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Keywords:

  • microtubule;
  • CLASP ;
  • endosomal sorting;
  • SNX1;
  • actin filament;
  • kinesin;
  • myosin;
  • Arabidopsis thaliana ;
  • Nicotiana ;
  • characean algae

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

Movement of secretory organelles is a fascinating yet largely mysterious feature of eukaryotic cells. Microtubule-based endomembrane and organelle motility utilizing the motor proteins dynein and kinesin is commonplace in animal cells. In contrast, it has been long accepted that intracellular motility in plant cells is predominantly driven by myosin motors dragging organelles and endomembrane-bounded cargo along actin filament bundles. Consistent with this, defects in the acto-myosin cytoskeleton compromise plant growth and development. Recent findings, however, challenge the actin-centric view of the motility of critical secretory organelles and distribution of associated protein machinery. In this review, we provide an overview of the current knowledge on actin-mediated organelle movement within the secretory pathway of plant cells, and report on recent and exciting findings that support a critical role of microtubules in plant cell development, in fine-tuning the positioning of Golgi stacks, as well as their involvement in cellulose synthesis and auxin polar transport. These emerging aspects of the biology of microtubules highlight adaptations of an ancestral machinery that plants have specifically evolved to support the functioning of the acto-myosin cytoskeleton, and mark new trends in our global appreciation of the complexity of organelle movement within the plant secretory pathway.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

The intracellular transport of organelles in secretory pathways is one of the most critical and yet least understood processes in cells. Organelle transport in animal cells depends on microtubules and associated motor proteins, such as dynein and kinesin, whereas in plants actin and myosin move organelles. At first glance it is puzzling that there should be such a dramatic difference in the use of cytoskeletal tracks and motor systems in plant cells, except when one considers the specialization of plant cells and the peculiarities of their motors. It has been suggested that fast myosin motors enable plant cells, which are comparatively large and highly vacuolated, to achieve more efficient trafficking of metabolites (Gunning, 1999; Wasteneys, 2002). Land plants share common ancestry with the characean algae, the giant internodal cells of which can be up to 15 cm in length and achieve streaming rates approaching 100 microns per second, and from which the acto-myosin basis for cytoplasmic streaming was first established (Williamson, 1974; Kersey et al., 1976; Grolig et al., 1988). Indeed, the myosin motors found in characean algae may be the fastest myosin motors in existence, with tracking speeds up to 10 times the velocity of muscle contraction (Morimatsu et al., 2000). Moreover, it has been calculated that the high ATPase activity of Chara myosins can generate the characteristic cytoplasmic streaming in these cells with as little as 1% motor activity (Yamamoto et al., 2006), leaving little need for support from other motors. The evolution and adoption of such efficient and fast motors may have led to the predominant use of myosin in the green algal ancestors of land plants. It has been confirmed that this form of intracellular motility is conserved in land plants (Reddy, 2001).

Angiosperms and two subclasses of gymnosperms (Pinadae and Gnetidae) have adopted pollen for sperm dispersal, and have therefore lost the ciliary dynein motor proteins that drive flagellar beating. Cytoplasmic dyneins, which control rapid, minus end-directed organelle motility in animal and fungal cells, also appear to have been lost from most plant lineages. The fact that in the model genetic system Arabidopsis thaliana there are 61 genes encoding kinesin motors indicates a remarkable diversification of this class of motors. In fact, as outlined in a recent comprehensive review of plant kinesins, several families of kinesins, particularly those involved in long-distance organelle transport, are not found in plants, whereas others, such as the kinesin–7 and -14 families have expanded (Zhu and Dixit, 2012). Although kinesins have been found to be instrumental for the construction of microtubule arrays, such as spindles and phragmoplasts, the kinesin-dependent motility of organelles has been difficult to identify, and in some cases the microtubule involvement in the function of the kinesins characterized remains unclear.

In this article, we explore microtubule–membrane interactions, including endomembrane and organelle trafficking, as well as plasma membrane connections. We begin with an overview of the plant secretory system then discuss the relative importance of actomyosin-mediated trafficking, before reviewing what is emerging about the role microtubules play in organelle positioning and function. We highlight cortical microtubule involvement in the modulation of cellulose synthesis and auxin polar transport via the tethering of cytoplasmic cellulose synthase complexes, and the retromer complexes that traffic PIN2 auxin efflux carriers, respectively. Finally, we discuss the importance of microtubule–membrane interaction for the self-organization of microtubule arrays during critical stages of plant cell development.

