CLASP: a microtubule-based integrator of the hormone-mediated transitions from cell division to elongation

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ScienceDirect CLASP: a microtubule-based integrator of the hormone-mediated transitions from cell division to elongation Yuan Ruan and Geoffrey O Wasteneys Plants use robust mechanisms to optimize organ size to prevailing conditions. Modulating the transition from cell division to elongation dramatically affects morphology and size. Although it is well established that auxin, cytokinin and brassinosteroid mediate these transitions, recent works show that the cytoskeleton, which is normally thought to act downstream of these hormones, plays a key role in this regulatory process. In particular, the microtubule-associated protein CLASP has a dual role in meristem maintenance. CLASP modulates levels of the auxin efflux carrier PIN2 by tethering SNX1 endosomes to cortical microtubules, which in turn fine tunes auxin maxima in the root apical meristem. CLASP is also required for transfacial microtubule bundle formation at the sharp cell edges, a feature strongly associated with maintaining the capacity for further cell division. Addresses The University of British Columbia, Department of Botany, 6270 University Blvd, Vancouver, BC V6T 1Z4, Canada Corresponding author: Wasteneys, Geoffrey O ([email protected])

Current Opinion in Plant Biology 2014, 22:149–158 This review comes from a themed issue on Cell biology Edited by Shaul Yalovsky and Viktor Zˇa´rsky´ For a complete overview see the Issue and the Editorial Available online 19th November 2014 http://dx.doi.org/10.1016/j.pbi.2014.11.003 1369-5266/# 2014 Elsevier Ltd. All right reserved.

Introduction — meristem maintenance and plant development All plant tissues originate from meristems located at shoot and root apices. Both the shoot apical meristem (SAM) and root apical meristem (RAM) are highly dynamic, integrating versatile developmental as well as environmental inputs into overall plant architecture [1,2]. The SAM is dome shaped and organized into three layers (L1– L3) and three zones (central, peripheral and rib) [3]. The central zone resides at the summit of SAM and slow division events occurring here serve to keep a constant number of stem cell initials. In contrast, peripheral cells sense positional cues, translating them into accelerated proliferation and directional growth as they are recruited www.sciencedirect.com

into rapidly outgrowing lateral primordia, defined by the surrounding growth-arrested organ boundary cells [4]. In the RAM, the stem cell niche comprises a mitotically inactive quiescent centre surrounded by initial cells that produce different tissues. Daughter cells undergo rapid elongation and differentiation when they are displaced from the root tip, a process that simultaneously builds the protective root cap and drives the root tip deeper into the soil [5]. Ultimately, meristem size control, which is coordinated by overlapping transcription factor networks [6], relies on the precise balance between maintaining a reservoir of undetermined cells and cell differentiation, giving rise to new tissues and/or organs through a wide spectrum of auxin-dependent activities and hormonal cross-talk [7,8,9,10,11]. In this article, we highlight recent findings that open new avenues to understanding the mechanisms that control meristem activity. In particular, we focus on how hormonal signals are both processed and integrated by cytoskeletal machinery.

Hormones, auxin transport and meristem maintenance Previous studies have focused on auxin and cytokinin, the two key participants in meristem development, providing evidence for an underlying complex hormone network. The auxin to cytokinin ratio is critical for meristematic cell fate determination, as evidenced by the fact that an auxin maximum at the root tip promotes cell division by upregulating cell cycle genes governing transition states, whereas cytokinin stimulates differentiation, thereby antagonizing auxin’s role. Conversely, relatively high auxin activities are required for lateral organogenesis while high levels of cytokinins are linked with maintenance of uncommitted cells in the SAM [12–14]. Recent studies have elucidated an intriguing involvement of the brassinosteroid (BR) hormone brassinolide (BL) in the morphogenesis of both the SAM and RAM. Low levels of BL or loss of BL perception in the SAM lead to deeper clefts between the main stem and axillary organs in contrast to the organ-fusion phenotypes, which result from loss of primordium boundaries in BL-hypersensitive mutants [15,16]. These studies establish the importance of BR homeostasis in the expression of boundary identity genes. By analogy, BL affects quiescent centre activity and promotes cell cycle progression in the RAM. Disturbance to normal BR signalling reduces RAM size either because of lower division rates in the initial cells or premature differentiation of meristematic cells [17,18]. Current Opinion in Plant Biology 2014, 22:149–158

