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Keywords:

  • Arabidopsis;
  • ARC6;
  • cyanobacteria;
  • FtsZ;
  • Min system;
  • organelle;
  • PDV1;
  • PDV2;
  • plastid division

Abstract

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

Chloroplasts are descendants of cyanobacteria and divide by binary fission. Several components of the division apparatus have been identified in the past several years and we are beginning to appreciate the plastid division process at a mechanistic level. In this review, we attempt to summarize the most recent developments in the field and assemble these observations into a working model of plastid division in plants.


Plastids: Characteristics, origins and distribution

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

Without chloroplasts, plant and animal life would not exist. Plastids have been studied most extensively with regard to their role in photosynthesis, and are essential to the plant itself, being required for a select of amino acid and lipid metabolism, assimilation of nitrogen and sulfur into organic compounds, signaling in response to environmental cues and biosynthesis of plant hormones (1–5). Despite all we have learned over the past century regarding this organelle, we still know relatively little about its replication—which, superficially, seems a simple process of binary fission. Because plastid division and biogenesis are an integral part of plant growth and development, understanding of the division process is paramount to our understanding of plant and organelle biology.

Plastids arose from an endosymbiotic relationship between a primitive biciliate protozoan and an ancient cyanobacterium that began 1.2–1.5 billion years ago (Figure 1) (6,7). Initially, each partner derived some mutual benefit: the cyanobacterium occupied and exploited an untapped protective niche and the host extracted reduced carbon and other nutrients from its new tenant. Time, selective pressure and mutation bring us to the present day where the chloroplast is a prisoner in its own home, having lost more than 90% of the gene content present in its free-living ancestor. Most of the plastid division genes identified to date are cyanobacterial in origin and encode division proteins localized inside the organelle. These are directed to the plastid post-translationally by N-terminal transit peptides that are cleaved upon import. A few components of the division complex were invented by the host after endosymbiosis and function on the surface of the organelle in contact with the cytosol; the mechanisms targeting these proteins to the chloroplast surface are unknown. In land plants, all known plastid division genes reside in the nucleus—none are found in the plastid genome—although in at least one unicellular green alga two plastid division genes remain associated with the plastid genome (8).

image

Figure 1. Origin of plastids. All plastids are derived from a single endosymbiotic relationship in which a biciliate protozoan engulfed a cyanobacterium. After some time, this primary plastid-bearing lineage diverged into glaucophyte algae, red algae and green algae/plants. Secondary and tertiary plastids are derived by engulfment of one of the primary lineages once or multiple times by another eukaryote (not shown). Adapted from Cavalier-Smith (6).

The terms chloroplast and plastid are often used interchangeably, but in vascular plants chloroplasts are actually a subset of plastids, each of which is specialized for a given set of functions within a specific cell type. All plastids are surrounded by two envelope membranes and are derived by division from a population of proplastids within the meristematic (stem) cells of the plant. Beyond land plants, red algae, green algae and diatoms also contain plastids. Even apicomplexan human pathogens like Toxoplasma gondii and Plasmodium falciparum contain plastids that are distant cousins of the chloroplast, making fundamental plastid biology relevant to our understanding of human diseases (9,10). Diatoms and apicomplexans obtained their plastids through ‘secondary’ endosymbiosis—engulfment of a plastid-containing eukaryote by another eukaryotic cell (10). This is in contrast to the primary plastid-bearing lineage, which includes glaucophyte algae, red algae, green algae and plants (6).

Organisms used to study plastid division

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

Most of the known components of the plastid division apparatus have been identified in Arabidopsis using both forward and reverse-genetic approaches, but the study of plastid division has benefited from the use of several other photosynthetic organisms, including Cyanidioschyzon merolae (a unicellular red alga), Physcomitrella patens (a multicellular moss) and several cyanobacterial genera. As we describe our current understanding of the division process in this review, we will attempt to highlight the advantages and disadvantages of each system.