Secretory Organelles Undergo Remodeling of Constituents and Shapes

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

The plant cell secretory pathway is made of functionally interlinked and morphologically distinct organelles that are highly dynamic: not only are they for the most part largely motile, their membrane and lumenal constituents also undergo constant remodeling (Matheson et al., 2006). Membranes and proteins synthesized in the endoplasmic reticulum (ER) are subsequently exported to the Golgi for modification and delivery to post-Golgi organelles, which include the trans-Golgi network (TGN), prevacuolar compartments, vacuole and the plasma membrane (Figure 1). A retrograde endocytic flow of membrane and proteins from the plasma membrane counterbalances the anterograde movement of these constituents to maintain cellular homeostasis, and to allow cells to communicate with the external environment. The movement of membranes, which may occur through vesicles, tubules or direct connections, through the anterograde and retrograde routes, is coupled with the exchange of lumenal content between organelles. Despite such a flow of constituents among organelles, the secretory pathway maintains its dynamic homeostasis throughout the life of the cell.

image

Figure 1. Schematic illustration of the secretory pathway organization and protein trafficking routes in plant cells. Proteins are synthesized at the endoplasmic reticulum (ER), which is contiguous with the nuclear envelope (NE). From the ER, proteins are transported to the Golgi apparatus, which in plants is composed of multiple polarized stacks of cisternae. If proteins are not retrieved to the ER, they are transported to distal compartments, which include the trans-Golgi network (TGN) and the prevacuolar compartment (PVC). To reach the central vacuole, proteins can be transported from the PVC to the late PVC (LPVC), although direct pathways from the Golgi and the ER are known to exist. From the TGN, proteins can reach the plasma membrane, which is surrounded by the cell wall. The TGN is also involved in endocytic processes. A transport route from the Golgi to chloroplasts is also known to exist in plant cells.

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With little doubt, a peculiarity of the plant secretory pathway is the dynamism of the shape of most of its constituents. For example, the ER is composed of an interconnected network of tubules and cisternae that undergo continuous remodeling by tubule formation, elongation, sliding and fusion, as well as incessant streaming (Sparkes et al., 2009a, 2011; Ueda et al., 2011; Stefano et al., 2012). In plants, the Golgi apparatus is composed of numerous polarized stacks of membranous cisternae that are motile, and, at least in highly vacuolated cells, are attached to the underlying ER (Boevink et al., 1998; Moreau et al., 2007; Sparkes et al., 2009b). The TGN, by contrast, is a distinct organelle that can appear in association with the Golgi, but that can also be found as an independent organelle in the cytosol (Dettmer et al., 2006; Lam et al., 2007; Chow et al., 2008; Viotti et al., 2010; Contento and Bassham, 2012). Similarly, the prevacuole and late prevacuole compartments, which function as intermediate organelles for the transport of cargo and membranes to the vacuole, exist as motile organelles (Tse et al., 2004; Miao et al., 2008; Bottanelli et al., 2011). The central vacuole undergoes remodeling through the dynamic reshaping of its delimiting membrane: the tonoplast (Hicks et al., 2004; Reisen et al., 2005; Saito et al., 2005). The plasma membrane envelopes the entire cell and serves as a selective barrier between the cell wall and the cell interior. Although the plasma membrane is not motile, its make-up is continually remodeled with the acquisition of membrane and cargo through the exocytic pathway, as well as invagination and vesicle budding through the endocytic pathway. The lipid bilayer of the plasma membrane is also remodeled by lateral movement of cell wall building enzymes and other integral proteins, through dynamics that are constrained by the cell wall (Martiniere et al., 2012).

Involvement of Actin in Organelle Movement and Membrane Traffic

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

As it occurs for most of the plant organelles, the movement of plant secretory organelles is mainly dependent on the actin filament network, as demonstrated by evidence that the chemical inhibition of actin cables leads to the disruption of the remodeling of ER tubules (Sparkes et al., 2009a), and movement of the Golgi (Boevink et al., 1998; Nebenführ et al., 1999; Brandizzi et al., 2002; Akkerman et al., 2012) and the TGN (Asaoka et al., 2012). A recent study has shown that in elongated root cells that have completed growth, two distinct actin filament configurations co-exist: actin filament bundles that are interspersed with areas containing fine filamentous actin (F–actin) and areas without F–actin. In areas in which actin is bundled, the Golgi can reach maximal velocity (up to 7 μm s−1); however, in regions populated by fine F–actin, the Golgi exhibit a wiggling type of motility (Akkerman et al., 2012). These observations imply that the configuration of actin filaments has a direct relevance on organelle movement, at least for the Golgi. Although the actin cytoskeleton is important for movement, its disruption does not impair short-distance membrane flow in the early secretory pathway. For example, ER/Golgi protein exchange can occur in the absence of actin, as demonstrated by separate approaches in cells treated with actin-disrupting agents. Specifically, fluorescence recovery was reported in the Golgi after the photobleaching of fluorescent protein fusions of Golgi-localized proteins, and studies based on treatment with the traffic inhibitor brefeldin A showed the re-absorption of Golgi membranes into the ER (Brandizzi et al., 2002; Saint-Jore et al., 2002). Similar results were obtained in the presence of microtubule-disrupting agents (Brandizzi et al., 2002; Saint-Jore et al., 2002). On the one hand, it is likely that because of their close association, the ER and Golgi organelles do not require cytoskeletal elements to communicate; nonetheless, it is yet to be established whether ER–Golgi membrane exchange could be more efficient in the presence of an intact cytoskeleton. On the other hand, the lack of an intact actin cytoskeleton is known to affect post-Golgi traffic to the vacuole (Kim et al., 2005), perhaps because intermediate organelles cannot move efficiently towards their respective target compartments in the absence of actin.