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It was recently shown that the R2R3 transcription factor BRAVO (Brassinosteroids at Vascular and Organizing Center) is expressed within the QC and counteracts BR-stimulated QC division [19]. This discovery identifies a cell-specific molecular mechanism that explains how the BR signalling machinery can be dampened in a cell-specific manner, which in the case of the cells in the quiescent centre, is critical for maintaining stem cell identity in the surrounding region. Auxin gradients and local auxin maxima and minima, which drive the differential expression of genes for the body plan, are built up and retained mainly by ratelimiting auxin efflux carriers, most notably the PIN family proteins, which generate directional flow of auxin through their asymmetric (polar) distribution on the plasma membrane [20,21]. PIN1 in the SAM is positioned towards cells accumulating higher concentrations of auxin in the L1 layer and in the developing leaf vasculature, accounting for high auxin flow through the incipient organ promordia and drainage through developing vascular strands [22,23]. Recent work explains the defective leaf production in an erecta family gene triple mutant [24,25]. Attenuation of PIN1 causes homogenous accumulation of auxin in the L1 layer, and prevents its movement into the vasculature. The enlarged SAM in these mutants can be explained by either reduced recruitment of stem cells or the dramatic expansion of cells in the L1 layer [24,25] as a consequence of higher than normal auxin concentrations. In roots, the flow of auxin is critical to the maintenance of RAM identity and organ elongation. This upside-down fountain pattern is coordinated synergistically through the tissue-specific polar distribution of PINs 1, 2, 3, 4 and 7. Apically polarized PINs 1, 3, 4 and 7 in the stele direct auxin towards the root tip. PINs 3 and 7 account for lateral redistribution in the root cap. PIN2 is instrumental for basipetal movement of auxin through the epidermal layer as well as elongating lateral root cap and cortex tissues. Finally, PINs 2, 7 and 3 complete the loop by diverting shoot-bound auxin into the stele to maintain auxin homeostasis in the root tip [22]. It is now well established that perturbing polar auxin transport with chemical inhibitors or mutations affecting the activity of auxin transporters leads to auxin imbalances and concomitant meristem abnormalities. Modulating meristem activity in response to developmental programs and environmental cues is carried out by many complex and overlapping molecular mechanisms, the understanding of which remains far from clear. Consistent with meristem activity being integrated with the circadian clock, it was recently reported that PLT1/2 and consequently PIN1, 2, 3 and 7 gene expression are downregulated in the loss of function tic-2 mutant [26]. This reduces overall auxin abundance in the root, producing a Current Opinion in Plant Biology 2014, 22:149–158

smaller meristem by compromising stem cell competence to divide [26]. Perturbing just one PIN protein within the network can have similar consequences and may be of particular relevance to modulating meristem activity under changing environmental conditions. The recent discovery that the microtubule-associated protein CLASP has a role in controlling PIN2 levels post-translationally, identifies a previously unknown regulatory mechanism for meristem activity [27]. In mutants lacking CLASP gene expression, specific depletion of PIN2 perturbs the auxin reflux mechanism, causing auxin to pool in the meristem, a feature that is associated with precocious differentiation. This, and other related phenomena are discussed in the following section.

Auxin-driven microtubule entrainment It has long been known that the cytoskeleton serves to convert auxin signals into productive work such as unidirectional growth or organ bending in response to light, touch or gravity stimuli [28,29,30]. The findings of Xu et al. have clarified the underlying molecular mechanism of interpreting auxin as an extracellular messenger. In this study, auxin was shown to trigger the formation of the ABP1 (auxin-binding protein 1)–TMK (transmembrane kinase receptor like kinases) complex near the cell surface, which in turn activates the ROP GTPase (Rho Of Plants guanosine triphosphatase) signalling cascade to bring about fundamental changes in cytoskeleton configuration and cell shape [31] (see Figure 1a). An important insight into how ROP activity can generate microtubule alignment was revealed by Lin et al. [32], who determined that the ROP effector RIC1 interacts directly with and promotes the activity of the microtubule severing protein katanin. Evidently the release of newly nucleated microtubule branches facilitates their self organization through encounter-based entrainment. Computational simulations predicted a fourfold increase in the time to achieve parallel order without kataninbased severing and release of branch microtubule minus ends [33]. Subsequent experimental studies with katanin mutants have borne out this prediction, showing that either the normal establishment of parallel order [32] or realignment following changes in stress patterns [34] in leaf pavement cells or the blue light-induced shift from transverse to longitudinal orientation in hypocotyls [28] are delayed in katanin mutants. Consistent also with the idea that branch-form nucleation generates discordant microtubules for recruitment to new orientations, a recent study indicates that the loss of TON2/FASS-mediated branch form nucleation reduces the rate at which microtubules can reorient in response to hormone treatments [35].

Cytoskeletal control of auxin levels through PIN endocytosis More and more studies are demonstrating that the auxin–cytoskeleton relationship can be bidirectional, www.sciencedirect.com

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Figure 1

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Cytoskeletal regulation of PIN. (a) Leaf pavement cell. ROP6/RIC1 stabilizes microtubules at the neck region restraining outgrowth while ROP2/ RIC4 stabilizes actin filaments at the lobe tip promoting local growth. Stabilized F-actin inhibits PIN1 internalization. (b) Root cell. ROP6/RIC1 stabilizes actin filaments and inhibits PIN2 endocytosis. CLASP facilitates PIN2 recycling to the plasma membrane by direct interaction with SNX1, preventing it from BLOC-1 mediated vacuolar degradation.

incorporating complex feedback mechanisms that amplify signals or transport mechanisms. For example, auxin-mediated realignment of actin filaments can further consolidate auxin movement [36]. In the SAM, the close correspondence between microtubule orientation and PIN1 polarity suggests that microtubule-directed cellular expansion is integrated with auxin-induced organ outgrowth [37]. And finally, cellulose-dependent cell wall www.sciencedirect.com

integrity has been shown to be an important factor in controlling the polar distribution of PIN proteins [38]. Thus, PIN-dependent auxin transport, through its control of microtubule entrainment and consequent mechanochemical wall properties and cell shape, can reinforce its own polarity. Whereas cytoskeletal involvement in auxin responses as well as endocytosis-mediated control of auxin transport has been documented [39–41], whether Current Opinion in Plant Biology 2014, 22:149–158