FtsZ and control of Z-ring assembly and placement

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

FtsZ

Because of the prokaryotic heritage shared by all plastids, bacterial cell division systems (11,12) have been used to guide investigation of the plastid division apparatus. The first bona fide component of the plastid division apparatus was identified through a reverse-genetic approach when an FtsZ-like sequence with similarity to Escherichia coli FtsZ was found in an Arabidopsis expressed sequence tag (EST) collection (13). Escherichia coli FtsZ is a tubulin-like, polymer-forming GTPase (14) that is essential for cell division and forms a ring-shaped structure (the ‘Z-ring’) at the division site (15). FtsZ is thought to be one of the first protein components to arrive at the division site, where it acts as a scaffold for other division proteins (16). It may also provide a contractile force that facilitates cytokinesis (12). FtsZ is conserved among almost all prokaryotes, including the cyanobacteria, and all sequenced cyanobacterial genomes have only one FtsZ gene (17). In photosynthetic eukaryotes, plastidic forms of FtsZ seem to be encoded by at least two gene families (18), suggesting that FtsZ gene duplication may have been critical for evolution from endosymbiont to organelle. In a small number of unicellular eukaryotes, FtsZ proteins derived from the α-proteobacterial ancestor of mitochondria also function in mitochondrial division, but mitochondrial FtsZ has been lost from most eukaryotes, including plants (19).

In plants, plastid division is mediated by two phylogenetically distinct FtsZ families called FtsZ1 and FtsZ2 (18,20). Both proteins reside in the stromal compartment of the chloroplast and localize to rings at the plastid division site (21–23). The two most noted differences between the FtsZ1 and FtsZ2 proteins are (a) the variability of a single amino acid residue in the otherwise conserved ‘tubulin signature motif’ (24) and (b) the conservation in FtsZ2 proteins, but not FtsZ1, of a short C-terminal motif found in most bacterial FtsZ proteins. In E. coli, this motif mediates an interaction between FtsZ and two membrane-associated Z-ring stabilizing factors, FtsA and ZipA (25–27). The significance of the difference between FtsZ1 and FtsZ2 in the tubulin signature motif is not yet known, but the C-terminal motif in FtsZ2 has been shown to mediate an FtsZ2-specific interaction with the chloroplast division protein ARC6 (28) (described below). Neither ZipA nor FtsA are found in plants, but the observation that ARC6 stabilizes plastidic FtsZ filaments in vivo (29) suggests it could be functionally related to one of these proteins. One possibility is that ARC6 could promote bundling of FtsZ protofilaments similar to EcZipA (27), thereby promoting Z-ring assembly.

In addition to the differences in amino acid sequence, there is some evidence that FtsZ1 and FtsZ2 differ in their biochemical properties and in vivo behavior. El-Kafafi et al. (30) have shown that purified recombinant FtsZ1 is capable of forming high-molecular weight complexes in the presence of GTP, and electron microscopy shows that these complexes resemble filaments formed by recombinant E. coli FtsZ (31,32). FtsZ2 did not form any filament-like structures in vitro in these experiments, but seemed instead to form aggregates in the presence of GTP. While these are intriguing differences, all the above experiments were performed with precursor proteins bearing the transit peptides; thus their relevance to the in vivo behavior of mature FtsZ1 and FtsZ2 are unclear. El-Kafafi et al (30) also found that FtsZ1 and FtsZ2 have differing membrane association characteristics upon fractionation of isolated spinach chloroplasts (30); FtsZ1 was mostly present as a soluble stromal protein, while most of the FtsZ2 was tightly associated with the envelope membranes. We have obtained similar results in some experiments (J. M. Glynn, B. Olson and R. S. McAndrew, unpublished observations) but not others (22). The reasons for the variation in FtsZ2 membrane association are unclear. One possibility is that FtsZ2 localization varies at different stages of the plastid division cycle, although this has not been investigated. Typical chloroplast preparations from Arabidopsis or other higher plants are likely heterogenous; they do not contain a uniform population of dividing plastids, frozen at the same stage of division. It is likely that the Z-ring and the other components of the division apparatus are undergoing dynamic assembly and disassembly throughout the organelle division cycle. Therefore, synchronizable algal experimental systems may yield important insights into FtsZ properties and dynamics, possibly linking FtsZ behavior with a discrete stage of plastid division.

Experiments showing that either overexpression or depletion of FtsZ1 or FtsZ2 causes dose-dependent defects in chloroplast division, as indicated by reduced numbers of oversized chloroplasts (20,33), suggest that their stoichiometry may be important for Z-ring function (33). Consistent with this possibility, quantitative analysis of FtsZ1 and FtsZ2 protein levels in isolated Arabidopsis chloroplasts reveals an approximate 1:2 molar ratio (34). Whether this reflects the stoichiometry in the Z-ring of dividing chloroplasts is not yet known and further work is needed to clarify this matter as well.