Involvement of Myosins in Organelle Movement

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

Myosins are the motor proteins driving movement along actin filament tracks. In the Arabidopsis thaliana genome there are numerous myosin-encoding genes (17), which have been categorized by phylogenetic analyses into two plant-specific subfamilies: myosin VIII, with four members; and myosin XI, with thirteen members (Reddy and Day, 2001). Although the subcellular localization of most myosins has yet to be established (Reisen and Hanson, 2007), immunocytochemical analyses with an antibody for tobacco 175–kDa myosin (a homolog of Arabidopsis myosin XI) showed the distribution of this myosin with the cortical ER in BY–2 cells (Yokota et al., 2009). These data are in agreement with biochemical analyses on extracts from Arabidopsis seedlings showing that myosin XI–K co-fractionated with the ER (Ueda et al., 2011) as well as with live cell imaging of a functional fluorescent protein fusion to myosin XI-K, which was found to associate with structures that aligned and co-fractionated with an ER subdomain in Arabidopsis (Peremyslov et al., 2012). They are also consistent with earlier immunolocalization studies with the giant internodal cells of characean algae, which identified a Ca2+-dependent association of myosin with the prominent actin subcortical actin bundles (Grolig et al., 1988), as well as its association with motile nuclei (Wasteneys et al., 1996) and the ER tubules (Foissner et al., 2009). Characean myosins fall into the myosin–XI class (Ito et al., 2007), consistent with the common adoption of myosin-based cytoplasmic streaming by algal and plant lineages.

It is yet to be shown whether myosins interact directly with organelle membranes or indirectly by means of receptors/adaptors, such as RAB-GTPases, which control myosin–organelle linkage in mammalian cells (Seabra and Coudrier, 2004; Roland et al., 2011). Nonetheless, the localization data demonstrate that class–XI myosins are associated with secretory organelles, and are likely to influence their movement. In support of this, live-cell imaging-based analyses of truncated versions of all the myosins in A. thaliana and Nicotiana spp. revealed that when overexpressed, the tail fragments of myosins MYA1, MYA2, XI–C, XI–E, XI–I and XI–K can exert a negative effect on the movement of Golgi stacks and mitochondria (Avisar et al., 2008, 2009). Although these results highlight that myosin motors are in place to move plant secretory organelles, subsequent analyses in Arabidopsis knock-outs of XIC, XIE, XI–K, XI–I, MYA1 and MYA2/XI–2 showed that only the loss of myosin XI–K leads to the disruption of Golgi movement (Avisar et al., 2012). In addition, myosin XI–K was shown to have a role in the movement of the ER (Ueda et al., 2011). The dependence on myosin XI–K for the dynamics of post-Golgi organelles, such as the TGN, endosomes and prevacuole, has also been shown through the analysis of mutants (Avisar et al., 2012). The evidence that cytoplasmic streaming is inhibited in the myosin XI–K mutants suggests that this myosin is a major player in cytoplasmic motility in plant cells in general. One of the major challenges in characterizing myosin function is the high degree of redundancy. Major developmental defects only emerge when triple or quadruple mutants of class–XI myosins are generated (Peremyslov et al., 2010).