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and how the cytoskeleton regulates auxin and auxin polar transport has remained obscure or controversial. Unlike the polar secretion mode described for epithelial cells [42], Arabidopsis PIN proteins are targeted symmetrically to both sides of the cell plate during cytokinesis under the guidance of the microtubule-based phragmoplast and subsequently, PIN polarity is achieved post-mitotically through retrieval from selective cell faces by clathrinmediated endocytosis (CME) [43,44]. In yeast and animal cells, the actin cytoskeleton is implicated in CME [45,46]. High resolution microscopy has revealed, for example, that actin patches help the endocytic process by constricting and elongating the neck of clathrin-coated vesicles in cultured mouse cells [46]. In plant cells, two prominent studies have elucidated the rudimentary role of actin in PIN endocytosis [47,48]. These articles show that stabilized actin filaments resulting from the activity of ROP GTPases and their effectors impede clathrin-mediated PIN protein internalization, with ROP2 and RIC4 controlling PIN1 retention in pavement cell lobe regions [47] and ROP6 and RIC1 controlling PIN2 in root cells [48] (Figure 1a,b). Integrating the knowledge that auxin can stimulate ROP GTPase activity, the auxin-ROP/RICPIN-auxin loop can be described as a self-regulating positive feedback system [31]. Although it was previously shown that RIC1 binds to and promotes microtubule ordering in leaf pavement cells [49], according to Lin et al., PIN2 retention on the plasma membrane is microtubule-independent [48]. Following internalization, PIN proteins are continuously shuttled between the plasma membrane and endosomal compartments. An intact actin cytoskeleton is necessary for both apical and basal targeting, despite the fact that delivery towards the apical side is more sensitive to actin depolymerization [50]. Vesicles carrying PIN proteins have been identified in the phragmoplast, resulting in an unbiased secretion to the expanding cell plate of dividing cells [43]. In interphase cells, short-term oryzalin treatment to destabilize microtubules was shown to have little or no effect on PIN polarity, suggesting that microtubules play no role in PIN localization [51]. Prolonged microtubule perturbation provoked polarity alteration preferentially for basal localization [50] but this could be due to profound changes to the growth axis and the complete blocking of cytokinesis. In 2013, the description of a function for microtubules, via the microtubule-associated protein CLASP, in the modulation of PIN2 protein levels [27] broadened our understanding of microtubule involvement in the fine tuning of polar auxin transport (Figure 1b). Maintaining PIN homeostasis is achieved both by endocytosis and proteolytic activity. The putative BLOC-1 complex in Arabidopsis mediates the transport of PIN2 for degradation in the vacuole through a direct interaction with SNX1 endosomes [52–54]. CLASP associates with SNX1 Current Opinion in Plant Biology 2014, 22:149–158

to increase endosome stability along microtubules, thereby facilitating the recycling of PIN2 to the plasma membrane and consequently impeding its transit to the vacuole [27]. In mutants lacking CLASP gene transcription, PIN2 is drastically reduced at the plasma membrane (whereas PIN1 quantity and polarity are unaffected), causing an expanded region of auxin accumulation in the root tip, consistent with impaired PIN2-dependent shootward reflux. Expanded auxin maxima in the RAM are correlated with meristem collapse [27,55,56]. BLOC-1 complex-related protein degradation is evolutionarily conserved in eukaryotes [57,58]. The CLASP-SNX-BLOC-1 mechanism’s role in auxin transport appears to be specific to PIN2 and does not influence PIN2 polarity establishment or maintenance. Maintaining the polarity of PIN proteins is also critical for auxin flux patterns. In this regard, the fluid nature of the plasma membrane and the integrity of the plant cell wall are both important. Notably, a deficiency in membrane sterol biosynthesis seems to obliterate PIN polarity [59]. High resolution imaging and computational simulation analysis suggest that non-mobile domains at the plasma membrane minimize PIN lateral diffusion. PIN proteins that occasionally escape from these regions can be retrieved by clathrin-mediated endocytosis [60,61]. One obvious mechanism for establishing plasma membrane heterogeneity is the direct targeting of ER-derived and/or Golgi-derived plasma membrane components but the lack of asymmetric distribution of cytoplasmic organelles suggests that other mechanisms are at play [51]. Indeed, it appears that the cell wall and, to some extent, the cytoskeleton can constrain lateral diffusion of PIN proteins, suggesting that the cytoskeleton may facilitate connections between the plasma membrane and the cell wall in order to create spatially discrete domains [38,62,63]. COBRA, a potential integrator of microtubules and the cellulosic cell wall [64], warrants further study.

Microtubule control of cell division and its link to cellular geometry The microtubule-based machinery that coordinates mitosis and cytokinesis is an essential feature of auxin-induced cell division and meristem maintenance. The breakdown and rebuilding of these arrays through the cell cycle are controlled by the activity of microtubule nucleating complexes, and various associated proteins (MAPs) that can promote polymer addition, catastrophe, severing and bundle formation. Recent reviews have already offered excellent information with respect to cytoskeleton and associated protein-dependent PPB formation and phragmoplast guidance, mitotic spindle assembly and the mitogen-activated protein kinase (MAPK) cascade during cytokinesis [65,66,67]. Hence, we focus here on emerging ideas about cell geometry-based microtubule organization and the potential role that it plays in meristem maintenance. www.sciencedirect.com

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Four distinct microtubule arrays are typically associated with dividing cells. They include the interphase cortical array (CMT) that reinforces the tissue axis between divisions, the preprophase band (PPB) that forms in G2 to determine the future division plane, the mitotic spindle that separates chromosomes and finally, the phragmoplast, which completes cytokinesis by building the cell plate [68]. The interphase cortical microtubule array situated on the periclinal and radial faces is stereotypically described as being transverse to the cell’s axis of elongation. While this is certainly common post-mitotically in the elongation zones of organs, it has been noted for some time now that the arrangement of interphase microtubules in cells within the division zone does not conform to this rule [69]. Recent studies, which we outline below, have identified three mechanisms that explain how these non-transverse patterns are established and hint at the functional relevance in maintaining cells in an undifferentiated state.