Control of Z-Ring assembly and placement: The Min system

In bacteria, Z-ring positioning is regulated by the Min system (named for the mutant minicelling phenotype). The Min system allows Z-ring assembly at the mid-cell and prevents Z-ring assembly near the cell poles, ensuring symmetric division (12). An evolutionarily divergent Min system appears to actively position the placement of the Z-ring in the chloroplast.

The Min system has been studied extensively in the Gram-negative bacterium E. coli, in which it is composed of three protein components: MinC, MinD and MinE (12). MinC binds directly to and inhibits polymerization of FtsZ (35–37), but MinC activity is restricted to the cell poles by the concerted action of MinD and MinE (38,39). MinD, a polymer-forming ATPase, assembles on the inner leaflet of the cell membrane near the cell poles, forming a polar zone (12,40). By binding to MinD, MinC becomes tethered to the membrane in the polar zones, inhibiting FtsZ polymerization at the cell poles (41,42). MinD, and hence MinC, is prevented from localizing to the division site by MinE, which forms a dynamic ring-like structure near the mid-cell at the edge of the polar zone (43,44). By stimulating MinD ATPase activity, MinE causes MinD, and hence MinC, to dissociate from the membrane, keeping the concentration of membrane-associated MinD/MinC low near the cell center. The absence of MinC at the cell center allows FtsZ polymerization and Z-ring assembly to occur at this position, initiating division. In E. coli, the positioning of the Min system itself is directed in part by an oscillatory mechanism wherein all three Min proteins move from pole-to-pole (45). Oscillation of the Min system is critical both for inhibiting Z-ring assembly at the cell poles and permitting assembly at the cell center in E. coli (12). However, MinE is not found in the Gram-positive bacterium Bacillus subtilis. Instead, MinCD activity is restricted to the cell poles, and hence Z-ring assembly to the cell center, by the action of DivIVA in a distinct mechanism that does not involve pole-to-pole oscillation of MinC and MinD (11,12).

Homologs of MinD and MinE have been identified in plants and green algae and the Arabidopsis proteins AtMinD and AtMinE have been functionally characterized by Møller and colleagues (8,46–48). Mature AtMinD and AtMinE are stromal proteins, and green fluorescent protein (GFP) fusion proteins localize near the poles of chloroplasts (46,47,49). AtMinD and AtMinE are capable of interacting in vivo (28) and their biochemical characteristics are mostly consistent with those of their bacterial cousins, although AtMinD ATPase activity is calcium rather than magnesium dependent (50,51). Unlike EcMinD, AtMinD ATPase activity is not stimulated by phospholipid (50)—perhaps unsurprising because the plastid inner envelope membrane contains very little phospholipid (52). The same group found that AtMinE can form homodimers in yeast and in planta and propose that AtMinE homodimers near the mid-plastid might quench the activity of an AtMinD-containing FtsZ-inhibitory complex, much like the Min system of E. coli (12). How the reported polar localization of AtMinE (49) relates to this hypothesis is unclear, however. Whether AtMinD and AtMinE form oscillating complexes in chloroplasts is unknown.

Mutant analysis in Arabidopsis has shown that both AtMinD and AtMinE play roles in positioning of the chloroplast division site (47,48). Consistently, assembly and placement of the plastidic Z-ring is altered in the corresponding mutants (D. W. Yoder, S. Vitha, K. W. Osteryoung, unpublished observations) further suggesting that the Arabidopsis proteins have roles analogous to MinD and MinE in E. coli. Depletion of AtMinD by expression of an antisense transgene results in multiple asymmetric constrictions within a single chloroplast, reminiscent of E. coli minD mutants (48); the position of each Z-ring coincides with a constriction (29). A mutant allele of AtMinD, arc11, confers similar Z-ring phenotypes (Figure 2). In contrast, when AtMinD is overexpressed, FtsZ is not observed in rings, but is instead detected in short filaments inside a greatly enlarged plastid (29). These results are consistent with a role for plastidic MinD in preventing Z-ring assembly away from the mid-plastid. AtMinE has the opposite effect on plastidic Z-ring formation. Recently, arc12 was identified as an allele of AtMinE (S. Miyagishima, D. W. Yoder, K. W. Osteryoung, unpublished observations). A frameshift mutation in arc12 introduces a premature stop codon into the AtMinE open reading frame (Figure 2, legend), and the wild-type AtMinE gene complements the arc12 chloroplast division defect. arc12 mutants have short FtsZ filaments within a single oversized plastid, like AtMinD overexpressors (29), whereas AtMinE overexpressors have multiple sites of constriction, and presumably multiple Z-rings, within the chloroplasts (49,53), similar to AtMinD depletion and arc11 mutants (48). Taken together, the morphological and FtsZ localization phenotypes in AtMinD and AtMinE overexpressors and mutants suggest that plastidic MinD and MinE antagonistically regulate Z-ring assembly and placement in plant cell chloroplasts.

