab1b (N121I), disrupted the normal movement of Golgi stacks on the ER networks in tobacco [29]. Although the expression of wildtype AtRab1b did not affect Golgi movement, coexpression of wildtype AtRab1b with AtRab1b (N121I) rescued the dominant-negative effect of AtRab1b (N121I), suggesting that AtRab1b activity is necessary for plant Golgi movement. Members of this subclass of Rab GTPases have been implicated in vesicular trafficking in the secretary pathway in yeast, mammals and plants [30,31]. This novel function of AtRab1b suggests an exciting relationship between Golgi movement and vesicular trafficking. A recent paper demonstrated that aggregation of ER networks and Golgi accumulation occurred at the infection site of oomycete pathogen in A. thaliana [32]. At oomycete penetration sites, AFs but not MTs dynamically re-organized and formed large bundles. Takemoto et al. [32] suggested that the accumulation of ER at the penetration site serves as the ‘stop signal’ for Golgi movement, resulting in Golgi accumulation at the penetration site. ER–Golgi accumulation at the pathogen infection site may serve as secretion site for pathogen resistance.
Conclusions Organelle movements in seed plants are mediated mostly by AFs rather than by MTs, although most of the work carried out to date has involved inhibitor studies that have not provided detailed information about mechanisms. Precise studies on CHUP1 proteins that are localized on the chloroplast outer envelope may provide a strong clue to shed light on the mechanism of organelle movements, at least on chloroplast movement (Figure 2; [16]). The mechanism of movements may not be so difficult to determine because the involvement of AFs is clear. The most important questions are how organelles recognize where to go and stay, and how organelles are fixed at their appropriate positions. During chloroplast positioning within the cells of fern gametophytes [33,34] and Arabidopsis mesophyll cells [17], ring-like structures made of AFs appear and seem to connect chloroplasts to the cell surface when they have reached their position. We do not know, however, whether the ring-like AFs bind chloroplasts to the cell surface to anchor them. Recently, western blot and immunofluorescence studies using the maize myosin XI antibody have revealed that maize myosin XI co-localizes with mitochondria and plastids but not with nuclei, Golgi, ER, and peroxisomes [35]. Therefore, chloroplast or mitochondrial movement may be mediated by the acto-myosin system (Figure 2). Although no information is available yet about the posCurrent Opinion in Plant Biology 2004, 7:626–631
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sible association of intermediate filaments or cytoskeletal proteins with the cytoplasmic face in plants, the recent identification of a bacterial intermediate-filament-like protein opened the possibility that plants also have intermediate filaments [36,37]. The fine structure underneath the plasma membrane and the genes that are involved in constructing this structure must be studied if we are to understand organelle movement and positioning.
Acknowledgements This work was partly supported by Grand-in-Aid for Scientific Research (on Priority Areas, no.s 13139203 and 13304061) from the Ministry of Education, Sports, Science and Technology (MEXT) of Japan, and Grant-in-Aid for Scientific Research (no. 16107002) from the Japan Society for the Promotion of Science to MW, and by a grant from the Research Fellowships of the Japan Society for the Promotion of Science for Young Scientists to NS.
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