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Computational simulations of array establishment utilizing microtubule dynamics measurements have shown that cellular geometry can predict the orientation of CMTs [70]. In recently divided cells, the newly formed sharp edges are a formidable barrier to microtubules that encounter them, causing microtubules to undergo catastrophe. Consequently, microtubules become aligned parallel to the sharpest edges of cells. The microtubule-associated protein CLASP, which is enriched at the newly formed sharp cell edges, appears to overcome this default orientation pattern. In addition, CLASP is found along microtubules, and is involved in the anchoring of microtubules to the cortex. Paradoxically, more frequent microtubule detachment in clasp-1 mutants increases the likelihood that microtubule encounters will lead to bundle formation, contributing to a hyperparallel organization [71]. Thus, in mutants lacking CLASP altogether, microtubules assume transverse orientation very soon after division (Figure 2a). Importantly, this is strongly correlated with precocious exit of cells into terminal elongation, resulting in smaller meristems and shorter roots. In effect, the self-organizing capacity of microtubules can be predicted by the dynamic instability of microtubule polymers and the shape of the polyhedron in which they are assembling. By overcoming the catastrophe-inducing effects of the sharp cell edge, CLASP promotes the formation of robust transfacial bundles that account at least in part for the longitudinal microtubules that are seen at the outer periclinal face of root epidermal cells (Figure 2b). At least two other mechanisms involving microtubule nucleation contribute to non-transverse microtubule orientation patterns. Components of the g-tubulin ring complex are found at the newly formed transverse edges, and serve to generate microtubules that polymerize away from the edge in the longitudinal direction [72] (Figure 2c). Edge www.sciencedirect.com

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Non-transverse CMT arrays in post-cytokinetic cells. (a) clasp-1 mutants have transverse CMT (red lines) organization coincident with premature entry to elongation phase. The grey colour indicates the newly formed cell plate between adjacent cells. (b) CLASP enables CMT growth around sharp edges resulting in the formation of transfacial bundles. The blue colour represents CLASP and the adjacent red lines indicate transfacial bundles. (c) g-Tubulin ring complex-directed sharp edge nucleation generates CMTs in the longitudinal direction. The orange dots are GCP2/3 proteins, and microtubules growing away from them are shown in red colour. ‘‘+’’ is representative of microtubule plus ends. (d) Endoplasmic microtubules emanating from the nuclear surface-localized GCP2/3 splay outward to produce a bipolar CMT array when they reach the cell cortex. The brown colour designates nuclei, dotted red lines are endoplasmic microtubules and red lines are bipolar CMTs.

nucleation might aid in CLASP-dependent transfacial bundle initiation and/or maintenance. Consistent with this, GCPs disappear from edges as cells proceed into elongation and this is coincident with transfacial bundle disappearance from the transverse edges [72]. Radial microtubule arrays that form at the nuclear surface in early G1 are also recognized as a distinct array, generally thought to precede and help to populate the cortex with microtubules and nucleating complexes. A recent Current Opinion in Plant Biology 2014, 22:149–158

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study indicates that the G1 nucleus can also influence the orientation of cortical arrays [73]. As shown for epidermal cells of the root and leaf promoridia, immediately after division, perinuclear microtubule nucleating complexes spawn microtubules that, upon reaching the cell cortex, splay outward to form a bipolar array that is generally longitudinal with respect to the cell long axis (Figure 2d). Bipolar CMT arrays have been described in other recent studies [74–76] though any relationship between the nucleus and their formation has not been determined. The bipolar CMT array is a transient feature of recently formed cells, and is lost as nucleating activity at the nuclear surface diminishes. Intriguingly, the nucleus becomes active once again in populating the cell with a bipolar array in root hair-forming cells, and this generates the longitudinally aligned cortical microtubules that persist throughout elongation [73]. This suggests that the activity of microtubule nucleating complexes can alternate between the nucleus and the cortex in a development-dependent manner.

spindles and shorter phragmoplasts [55]. Despite these defects, clasp-1 mutants are still able to complete mitosis and cytokinesis with regular chromosome segregation and cell plate positioning, in contrast to the severe mitotic defects detected in clasp mutants of yeast and animal cells [81,82]. How SABRE affects microtubule structures through CLASP remains obscure, regardless of the synergistic phenotype of sab clasp-1 double mutant and the disrupted CLASP deposition in sab mutants during PPB formation. In meristematic cells of the RAM, SABRE is non-uniformly distributed on apical and basal plasma membranes, forming concentrated patches. It would be interesting to investigate the spatial relationship between these patches and the CLASP-enriched areas that give rise to transfacial microtubule bundles. Strikingly, the distorted cell plates observed in sab mutants are similar to those in mutants of TTP components, necessitating a sequential arrangement of TTP, SABRE, and CLASP guided division events.