image

Figure 2. Phenotypes associated with arc3 and AtMinD/E alleles. Top panel. A–D) Immunofluorescent micrographs of AtFtsZ2-1 filament morphology in chloroplasts of mature leaf mesophyll cells from wild-type Col-0 (A), arc11 (an allele of AtMinD) (B), arc12 (an allele of AtMinE) (C) and arc3 (D). Bottom panel. E–H) Light micrographs of chloroplast morphology in mature leaf mesophyll cells from wild-type Col-0 (E), arc11 (an allele of AtMinD) (F), arc12 (an allele of AtMinE) (G) and arc3 (H). Arrowheads in (H) show points of constriction in arc3 that are probable sites of Z-ring formation. The mutant arc12 harbors a lesion (1196GC to A–) in AtMinE creating a frameshift and premature stop, changing 108AWKI111 to 108IGRStop111. Size bars = 20 μm.

Although MinD and MinE are conserved in plants, no MinC-like sequences have been found in plant genomes or EST collections. This is somewhat surprising because all three Min components are conserved among the cyanobacteria (17). In E. coli, Z-ring assembly is inhibited at the cell poles by direct interaction of FtsZ with MinC, which is thought to destabilize FtsZ polymers (54). Neither AtMinD nor AtMinE binds FtsZ1 or FtsZ2 (28), suggesting they do not have a MinC-like function—therefore, it seems plausible that a functional analog of MinC may exist in plants that are divergent from cyanobacterial MinC. The chloroplast division protein ARC3, a plant-specific protein with some similarity to FtsZ (55), has been suggested as a possible chloroplastic MinC replacement based on its mutant phenotype and ability to interact with AtMinD, AtMinE and FtsZ1 (56). Further, we have observed that arc3 mutants exhibit multiple Z-rings within a single enlarged chloroplast (Figure 2D), similar to arc11 (Figure 2B) and AtMinD antisense mutants (29). Taken together, these findings suggest that ARC3 may indeed regulate Z-ring positioning and assembly in a MinC-like manner. Although ARC3 has a weakly predicted N-terminal transit peptide and is reported to be cytosolic (55), the above findings also suggest that ARC3 is a stromal protein. In agreement with this likelihood, we find that a rice ortholog of ARC3 is strongly predicted by TargetP (57) to be chloroplast-targeted. While the reported localization of ARC3 at the division site (55) may seem inconsistent with its proposed MinC-like function and colocalization with AtMinD near the poles of the chloroplast (49,56), mid-cell and polar localization of GFP-MinC has been reported for B. subtilis (58). Moreover, cyanobacteria encode relatives of both MinE (found in Gram-negative bacteria) and DivIVA (found in Gram-positive bacteria) (17), suggesting that cyanobacteria and plastids employ a mechanism for Z-ring placement that is distinct from those used by E. coli or B. subtilis. Further work on ARC3 is needed to verify its localization, topology and function in chloroplast division.

Beyond the Min system, a second mechanism of Z-ring placement, called nucleoid occlusion, is also active in many bacteria (12). The nucleoid occlusion system inhibits assembly of the Z-ring over the bacterial chromosome, averting scission of the genome during cytokinesis (59,60). However, recent evidence suggests that a nucleoid occlusion mechanism does not operate in the cyanobacterium Synechococcus elongatus, as Z-rings were noted to form around the chromosome in this organism (17). There is no strong evidence to support the existence of an analogous system in other cyanobacterial species or plastids based on blast searches using E. coli SlmA (59) as a query sequence (J. M. Glynn, Michigan State University, East Lansing, Michigan, unpublished observations).