Role of Kinesins and Microtubules in Organelle Positioning

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

The microtubule cytoskeleton appears to have very little involvement in organelle movement and cargo transport in the secretory pathway of plant cells, which is in remarkable contrast to other eukaryotic systems. For example, microtubules have prominent roles for Golgi integrity and ER dynamics in mammalian cells (Ward and Brandizzi, 2004; Friedman et al., 2010). Recently, however, microtubules have been found to be important for the motility and morphology of the ER located in the cortex of characean algal cells (Foissner et al., 2009). This involvement is limited to the early stages of cell elongation. In addition, microtubules, which are mainly confined to the cortex in these giant cells, do not appear to have any direct involvement in the rapid motility of the ER network found deeper in the cytoplasm (Foissner et al., 2009), in contrast to a previous finding that they contribute to mitochondrial motility (Foissner, 2004). It remains to be determined whether analogous microtubule-dependent cortical ER organization occurs in the cells of land plants, although there is evidence for a strong ER association with microtubule preprophase bands (Giannoutsou et al., 2012). Despite a lack of evidence for microtubule involvement in the long-distance trafficking of organelles or endomembrane compartments in land plants, there is mounting evidence that some microtubule-associated motor proteins of the kinesin superfamily are involved in some activities of the plant secretory pathway. As outlined in two recent review articles, although rapid organelle distribution, characterized by long-distance transport, is clearly dependent on myosin motor activity on the actin filament network, it has been suggested that microtubules and their motors could specialize to anchor or slow down organelles such that they are targeted to functionally appropriate subcellular destinations (Cai and Cresti, 2012; Zhu and Dixit, 2012).

In general, kinesins have been implicated in a variety of important cellular events, including unidirectional transport of vesicles and organelles, cytokinesis, signal transduction, morphogenesis as well as key steps during cell division (Lee and Liu, 2004; Hirokawa et al., 2009; Li et al., 2012; Zhu and Dixit, 2012). Consistent with this, during cell division the kinesin AtPAKRP2 is involved in mediating the transport of Golgi-derived carrier materials to the phragmoplast (Lee et al., 2001). Another kinesin, AtKinesin-13A, has been found to be distributed over the entire Golgi apparatus in Arabidopsis trichomes and cotton fibers (Lu et al., 2005; Wei et al., 2009). A lack of AtKinesin–13A was reported to lead to the aggregation of Golgi stacks in trichomes as well as to a sharp decrease of the size and number of Golgi vesicles and changes to the morphology of Golgi stacks in root-cap peripheral cells, but not in the meristematic cells and columella cells of the root cap (Lu et al., 2005; Wei et al., 2009). These data indicate that, despite their large number, plant kinesins have relatively few and highly specialized functions. The data also indicate that, at least in some cell types, AtKinesin–13A may be involved in Golgi-related processes, including either Golgi morphology and/or membrane transport.

Although the studies described so far imply that motors associated with microtubules have roles in Golgi distribution and function, some caution is warranted because it is possible that some plant kinesins have acquired non-conventional functions that are unrelated to microtubules. For example, two members of the kinesin–14 family, AtKAC1/KLP2/KCA1/GRIMP/KSN1OsKCH1 and AtKAC2/KCA2 have been implicated in actin-based chloroplast movement; furthermore, OsKCH1 simultaneously binds to actin filaments and controls nuclear positioning and the onset of mitosis (Frey et al., 2009, 2010). One rice kinesin, GDD1, has transcription factor activity that controls gibberellic acid biosynthesis and cell elongation (Li et al., 2011); therefore, it cannot yet be excluded that the association of AtKinesin–13A with the plant Golgi may be related to actin-based activities. It may also be the case, however, that AtKinesin–13A has a dual function in the Golgi network. By way of precedent, it has been shown that a cotton kinesin, KCH1, interacts with actin filaments in developing cotton fibers, and plays a putative role in dynamic microtubule–microfilament cross-linking (Preuss et al., 2004). Similarly, another cotton kinesin, GhKCH2, has been shown to bind actin filaments and to cross-link them with microtubules in vitro (Xu et al., 2009). These data support a model whereby certain kinesins serve as linkers between the actin and microtubule cytoskeleton. This might explain previous studies in Nitella internodal cells, which determined that despite the fact that depolymerization of microtubules alone has no effect on cytoplasmic streaming, it can nonetheless increase 10–fold the sensitivity of cells to actin inhibitors, and dramatically delay recovery from the cytochalasin-induced cessation of cytoplasmic streaming (Wasteneys and Williamson, 1991; Collings et al., 1996). A similar synergy between the microtubule and actin cytoskeleton has been documented in Arabidopsis thaliana in terms of root cell expansion (Collings et al., 2006), suggesting that functional synergies between microtubule-based and actin-based motility systems are conserved.

Analyses of the movement of the plant Golgi have revealed that the stacks undergo stop-and-go movement, whereby phases of movement alternate with stationary phases (Boevink et al., 1998; Nebenführ et al., 1999). It has been suggested that during the stationary phases the Golgi receive cargo from the ER (Nebenführ et al., 1999). It was subsequently shown that ER-to-Golgi protein transport can occur while the Golgi are moving (daSilva et al., 2004), implying that the stationary phases of Golgi movement are not strictly necessary for ER export to the Golgi. Despite this finding, a stationary phase would be expected to tune an efficient and localized deposition of cell wall materials. For example, kinesins, such as AtKinesin–13A could function to anchor and to pause Golgi stacks through an interaction between actin filaments and cortical microtubules, which would facilitate post-Golgi traffic of cell wall enzymes and matrix in specific regions of the cell cortex. Recent findings that microtubules are not required for Golgi pausing, however, may challenge such a view (Hamada et al., 2012), and raise the intriguing question on whether kinesins may have other functions at the Golgi, including serving as part of a Golgi matrix, which has been invoked to anchor the ER and the Golgi (Sparkes et al., 2009b). Clearly, more experimental data are required to understand these processes.

Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

Most wall polysaccharides, including pectins and hemicelluloses, are synthesized by glycosyl transferases, and are often modified by other enzymes located in the Golgi. Cellulose and callose, in contrast, are produced at the plasma membrane from multi-enzyme complexes; thus it is the complexes themselves rather than the polysaccharide product that is delivered to appropriate sites at the plasma membrane. The stop-and-go activity of Golgi stacks described above is consistent with findings that Golgi carrying newly assembled cellulose synthase complexes pause at cortical microtubules, and that the insertion of cellulose synthase complexes in the plasma membrane is associated with these pauses (Crowell et al., 2009; Gutierrez et al., 2009). Endocytosis of cellulose synthase complexes, presumably a normal aspect of plasma membrane-associated enzyme recycling, can also be induced by osmotic stress, by chemicals that inhibit cellulose synthesis (Crowell et al., 2009; Gutierrez et al., 2009) or at the restrictive temperature for the conditional CesA1 allele rsw1–1 (Arioli et al., 1998; Chen et al., 2010; Fujita et al., 2013). Under certain conditions, these endocytosed cellulose synthase complexes have been shown to associate with microtubules in compartments referred to as small CesA-containing compartments (SMACs; Gutierrez et al., 2009) or microtubule-associated cellulose synthase compartments (MASCs; Crowell et al., 2009). Interestingly, the retraction of cellulose synthase complexes in the rsw1–1 mutant at restrictive temperatures is correlated with a massive decrease in the movement of Golgi and other organelles (Fujita et al., 2013), suggesting that the integrity of these enzyme complexes at the plasma membrane is somehow intimately linked to the general regulation of myosin-based intracellular motility.

The mechanism by which compartments carrying cellulose synthase complexes are tethered to microtubules, either during the delivery or retraction process, remains unclear. It is possible, however, that kinesins are involved. FRAGILE FIBER 1 (FRA1) of Arabidopsis (Zhong et al., 2002) and BRITTLE CULM 12 (BC12) of Oryza sativa (rice) are class–4 kinesins that were identified in mutant screens for mutants with increased tendency for organ breakage under bending stress (Zhong et al., 2002; Li et al., 2011, 2012). The fra1 and bc12 mutants have altered microfibril deposition patterns, indicative of reduced cellulose production (Sugimoto et al., 2001). Both of these motor proteins have been shown, through in vitro microtubule-binding ATPase assays, to have bona fide motor activity (Li et al., 2011, 2012; Zhu and Dixit, 2011), but whether they contribute to cellulose production through the anchoring of cellulose synthase compartments to microtubules awaits further analysis. It remains possible that this is achieved through distinct mechanisms, such as by modulating the dynamic properties and organization of cortical microtubules, or through the transcriptional regulation of gibberellin synthesis, as implicated for BC12 (Li et al., 2011), which also has a strong influence on microtubule orientation and wall expansion.

As cellulose synthase complexes show a strong association with cortical microtubules (Paradez et al., 2006; DeBolt et al., 2007; Desprez et al., 2007; Chan et al., 2010; Gu et al., 2010), emphasis has been placed on a role for microtubules in cellulose deposition and orientation. Interestingly, there has been little consideration for a role in the deposition of other wall polysaccharides or their modification. Coordinated delivery of other wall components, including matrix polysaccharides, wall-modifying proteins and enzymes, could require microtubule-dependent delivery (Fujita et al., 2012).

Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

Cortical microtubules define regions in which the majority of cellulose synthase complexes (CSCs) track at the plasma membrane. According to a recent study, 63% of CSCs are coincident with microtubule domains (Fujita et al., 2011). In the mor1–1 temperature-sensitive mutant, the loss of microtubule polymer mass at the plasma membrane reduces the coincidence of CSCs at microtubule-rich domains to 48%. This means that a higher proportion of CSCs are active in microtubule-free domains. Intriguingly, the average velocity of CSCs was significantly increased in mor1–1 at a restrictive temperature, as was the proportion of crystalline cellulose produced under these conditions (Fujita et al., 2011). Given that CSC displacement velocity is a proxy for the rate of cellulose polymerization, these measurements suggest that microtubule domains reduce the rate of cellulose microfibril synthesis, but produce cellulose of optimum quality (Fujita et al., 2012). Changes in lipid composition can increase or decrease the activity of many membrane-bounded enzymes (Spector and Yorek, 1985). It has therefore been proposed that microtubule domains might influence the activity of cellulose synthase complexes by modulating the lipidic composition of the plasma membrane (Fujita et al., 2011, 2012).