Conclusion and future directions The transition to cell division To date, relatively little research has been conducted to assess the significance of the CMT pattern in the CMTPPB conversion yet this event establishes the cellular geometry that, as discussed above, influences post-mitotic CMT alignment. Recently, Spinner et al. identified a TTP (TON1-TRM-PP2A with FASS/TON2 as its regulatory subunit) protein complex as a positive effector for the CMT-PPB transition [77]. Previously characterized ton1 and fass mutants failed to form PPBs, yielding severely mis-positioned cell division and a loss of root organization [78,79]. The recently discovered TRMs (TON1 Recruiting Motif), a superfamily comprising 34 members [80], appear to function in directing TON1 and PP2A onto microtubules where the PP2A complex presumably exerts its phosphatase activity [77]. The critical role played by the TTP complex in PPB formation has been confirmed by mutational analysis of PP2A along with other subunits. These mutants exhibit impaired PPBs and cell plate misorientation. This work demonstrates that the TTP complexes found in acentrosomal plant cells are similar in composition to components of animal centrosomes. Importantly, it is still unclear how the TTP complex controls PPB formation but answers will come with the identification of PP2A substrates. Additionally, the large number of TRM members suggests functional diversification and tissue or organ specificity, which will require careful classification. The transmembrane protein SABRE has been identified as a factor in PPB, spindle and phragmoplast orientation as well as in cortical microtubule alignment, and evidence suggests that it acts upstream of CLASP [76]. CLASP has previously been shown to associate with mitotic and cytokinetic arrays and its depletion generates a series of disorders such as poorly developed PPBs, diamond-shaped Current Opinion in Plant Biology 2014, 22:149–158

Maintaining a population of stem cells, and the tissue initial cells that they give rise to, is a fundamental activity in all multicellular organisms. In plants, the number of cells within division zones is a critical determinant of organ size and growth rate. The key to modulating meristem populations appears to be the rate at which cells exit the division pathway and enter terminal growth and/or differentiation. The transition zones that define the border between dividing and elongating cells are now well known to be under the control of hormones, in particular auxin, cytokinin and brassinosteroids, as well as mechanical inputs that are derived both from the expanding organ itself and the environment that it encounters [34]. How some of these inputs converge to align microtubules and feedback upon themselves is shown in Figure 3. Although there are undoubtedly many players involved in developmental phase transitions, the microtubule-associated protein CLASP has generated much interest because it is important for both the integration of hormone signals and for fostering the formation of specific microtubule arrays that promote the continuation of cell division. In this article we highlighted CLASP’s ability to promote the formation of robust transfacial microtubule bundles, the construction of which is strongly associated with keeping cells mitotically active and inhibiting exit into the differentiation pathway. Overlaid with this is the intriguing and apparently plant-specific role that CLASP has in connecting SNX1 endosomes to microtubules, a process that maintains high levels and activity of the auxin efflux carrier PIN2, which is critical for cycling auxin in the root apical meristem. Thus, since auxin flux is a major determinant of meristem identity and size, CLASP has a second, very specific yet critical function in controlling auxin levels. www.sciencedirect.com

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Figure 3 Mechanical stress PIN reorientation Auxin

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Microtubule and cell wall-based feedback loops in auxin signalling. The black pathway indicates the activation of ROP/RIC and thus katanin-mediated microtubule orientation during auxin-induced expansion. CLASP-SNX1 and microtubule-mediated cell wall mechanical and chemical properties consolidate the function of auxin by promoting PIN stability as shown by the red colour. Microtubules reorient in response to mechanical stress (orange), which converge with auxin signalling at the ROP/RIC point. Meanwhile, mechanical stress (both intrinsic and extrinsic) modulates PIN polarity, influencing auxin distribution and action. Cellular geometry is determined initially by cell division planes, which are predicted by the formation of preprophase bands (PPB). This process (green pathway) involves recruitment of TON1 to microtubules by TRM as well as the activity of SABRE, which may act upstream of CLASP.

CLASP’s remarkable dual function in meristem activity corresponds closely with its expression, which is strongest in young, actively dividing tissues and tapers off as cells enter terminal differentiation [55,56]. How the expression of CLASP and/or CLASP protein stability is regulated is thus a critical next step for understanding meristem maintenance. Our own ongoing experiments are exploring a possible connection to the brassinosteroid signalling pathway. Intriguingly, the brassinosteroid receptor BRI1, like PIN2, has been identified in proteomic analysis of SNX1 endosomes [83]. In addition, our preliminary investigations indicate that clasp-1 mutants are hyposensitive to BL, just as they are to auxin. It is tantalizing to speculate that CLASP, which despite its similarity to homologues in other eukaryotic lineages [84], has evolved a unique function in plants to modulate the cross-talk between the hormones that mediate transitions from division to differentiation.

Acknowledgement The research from the Wasteneys laboratory is supported by the Natural Sciences and Engineering Research Council of Canada Discovery Grant 2014-06080. We thank Dr Chris Ambrose for many insightful discussions.

References and recommended reading Papers of particular interest, published within the period of review, have been highlighted as:  of special interest  of outstanding interest www.sciencedirect.com