Promoting Z-ring assembly: ARC6

Previous work from our laboratory has shown that ARC6 is a bitopic inner envelope membrane protein that acts as a positive regulator of Z-ring formation (29). ARC6-GFP localizes to a ring-like structure at the mid-plastid (29), similar to AtFtsZ1 and AtFtsZ2 proteins (23). ARC6 is a descendant of the cyanobacterial cell division gene Ftn2 (61), and ARC6 and its orthologs are only found in cyanobacteria and chlorophytes. ARC6 and Ftn2 proteins possess a conserved region at their N-termini with sequence similarity to J-domains, implicating them as possible Hsp70-associated co-chaperones. arc6 mutants have short FtsZ filaments within a single large chloroplast and ARC6 overexpressors have abnormally long, branched FtsZ filaments held within an oversized plastid. These phenotypes suggest that ARC6 could play a role in bundling of short FtsZ filaments into a ring at the chloroplast division site.

The N-terminus of ARC6 resides in the stroma (29) and a conserved N-terminal segment of ARC6 interacts with FtsZ2 but not FtsZ1 (28). This interaction represents an important functional difference between the FtsZ1 and FtsZ2 families. The interaction requires the short C-terminal motif in FtsZ2 described above, reminiscent of the FtsZ–ZipA and FtsZ–FtsA interactions observed in E. coli (62). The J-domain of ARC6 is not required for interaction with FtsZ2 (28). In contrast, Ftn2 is reported to require the J-domain for interaction with cyanobacterial FtsZ (63) but the significance of this difference is not yet understood.

Despite its relevance to our understanding of plastid division, only a few studies have identified components of the cyanobacterial cell division apparatus (17,61). While some of these are orthologs of division components from Firmicutes and Proteobacteria, others like ARC6 are unique to the cyanobacterial lineage and provide an opportunity for greater understanding of the chloroplast division apparatus. Discrete analysis of division components may be more convenient and efficient in cyanobacteria rather than Arabidopsis or other model systems because of the short generation time, ease of transformation and gene replacement, and the ability to obtain a near-synchronous culture.

GC1/AtSulA: An indirect regulator of the Z-ring

GIANT CHLOROPLAST 1 (also known as GC1 and AtSulA) was discovered on the basis of its sequence similarity to SulA-like proteins from cyanobacteria (64,65). GC1 was shown to be associated with the inner envelope and is likely to be a key regulator of the division process, although its exact function is still unknown. In a subset of bacterial systems, induction of SulA is one of many responses to DNA damage; SulA inhibits cell division by binding directly to FtsZ and occluding the protofilament interface, preventing FtsZ polymerization (66,67). However, unlike SulA, GC1 does not appear to possess an FtsZ-binding domain identical to that in Pseudomonas aeruginosa SulA (66) nor does it bind FtsZ1 or FtsZ2 directly (64). Although SulA inhibits cell division in bacteria, the published effects of GC1 on chloroplast division are contradictory: work from one group suggests that GC1 acts as a positive regulator of chloroplast division (64), while work from another indicates that it acts as a negative regulator (65). Further work on GC1 is needed to clarify its role in the division process.

Plastid-dividing rings and dynamin

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

The plastid-dividing rings

A great deal of what we know about plastid division has come from pioneering microscopic analysis of the division process in both plants and algae. Early observations of plant cells provided the first evidence that chloroplasts divide by binary fission and that this type of mechanism accounts for the increase in chloroplast number observed as leaf cells mature (68,69). In the mid-1980’s, two electron-dense plastid-dividing (PD) rings were shown to exist at the central constriction of dividing chloroplasts in oat leaves, building on previous observations in red algae (70,71). The outer PD ring is located on the cytosolic side of the outer envelope membrane and consists of fine filaments 5–7 nm in diameter (72,73), while the inner PD ring resides on the stromal side of the inner envelope membrane. Similar PD rings have also been observed in other species and it is widely believed they are a common component of the division apparatus, although C. merolae appears to have an additional PD ring (three PD rings in total) located between the inner and outer envelopes (74). The PD rings are discrete structures, distinct from the dynamin ring (discussed below) on the outside of the outer envelope and distinct from the FtsZ ring on the inner side of the inner envelope (75,76). The constituents of the PD rings have not been identified and would be a very valuable find for the field in general.