How might the presence of microtubules at the plasma membrane modify lipidic composition? Contact between the ER and the plasma membrane is one means by which lipid exchange might occur (Stefan et al., 2013). In this regard, the finding that cortical ER associates with the transversely oriented microtubules in early elongating internodal cells of the alga Nitella (Foissner et al., 2009) is certainly intriguing. Nevertheless an ER–microtubule association has not yet been observed in the elongating cells of land plants. Factors involved in lipid deposition at the plasma membrane, such as members of the synaptogamin protein family, are beginning to be characterized in plant cells (Schapire et al., 2008; Yamazaki et al., 2010; Zhang et al., 2011). Understanding the mechanisms controlling their spatial distribution and activity could provide clues as to how membrane lipid composition is determined.

CLASP-mediated Sorting Endosome Association with Microtubules

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

The elusive connection between endomembrane trafficking and microtubules became clearer this year, with the discovery that the microtubule-associated protein CLASP tethers the retromer-associated TGN/early endosomes to cortical microtubules via an interaction with sorting nexin 1 (SNX1; Ambrose et al., 2013; Figure 2a). SNX1 is a component of the retromer protein complex responsible for the recycling of the plasma membrane auxin efflux carrier PIN2 (Jaillais et al., 2006), and has been shown by immunogold transmission electron microscopy to localize to the TGN (Stierhof et al., 2013). In the absence of SNX1, PIN2 is trafficked to the lytic vacuole through the activity of the BLOC–1 complex, and is thus post-translationally regulated by the relative activities of the SNX1 retromer complex recycling pathway and the BLOC–1 lytic pathway (Cui et al., 2010). Ambrose and co-workers identified a direct interaction between the microtubule-associated protein CLASP and SNX1 based on both yeast two-hybrid and in vivo bimolecular fluorescence complementation analyses. In wild-type cells (Figure 2a), the CLASP-mediated tethering of SNX1 vesicles to microtubules is obvious under live imaging conditions. In contrast, SNX1 endosomes are scarce and their morphology is vastly different in mutants devoid of CLASP (Figure 2b), or when wild-type plants are treated with microtubule-disrupting drugs (Ambrose et al., 2013). The tethering of SNX1 compartments to cortical microtubules seen by live cell imaging resembles bunches of balloons on strings (Ambrose et al., 2013). This matches the description by transmission electron microscopy of the SNX1 and 2a-positive TGN as ‘bunches of grapes’ (Stierhof et al., 2013). Interestingly, endosomal association with cortical microtubules was also observed in living plant cells with the lipophilic dye FM4–64 (Ambrose et al., 2013), suggesting that the microtubule tethering of endosomes could be a general phenomenon.

image

Figure 2. CLASP-mediated microtubule-dependent recycling of the auxin efflux carrier PIN2. (a) PIN2 efflux carriers, after initial delivery to all regions of the plasma membrane, are endocytosed and trafficked to the trans-Golgi network (TGN), which becomes tethered to cortical microtubules via an interaction between the microtubule-associated protein CLASP and the retromer component sorting nexin 1 (SNX1). SNX1, through interaction with other retromer components, coordinates the redirection of PIN2 endosomes so that they become concentrated at one end of the cell. As depicted in this diagram, Golgi move along actin filament bundles to cortical sites, where they are seen to pause in close proximity to microtubules. (b) In the absence of CLASP, endocytosed PIN2, to a large extent, bypasses the recycling pathway and instead is trafficked to the lytic vacuole via the prevacuolar compartment (PVC). Thus, in clasp–1 mutants, PIN2 levels and auxin efflux activity are greatly reduced.

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PIN2 is responsible for tissue-specific polar auxin transport. Consequently, in the clasp–1 mutants with PIN2 levels dramatically reduced, there are unusual auxin accumulations in root tips and cotyledons (Ambrose et al., 2013). The altered auxin flow helps to explain the dwarf stature of clasp–1 mutants, their loss of apical dominance and other effects attributable to defects in auxin polar transport.