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BR-stimulated cell division in the quiescent centre (also shown in [17]) but this activity switches off to replenish damaged stem cells. 20. Blilou I, Xu J, Wildwater M, Willemsen V, Paponov I, Friml J, Heidstra R, Aida M, Palme K, Scheres B: The PIN auxin efflux facilitator network controls growth and patterning in Arabidopsis roots. Nature 2005, 433:39-44. 21. Petra´sek J, Mravec J, Bouchard R, Blakeslee JJ, Abas M, Seifertova´ D, Wisniewska J, Tadele Z, Kubes M, Covanova´ M et al.: PIN proteins perform a rate-limiting function in cellular auxin efflux. Science 2006, 312:914-918. 22. Petra´sek J, Friml J: Auxin transport routes in plant development. Development 2009, 136:2675-2688. 23. Reinhardt D, Pesce ER, Stieger P, Mandel T, Baltensperger K, Bennett M, Traas J, Friml J, Kuhlemeier C: Regulation of phyllotaxis by polar auxin transport. Nature 2003, 426:255-260. 24. Chen MK, Wilson RL, Palme K, Ditengou FA, Shpak ED: ERECTA  family genes regulate auxin transport in the shoot apical meristem and forming leaf primordia. Plant Physiol 2013, 162:1978-1991. This study revealed that ERECTA family genes can regulate leaf primodium formation and SAM size. The reduced leaf production rate and broader meristem observed in the er erl1 erl2 triple mutant are related to altered PIN1 and auxin distribution. 25. Uchida N, Shimada M, Tasaka M: Modulation of the balance between stem cell proliferation and consumption by ERECTAfamily genes. Plant Signal Behav 2012, 7:1506-1508. 26. Hong LW, Yan DW, Liu WC, Chen HG, Lu YT: TIME FOR COFFEE  controls root meristem size by changes in auxin accumulation in Arabidopsis. J Exp Bot 2014, 65:275-286. The authors of this article found that a global reduction of PIN expression in the root, caused by the tic mutation, resulted in compromised auxin flow towards the tip and a smaller meristem. 27. Ambrose C, Ruan Y, Gardiner J, Tamblyn LM, Catching A, Kirik V,  Marc J, Overall R, Wasteneys GO: CLASP interacts with sorting nexin 1 to link microtubules and auxin transport via PIN2 recycling in Arabidopsis thaliana. Dev Cell 2013, 24:649-659. This work extends the current understanding of how microtubules and associated proteins are involved in polar auxin transport. PIN2-containing SNX1 vesicles are tethered to microtubules through a direct interaction with CLASP. Without CLASP, PIN2 is sorted into the BLOC-1 route for vacuolar degradation. This establishes a direct role for microtubules and CLASP in PIN trafficking thus auxin distribution. 28. Lindeboom JJ, Nakamura M, Hibbel A, Shundyak K, Gutierrez R,  Ketelaar T, Emons AM, Mulder BM, Kirik V, Ehrhardt DW: A mechanism for reorientation of cortical microtubule arrays driven by microtubule severing. Science 2013, 342:1245533. Light-induced establishment of longitudinal microtubule arrays in hypocotyl epidermal cells was shown to be promoted by katanin, which localizes to the sites at which branch-form microtubules emerge from earlier formed microtubules. 29. Nick P, Scha¨fer E, Furuya M: Auxin redistribution during first positive phototropism in corn coleoptiles: microtubule reorientation and the Cholodny–Went theory. Plant Physiol 1992, 99:1302-1308. 30. Holweg C, Susslin C, Nick P: Capturing in vivo dynamics of the actin cytoskeleton stimulated by auxin or light. Plant Cell Physiol 2004, 45:855-863. 31. Xu T, Dai N, Chen J, Nagawa S, Cao M, Li H, Zhou Z, Chen X, De  Rycke R, Rakusova H et al.: Cell surface ABP1-TMK auxinsensing complex activates ROP GTPase signaling. Science 2014, 343:1025-1028. This article reports on a mechanism for transmitting extracellular auxin signals into intracellular developmental programs. Auxin-dependent ABP1-TMK1 interaction activates ROP GTPases and their effectors, leading to cytoskeleton-mediated directional expansion. Note that PIN endocytosis is inhibited by ROP-stabilized actin filaments, amplifying the auxin response. 32. Lin D, Cao L, Zhou Z, Zhu L, Ehrhardt D, Yang Z, Fu Y: Rho  GTPase signaling activates microtubule severing to promote microtubule ordering in Arabidopsis. Curr Biol 2013, 23:290-297. Current Opinion in Plant Biology 2014, 22:149–158