The role of dynamins in plastid division

A role for dynamin-related proteins in chloroplast division was realized upon isolation of the Arabidopsis ARC5 locus and the association of a dynamin-like protein with dividing plastids of C. merolae (77,78). The dynamins are a superfamily of GTPases that are involved in an array of membrane-altering events (79), including mitochondrial fission (80) and fusion (81). Dynamins can self-assemble into spiral-shaped polymers that facilitate membrane constriction by virtue of their mechanochemical activity (82–85). Chloroplasts in arc5 mutants are dumbbell shaped, indicating a late-stage arrest in the division process. In C. merolae, dynamin patches migrate from the cytosol to the middle of a dividing plastid, later forming a ring-like structure around the outside of the chloroplast that decreases in diameter as the plastid constricts (78). These dynamin-like proteins are directed to the outer surface of the outer envelope and therefore do not require an N-terminal transit peptide, but are recruited to the plastid by a unique mechanism (see below).

Work from the Kuroiwa laboratory has recently shown that a plastid-dividing dynamin FtsZ (PDF) ring can be isolated as an intact structure from synchronized C. merolae cells and that the linkage between all the rings is continuous and likely free of membrane lipids (76). Using optical tweezers, the Kuroiwa group showed that the major constrictive force for division likely comes from the dynamin portion of the PDF structure as only a dynamin-containing PDF structure is capable of snapping back to its original position after being stretched. They suggest a dual role for dynamin in the division process: (a) dynamin facilitates sliding of the 5–7 nm filaments during the early stages of division, acting at the outer surface of the outer PD ring; (b) following the primary constriction event, the dynamin filaments somehow move through the outer PD ring, making direct contact with the membrane itself and completing fission by acting as a ‘pinchase.’ Because of the ability to isolate a population of plastids at the same stage of division from C. merolae, which is not possible for land plants, this organism will continue to serve as an outstanding model for studies of plastid division.

Recruitment of dynamin to the division site by PDV1 and PDV2

A screen for Arabidopsis mutants with an arc5-like division arrest led to the identification of PDV1. Like ARC5, PDV1 is localized to a ring-shaped structure at the mid-plastid (86) (Figure 3). PDV2 was subsequently identified on the basis of sequence similarity to PDV1, and pdv2 mutants also exhibit arc5-like chloroplast division defects. PDV1 and PDV2 appear to have overlapping functions in recruitment of ARC5 to the division site as GFP-ARC5 is not recruited to the oversized chloroplasts present in the pdv1 pdv2 double mutant (86). However, it is not yet known if PDV1 and PDV2 interact with each other if either interacts with ARC5. PDV1 is associated with the outer envelope membrane. Both proteins have a single predicted transmembrane helix and the amino terminus of PDV1 faces the cytosol, but it is not known if either protein protrudes into the intermembrane space (IMS). If so, these proteins, along with ARC6, could mediate coordination between the inner and outer components of the division complex. Consistent with this possibility, we have identified a unique yeast two-hybrid interaction between PDV2 and ARC6 (J. M. Glynn, unpublished observations) that is now being verified and characterized in Arabidopsis.

image

Figure 3. Phenotype of pdv1-2 and localization of pPDV1-GFP-PDV1. Chloroplast morphology and number are significantly altered in the pdv1-2 (right) background, relative to wild type (left). GFP-PDV1 is localized to the division site of wild-type dividing chloroplasts (inset, left panel). The scale bar in the right panel is 10 microns.

Neither PDV1 nor PDV2 have orthologs in the red algae C. merolae or Galdieria sulphuraria, but orthologs of both PDV1 and PDV2 have been identified in Poplar, moss and rice, suggesting that PDV1/PDV2 were invented after the red/green divergence and that red algae and green plants may use different schema for recruitment of dynamin to the division site. Regardless, the identification of PDV1 and PDV2 reiterates the utility of forward mutant screens for discovery of novel plastid division genes in Arabidopsis.