A major question is whether the CLASP-mediated tethering of endosomal compartments is responsible for the polar distribution of PIN2. PIN proteins normally accumulate at one face of cells to orchestrate polar auxin efflux, and so far this process has been demonstrated to rely on the activity of the ADP ribosylation factor guanine nucleotide exchange factor (ARF-GEF) GNOM (Steinmann et al., 1999), as well as the PINOID serine–threonine protein kinase (Friml et al., 2004). A recent study claims that PIN2 polarity is reversed in the cortex cells of clasp–1 mutants, and that this polarity reversal is apparently correlated with an increased abundance of PINOID (Kakar et al., 2013). This polarity reversal claim, however, is in direct contrast with the findings of the original study of PIN2 distribution in clasp–1 mutants, in which PIN2 is clearly shown to remain at the basal/shootward ends of the lateral root cap and epidermal cells, and at the apical/rootward ends of cortex cells in the division zone of the root (Ambrose et al., 2013). Although further analyses are needed to establish the nature of these contradictory results, a definite role of CLASP in microtubule–endosome tethering has now been uncovered, and the findings support the hypothesis that plants may have adapted microtubule-associated proteins to suit specific steps in trafficking.

Previous studies in animal cells have identified sorting nexin associations with microtubules via dynein, either through dynactin–dynein complexes in the case of SNX5 and SNX6 (Hong et al., 2009; Wassmer et al., 2009), or the WWC1/KIBRA (kidney and brain expressed protein) in the case of SNX4 (Traer et al., 2007). Although CLASP could have an analogous function to dynein in endosomal sorting, in plant cells CLASP connects sorting endosomes to microtubules in a relatively static manner, whereas in animal cells these complexes are trafficked towards the perinuclear Golgi complex (Stephens, 2012).

CLASP is a ‘plus’-end tracking microtubule-associated protein, and microtubules themselves can undergo rapid growth and shrinkage, but movement of PIN2-containing endosomal compartments by ‘plus’-end microtubule polymerization is unlikely to achieve large-scale transport. The movement of sorting endosomes from the longitudinal flanks to the end walls of cells, although possibly facilitated by tethering to cortical microtubules, is most likely achieved through myosin motor activity along actin filament networks, which tend to run parallel with the growth axis. Previous work demonstrated that perturbing the actin cytoskeleton with latrunculin B or cytochalasin D prevented the repolarization of PIN1 after treatment with the ARF-GEF-targeted exocytosis inhibitor brefeldin A (BFA), which causes PIN proteins to accumulate in intracellular compartments (Geldner et al., 2001).

The CLASP binding domain of SNX1 has been identified as the region between the N–terminal lipid-interacting PHOX domain and the C–terminal BAR domain, which mediates dimerization with other sorting nexins (Ambrose et al., 2013). The SNX1-interacting domain of CLASP, however, has not yet been revealed. Continuing studies will elucidate the specific mechanism by which CLASP interacts with sorting nexin, and once this is known it will be possible to determine whether this interaction is conserved among eukaryotes or whether it is unique to plants.

Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

The identification of links between microtubules and endomembranes could provide insight into the nature of cortical microtubule associations with the plasma membrane, which is critical to microtubule self-organization into functional arrays that define the axis of cell extension as well as division planes (Wasteneys and Ambrose, 2009). Recently, a flurry of live cell imaging integrated with mathematical modeling has provided insight into plant cortical microtubule self-organization (for a comprehensive review, see Eren et al., 2012). Connecting microtubules to the plasma membrane such that they assemble on a two-dimensional surface increases the likelihood that growing microtubules will encounter each other. When the growing ‘plus’ end of a microtubule encounters the side of an obstacle microtubule, this can generate rapid and irreversible depolymerization (catastrophe), entrainment (co-alignment of two or more microtubules), crossing-over or severing, depending on the angle at which the encounter occurs (Dixit and Cyr, 2004), as well as the degree to which the growing microtubule is anchored to the cortex (Allard et al., 2010a).

Mathematical modeling indicates that self-organization of microtubules into parallel arrays is positively correlated with encounter frequency, which in turn is dependent on highly dynamic microtubules. In the mor1–1 temperature-sensitive mutant, for example, microtubule growth (and shrinkage) rates are dramatically reduced when the temperature exceeds 28°C (Kawamura and Wasteneys, 2008). Computer simulations that incorporate the dynamic parameters of mor1–1 mutants measured by live cell imaging recapitulate the short microtubules and disordered arrays of these mutants, whereas inputs from wild-type cells generate parallel order (Allard et al., 2010b).