This study links ROP GTPase signalling with katanin-mediated severing, which facilitates microtubule entrainment into neck-constraining bands in pavement cells. An observed genetic interaction between katanin and ROP6 led to the finding that the ROP6 effector RIC1, already known to associate with microtubules, can directly bind to katanin, and likely promote microtubule severing. 33. Allard JF, Wasteneys GO, Cytrynbaum EN: Mechanisms of selforganization of cortical microtubules in plants revealed by computational simulations. Mol Biol Cell 2010, 21:278-286. 34. Sampathkumar A, Krupinski P, Wightman R, Milani P, Berquand A,  Boudaoud A, Hamant O, Jo¨nsson H, Meyerowitz EM: Subcellular and supracellular mechanical stress prescribes cytoskeleton behavior in Arabidopsis cotyledon pavement cells. Elife 2014, 3:e01967. This article demonstrated the feedback at play between tissue-induced stress and microtubule orientation pattern formation in leaf pavement cells. This study also determined that katanin’s severing activity promotes the reorientation of microtubules following compression stress. 35. Atkinson S, Kirik A, Kirik V: Microtubule array reorientation in response to hormones does not involve changes in microtubule nucleation modes at the periclinal cell surface. J Exp Bot 2014. pii:eru325. 36. Nick P, Han MJ, An G: Auxin stimulates its own transport by shaping actin filaments. Plant Physiol 2009, 151:155-167. 37. Heisler MG, Hamant O, Krupinski P, Uyttewaal M, Ohno C, Jo¨nsson H, Traas J, Meyerowitz EM: Alignment between PIN1 polarity and microtubule orientation in the shoot apical meristem reveals a tight coupling between morphogenesis and auxin transport. PLoS Biol 2010, 8:e1000516. 38. Feraru E, Feraru MI, Kleine-Vehn J, Martinie`re A, Mouille G, Vanneste S, Vernhettes S, Runions J, Friml J: PIN polarity maintenance by the cell wall in Arabidopsis. Curr Biol 2011, 21:338-343. 39. Maisch J, Nick P: Actin is involved in auxin-dependent patterning. Plant Physiol 2007, 143:1695-1704. 40. Nick P: Probing the actin-auxin oscillator. Plant Signal Behav 2010, 5:94-98. 41. Paciorek T, Zazı´malova´ E, Ruthardt N, Petra´sek J, Stierhof YD, Kleine-Vehn J, Morris DA, Emans N, Ju¨rgens G, Geldner N et al.: Auxin inhibits endocytosis and promotes its own efflux from cells. Nature 2005, 435:1251-1256. 42. Mostov K, Su T, ter Beest M: Polarized epithelial membrane traffic: conservation and plasticity. Nat Cell Biol 2003, 5:287-293. 43. Geldner N, Friml J, Stierhof YD, Ju¨rgens G, Palme K: Auxin transport inhibitors block PIN1 cycling and vesicle trafficking. Nature 2001, 413:425-428. 44. Kitakura S, Vanneste S, Robert S, Lo¨fke C, Teichmann T, Tanaka H, Friml J: Clathrin mediates endocytosis and polar distribution of PIN auxin transporters in Arabidopsis. Plant Cell 2011, 23:1920-1931. 45. Yarar D, Waterman-Storer CM, Schmid SL: A dynamic actin cytoskeleton functions at multiple stages of clathrin-mediated endocytosis. Mol Biol Cell 2005, 16:964-975. 46. Collins A, Warrington A, Taylor KA, Svitkina T: Structural organization of the actin cytoskeleton at sites of clathrinmediated endocytosis. Curr Biol 2011, 21:1167-1175. 47. Nagawa S, Xu T, Lin D, Dhonukshe P, Zhang X, Friml J, Scheres B, Fu Y, Yang Z: ROP GTPase-dependent actin microfilaments promote PIN1 polarization by localized inhibition of clathrindependent endocytosis. PLoS Biol 2012, 10:e1001299. 48. Lin D, Nagawa S, Chen J, Cao L, Chen X, Xu T, Li H, Dhonukshe P, Yamamuro C, Friml J et al.: A ROP GTPase-dependent auxin signaling pathway regulates the subcellular distribution of PIN2 in Arabidopsis roots. Curr Biol 2012, 22:1319-1325. 49. Fu Y, Gu Y, Zheng Z, Wasteneys G, Yang Z: Arabidopsis interdigitating cell growth requires two antagonistic pathways with opposing action on cell morphogenesis. Cell 2005, 120:687-700. www.sciencedirect.com

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50. Kleine-Vehn J, Langowski L, Wisniewska J, Dhonukshe P, Brewer PB, Friml J: Cellular and molecular requirements for polar PIN targeting and transcytosis in plants. Mol Plant 2008, 1:1056-1066. 51. Boutte´ Y, Crosnier MT, Carraro N, Traas J, Satiat-Jeunemaitre B: The plasma membrane recycling pathway and cell polarity in plants: studies on PIN proteins. J Cell Sci 2006, 119:1255-1265. 52. Jaillais Y, Fobis-Loisy I, Mie`ge C, Rollin C, Gaude T: AtSNX1 defines an endosome for auxin-carrier trafficking in Arabidopsis. Nature 2006, 443:106-109. 53. Cui Y, Li X, Chen Q, He X, Yang Q, Zhang A, Yu X, Chen H, Liu N, Xie Q et al.: BLOS1, a putative BLOC-1 subunit, interacts with SNX1 and modulates root growth in Arabidopsis. J Cell Sci 2010, 123:3727-3733. 54. Kleine-Vehn J, Leitner J, Zwiewka M, Sauer M, Abas L, Luschnig C, Friml J: Differential degradation of PIN2 auxin efflux carrier by retromer-dependent vacuolar targeting. Proc Natl Acad Sci U S A 2008, 105:17812-17817. 55. Ambrose JC, Shoji T, Kotzer AM, Pighin JA, Wasteneys GO: The Arabidopsis CLASP gene encodes a microtubule-associated protein involved in cell expansion and division. Plant Cell 2007, 19:2763-2775. 56. Kirik V, Herrmann U, Parupalli C, Sedbrook JC, Ehrhardt DW, Hu¨lskamp M: CLASP localizes in two discrete patterns on cortical microtubules and is required for cell morphogenesis and cell division in Arabidopsis. J Cell Sci 2007, 120:4416-4425. 57. Hermann GJ, Scavarda E, Weis AM, Saxton DS, Thomas LL, Salesky R, Somhegyi H, Curtin TP, Barrett A, Foster OK et al.: C. elegans BLOC-1 functions in trafficking to lysosome-related gut granules. PLoS ONE 2012, 7:e43043. 58. Raposo G, Marks MS, Cutler DF: Lysosome-related organelles: driving post-Golgi compartments into specialisation. Curr Opin Cell Biol 2007, 19:394-401. 59. Willemsen V, Friml J, Grebe M, van den Toorn A, Palme K, Scheres B: Cell polarity and PIN protein positioning in Arabidopsis require STEROL METHYLTRANSFERASE1 function. Plant Cell 2003, 15:612-625. 60. Men S, Boutte´ Y, Ikeda Y, Li X, Palme K, Stierhof YD, Hartmann MA, Moritz T, Grebe M: Sterol-dependent endocytosis mediates post-cytokinetic acquisition of PIN2 auxin efflux carrier polarity. Nat Cell Biol 2008, 10:237-244. 61. Kleine-Vehn J, Wabnik K, Martinie`re A, Łangowski Ł, Willig K, Naramoto S, Leitner J, Tanaka H, Jakobs S, Robert S et al.: Recycling, clustering, and endocytosis jointly maintain PIN auxin carrier polarity at the plasma membrane. Mol Syst Biol 2011, 7:540. 62. Martinie`re A, Lavagi I, Nageswaran G, Rolfe DJ, Maneta-Peyret L, Luu DT, Botchway SW, Webb SE, Mongrand S, Maurel C et al.: Cell wall constrains lateral diffusion of plant plasmamembrane proteins. Proc Natl Acad Sci U S A 2012, 109:1280512810. 63. Martinie`re A, Gayral P, Hawes C, Runions J: Building bridges: formin1 of Arabidopsis forms a connection between the cell wall and the actin cytoskeleton. Plant J 2011, 66:354-365. 64. Roudier F, Fernandez AG, Fujita M, Himmelspach R, Borner GH, Schindelman G, Song S, Baskin TI, Dupree P, Wasteneys GO et al.: COBRA, an Arabidopsis extracellular glycosylphosphatidyl inositol-anchored protein, specifically controls highly anisotropic expansion through its involvement in cellulose microfibril orientation. Plant Cell 2005, 17:1749-1763. 65. Rasmussen CG, Wright AJ, Mu¨ller S: The role of the cytoskeleton and associated proteins in determination of the plant cell  division plane. Plant J 2013, 75:258-269. A comprehensive review of PPB prediction, formation with the cytoskeleton and its associated proteins as key determinants. 66. Masoud K, Herzog E, Chaboute´ ME, Schmit AC: Microtubule nucleation and establishment of the mitotic spindle in vascular  plant cells. Plant J 2013, 75:245-257. www.sciencedirect.com