Perspectives

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

Control of plastid volume

An intriguing observation from analysis of Arabidopsis chloroplast division mutants is that chloroplast plan area (and likely volume) remains fairly constant between wild-type and ‘big-plastid’ mutants (20,87,88), implying a mechanism for chloroplast ‘volume sensing.’ Plant homologs of the bacterial mechanosensitive ion channel protein MscS could be components of such a volume-sensing system. In bacteria, Msc proteins are involved in regulating osmotic potential across the membrane in response to increased membrane tension induced by osmotic shock (89). The Arabidopsis MscS-like (MSL) proteins MSL2 and MSL3 are localized to chloroplast membranes and msl2-1 msl3-1 double mutants have chloroplast morphology defects similar to those observed in chloroplast division mutants (90). MSL2 and MSL3 also colocalize with MinE in transient overexpression assays. MSL3 is capable of rescuing the osmotic shock sensitivity of a bacterial mechanosensitive ion channel mutant, suggesting that MSL proteins have a related function in chloroplasts. The authors propose MSL ion transporters respond to membrane tension by altering ion flux between the chloroplast and the cytosol. The phenotype observed in the msl2-1 msl3-1 double mutants might reflect a compensatory increase in chloroplast volume to alleviate elevated intra-chloroplast ion concentration, thereby reducing osmotic pressure on the organelle. Pyke has suggested a role for the MSL system as a mechanism for density-dependent control of chloroplast number and volume in leaf mesophyll cells (91). The potential connection between ion transport and chloroplast size is intriguing; further work on the plastidic MSL system may shed light on how plant cells regulate chloroplast volume.

Effects of photocollection and photoprotection on chloroplasts

The viability of chloroplast division mutants with large chloroplasts raises the question of why plant cells evolved to have multiple chloroplasts. The answer may have to do partly with the ability of multiple small chloroplasts to redistribute more effectively than fewer large chloroplasts inside the cell. Unlike motile unicellular algae with single chloroplasts, land plants are sessile and cannot relocate in response to sudden changes in light intensity. Chloroplasts in land plants avoid potentially damaging high light by moving to the cell periphery and orienting in columns parallel to the plane of incoming light; under low-light conditions, chloroplasts gather in periclinal layers to maximize light absorption (92,93). This movement is believed to be largely performed through actin reorganization (93–95). Consistent with the importance of plastid division for plastid movement, a mutant allele of Arabidopsis FtsZ1 shows diminished plastid movement in response to high light (S. DeBlasio, R. Hangarter, D. Yoder, K. W. Osteryoung, unpublished observations). Interestingly, the thylakoid morphology of big-plastid mutants is reminiscent of that in high light-adapted plants (96), a likely pleiotropic effect resulting from impaired plastid movement arising from defects in chloroplast division. Conversely, mutations in proteins influencing thylakoid organization also impair chloroplast division (97–99). The regulation of thylakoid morphology and chloroplast movement may be integrated into a system that maximizes photosynthetic output and minimizes light-induced damage in response to changing environmental stimuli (100,101).

Coordination of cell division and plastid number in land plants

Being essential organelles, plastids are presumably required by every plant cell at some point during development. For algal species bearing a single plastid, like C. merolae, plastid division must be coordinated with cellular division, but in Arabidopsis and other plants this coordination is less stringent, probably because multiple proplastids are present in every meristematic cell. It seems likely that plastid and cell division are coordinated to some degree, although the mechanisms involved are unknown. One possibility is that origin-licensing factors or other proteins influenced by the cell cycle might have dual roles in coordinating events in the nucleus with those in organelles. Cdc10-dependent transcript 1 (CDT1) has been identified as a member of the pre-replication complex in several eukaryotes, where it appears to act as a key regulator of nuclear DNA replication (102,103). Recent work in Arabidopsis suggests that a CDT1 paralog, CDT1a, may play a role in this coordination (104). Plants expressing an AtCDT1-RNAi construct show defects in plastid size and number, reminiscent of those in bona fide plastid division mutants. CDT1a was shown to be dually targeted to the nucleus and chloroplast. The authors suggest that AtCDT1a is a component of the nuclear pre-replication complex as in other eukaryotes, and that the plastidic form of the protein may regulate plastid Z-ring assembly through interaction with ARC6, based on two-hybrid experiments. Further work on CDT1 is required to determine the extent to which CDT1 is involved in coordinating cell and plastid division.

Developmental controls on plastid fission

Analysis in other vascular plants is also providing clues about the relationship between plastid division and plastid conversion. Examination of tomato has revealed suffulta, a recessive mutation leading to abnormalities in chloroplast number and content reminiscent of the arc6 mutation (29,105). However, as the fruit begins to ripen, the abnormal chloroplasts are converted to relatively normal chromoplasts. This finding suggests that a unique plastid fission pathway exists during fruit ripening that is developmentally coordinated with the conversion of chloroplasts to chromoplasts. The authors also show that a novel fission pathway may be active in guard cell development, as aplastidic stomatal guard cells can harbor a body that accumulates plastid-targeted GFP. They suggest this pathway may be related to those giving rise to the dynamic tubular extensions of the plastid membrane called stromules (106). Whether these alternative fission pathways involve canonical plastid division proteins such as FtsZ1 and FtsZ2 is not yet known.