CLASP, the same protein that controls the tethering of SNX1 vesicles to microtubules (Ambrose et al., 2013), has been found to have an additional function in keeping cortical microtubules in close contact with the cortex (Ambrose and Wasteneys, 2008). Paradoxically, despite the reduced contact with the plasma membrane, cortical microtubule arrays are often hyperparallel in clasp–1 mutant cells. Mechanochemical modeling has determined that the closer the anchor point of a growing microtubule is to the obstacle microtubule, the less likely it is that an encounter will result in the two microtubules undergoing entrainment, and the more likely that it will result in catastrophe (Allard et al., 2010a). With fewer anchor points, microtubules in clasp–1 mutants have more freedom to explore three-dimensional space and entrain with obstacle microtubules at greater frequency, and at incident angles as high as 60°. When computer simulations were run using the live cell inputs from clasp–1 mutants, however, orientation patterns emerged that did not match the hyperparallel order found in vivo (Allard et al., 2010b). This led to a closer examination of the distribution pattern of CLASP in cells, and an extension of the mathematical modeling of cortical microtubule arrays to all six faces of a cell, rather than just on one-two-dimensional surface. CLASP was observed to preferentially accumulate at the sharp edges of the ends of newly divided cells, and this distribution was found to be essential for the formation of prominent transfacial microtubule bundles (Ambrose et al., 2011). As with microtubules encountering an obstacle microtubule at right angles, microtubules polymerizing against a sharp edge in the absence of CLASP invariably undergo catastrophe. With these inputs the three-dimensional modeling recapitulated the in vivo patterns, demonstrating that the concentration of CLASP activity to specific cell edges is critical to cell-wide array organization. In elongating cells, CLASP relocates to the longitudinal edges of elongating cells, consistent with the transverse parallel orientation of microtubules, and, when cells prepare for division, CLASP is confined to the sites at which preprophase bands intersect cell edges (Ambrose et al., 2011; Ambrose and Wasteneys, 2012; Dhonukshe et al., 2012).

The mechanism by which CLASP mediates microtubule interaction with the plasma membrane is unknown, and clearly there are additional, separate mechanisms. The lipid hydrolysing enzyme phospholipase D (PLD) is also implicated in microtubule attachment to the plasma membrane and cortical array organization (Gardiner et al., 2001, 2003; Dhonukshe et al., 2003; Motes et al., 2005). Strategies involving chemical genetic screening might eventually elucidate the mechanisms that characterize the various factors that control this quintessential process in plant cells.

Looking Back, Current Perspectives and Future Directions

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References

Fifty years ago, when Ledbetter and Porter first described microtubules at the cortex of plant cells, the startling images depicting a long-awaited scaffold for the deposition of cellulose initiated a quest to understand how the same elements that form spindle fibers in all eukaryotic cells could specialize to control cellulose deposition in plant cells (Ledbetter and Porter, 1963). At that time, the idea that microtubules could serve as tracks for motile elements was at least considered, but it is only in the past few years that we have begun to understand the mechanisms by which microtubules control activity at the plant cell cortex. The recent discoveries highlighted in this article, including the tethering of compartments carrying cellulose synthase complexes to microtubules, and the CLASP–SNX1 association that links microtubules to the recycling of the auxin efflux carrier PIN2, expand the known function of microtubules in plant cells. In addition, these newly identified mechanisms set the stage for a much broader understanding of endomembrane trafficking in plant cells.

There remain many gaps in understanding the processes that drive the plant secretory pathway. Despite much clearer evidence for myosin-mediated transport in plant cells, we still know very little about how myosin motors are connected to their cargoes. Future research needs to identify specific functions for different myosins, as well as the numerous microtubule motors and microtubule-associated proteins. Advances in cell imaging, including super-resolution microscopy, will aid in this direction, as well as reduce the distance that exists between findings obtained in live-cell fluorescence imaging and electron microscopy studies. Although the latter provides insights at a higher resolution than fluorescence microscopy, to date it has provided limited advances in deciphering the acto-myosin system, compared with microtubules. The surprising recent finding that the microtubule-associated protein CLASP has multiple roles, including the tethering of SNX1 vesicles to microtubules, linking microtubules to the plasma membrane, and promoting the formation of the transfacial microtubule bundles that coordinate cell-wide microtubule organization, suggests that despite its general conservation across all eukaryotes, this protein has adopted novel attributes as plants have evolved. Although clearly subordinate to the actin-based myosin motors, in terms of the rapid transport of organelles and vesicular cargo, plant kinesins are likely to have critical roles in the self-organization of microtubule arrays as well as specialized involvement in endomembrane trafficking events.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Secretory Organelles Undergo Remodeling of Constituents and Shapes
  5. Involvement of Actin in Organelle Movement and Membrane Traffic
  6. Involvement of Myosins in Organelle Movement
  7. Role of Kinesins and Microtubules in Organelle Positioning
  8. Microtubule-dependent Insertion and Tethering of Cellulose Synthase Complexes
  9. Do Microtubules Control Plasma Membrane Properties to Modulate the Activity of Cellulose Synthase Complexes?
  10. CLASP-mediated Sorting Endosome Association with Microtubules
  11. Microtubule–plasma Membrane Associations, Microtubule Organizing Centers at Cell Edges and the Self-organization of the Cortical Microtubule Array
  12. Looking Back, Current Perspectives and Future Directions
  13. References
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