Bacterial cell wall synthesis in plastids

Analysis of the moss P. patens revealed that homologs of bacterial Mur genes, which are involved in production of the periplasmic peptidoglycan-containing cell wall, may also affect plastid number and morphology. This analysis was preceded by the observation that moss chloroplast size and number can be influenced by treatment with compounds known to inhibit to bacterial cell wall biosynthesis (107,108). Mining of the Physcomitrella genome revealed nine homologs of bacterial cell wall synthesis genes, and interruption of some of these genes led to defects in chloroplast morphology and number reminiscent of phenotypes seen following treatment with β-lactams (109). This is rather surprising because land plants have no peptidoglycan layer within the boundaries of the chloroplast, although other plastid-bearing lineages like the glaucophyte algae have retained a peptidoglygan-containing cell wall (110). Curiously, Arabidopsis and rice contain a four-member subset of these putative cell wall synthesis genes (109), but Arabidopsis (J. M. Glynn, unpublished observations) and tomato are unresponsive to β-lactam antibiotics (108). Collectively, these observations suggest that: (a) methods for cell wall detection lack sensitivity or are incapable of recognizing the unique cell wall layer possessed by higher plant chloroplasts or (b) land plants have no plastidic cell wall, but they have retained a subset of the cell wall biosynthesis genes for some other function. Either prospect is interesting and will have significant impacts on how we envision the operation of the plastid division machinery.

Working model of chloroplast division

In Figure 4, we present a working model of chloroplast division in plants that synthesizes much of the information described above. The model postulates the following: FtsZ1 and FtsZ2 assemble into short filaments in an unknown configuration. FtsZ filament formation is inhibited at the poles by the concerted action of MinD and possibly ARC3 (not shown in the model). MinE quenches the FtsZ polymer-inhibiting activity near the mid-plastid, allowing FtsZ filament formation. FtsZ filaments are bundled at the division site to form the plastidic FtsZ ring. Bundling activity is promoted by the inner envelope protein ARC6. The Z-ring undergoes constant remodeling during the division cycle and its assembly and dynamics are influenced by ARC6, ARC3, MinD and MinE. Assembly of the Z-ring is followed by formation of the inner PD ring and then the outer PD ring; the compositions and functions of the PD rings are unknown. The C-terminal IMS domain of ARC6 may mediate interaction with factors in the IMS and/or signaling between the stroma and IMS, possibly through PDV2 or other unknown factor. PDV1 and PDV2 together mediate the recruitment of the dynamin, ARC5. The roles of GC1, MSL2 and MSL3 during division are unclear at this time. Late membrane constriction is driven largely by ARC5 (dynamin) with progressive widening of the outer PD ring. During this final squeeze, dynamin filaments move to the inner face of the outer PD ring, making direct contact with the outer membrane surface to complete the fission process.

image

Figure 4. Tentative model of chloroplast division. The tentative model shown here starts with a pre-assembled division complex and does not show the order of assembly. For figure clarity, not all proposed interactions are shown here; for further information, the reader should see Aldridge et al. (53) and Maple and Moller (56). The number and size of individual protein molecules shown is not to scale. See text for full description.

This model is still largely speculative and will clearly undergo considerable revision as additional data are obtained.

Closing remarks

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

Given all the above, the regulation of plastid division is no simple matter. While we have seen examples where developmental programs seem to dictate plastid type, number and shape, the precise connections between development, inter-organelle signaling, light perception, photosynthesis and metabolic flux as they relate to chloroplast division are still unknown. Genetic screens, proteomics and other approaches aimed at identification of the major control mechanisms will likely reveal key regulatory players and may help us connect plastid division to several other fundamental processes in plants.

Acknowledgments

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References

We extend our apologies for omission of many relevant articles because of space constraints. J. M. G. thanks Aaron Schmitz for critical review and advice on the manuscript. This work was supported by grants from the National Science Foundation and the United States Department of Energy to K. W. O.

References

  1. Top of page
  2. Abstract
  3. Plastids: Characteristics, origins and distribution
  4. Organisms used to study plastid division
  5. FtsZ and control of Z-ring assembly and placement
  6. Plastid-dividing rings and dynamin
  7. Perspectives
  8. Closing remarks
  9. Acknowledgments
  10. References