A comprehensive review of microtubule-directed spindle assembly and dynamics. 67. Sasabe M, Machida Y: Regulation of organization and function of microtubules by the mitogen-activated protein kinase cascade during plant cytokinesis. Cytoskeleton 2012, 69:913-918. 68. Wasteneys GO: Microtubule organization in the green kingdom: chaos or self-order? J Cell Sci 2002, 115:1345-1354. 69. Sugimoto K, Williamson RE, Wasteneys GO: New techniques enable comparative analysis of microtubule orientation, wall texture, and growth rate in intact roots of Arabidopsis. Plant Physiol 2000, 124:1493-1506. 70. Ambrose C, Allard JF, Cytrynbaum EN, Wasteneys GO: A CLASPmodulated cell edge barrier mechanism drives cell-wide cortical microtubule organization in Arabidopsis. Nat Commun 2011, 2:430. 71. Ambrose JC, Wasteneys GO: CLASP modulates microtubulecortex interaction during self-organization of acentrosomal microtubules. Mol Biol Cell 2008, 19:4730-4737. 72. Ambrose C, Wasteneys GO: Cell edges accumulate gamma tubulin complex components and nucleate microtubules following cytokinesis in Arabidopsis thaliana. PLoS ONE 2011, 6:e27423. 73. Ambrose C, Wasteneys GO: Microtubule initiation from the nuclear surface controls cortical microtubule growth polarity  and orientation in Arabidopsis thaliana. Plant Cell Physiol 2014, 55:1636-1645. This article describes the formation of a bipolar CMT array when endoplasmic microtubules emanating from the nuclear surface reach the cortex in proximity to the outer periclinal wall. Together with CLASPdependent transfacial bundles [64] and edge-nucleated microtubules [66], these arrays contribute to the mixed CMT orientation pattern seen in post-cytokinetic cells. 74. Sambade A, Pratap A, Buschmann H, Morris RJ, Lloyd C: The influence of light on microtubule dynamics and alignment in the Arabidopsis hypocotyl. Plant Cell 2012, 24:192-201. 75. Vineyard L, Elliott A, Dhingra S, Lucas JR, Shaw SL: Progressive transverse microtubule array organization in hormoneinduced Arabidopsis hypocotyl cells. Plant Cell 2013, 25:662-676. 76. Pietra S, Gustavsson A, Kiefer C, Kalmbach L, Ho¨rstedt P, Ikeda Y, Stepanova AN, Alonso JM, Grebe M: Arabidopsis SABRE and CLASP interact to stabilize cell division plane orientation and planar polarity. Nat Commun 2013, 4:2779. 77. Spinner L, Gadeyne A, Belcram K, Goussot M, Moison M, Duroc Y,  Eeckhout D, De Winne N, Schaefer E, Van De Slijke E et al.: A protein phosphatase 2A complex spatially controls plant cell division. Nat Commun 2013, 4:1863. This study identified a TON-TRM-PP2A complex, which is homologous to animal centrosomal components. TON and PP2A are recruited to microtubules via physical association with TRM. Mutations in the complex lead to complete loss of or mis-oriented PPBs, suggesting that nucleation activity is essential for PPB formation. 78. Azimzadeh J, Nacry P, Christodoulidou A, Drevensek S, Camilleri C, Amiour N, Parcy F, Pastuglia M, Bouchez D: Arabidopsis TONNEAU1 proteins are essential for preprophase band formation and interact with centrin. Plant Cell 2008, 20:2146-2159. 79. Camilleri C, Azimzadeh J, Pastuglia M, Bellini C, Grandjean O, Bouchez D: The Arabidopsis TONNEAU2 gene encodes a putative novel protein phosphatase 2A regulatory subunit essential for the control of the cortical cytoskeleton. Plant Cell 2002, 14:833-845. 80. Drevensek S, Goussot M, Duroc Y, Christodoulidou A, Steyaert S, Schaefer E, Duvernois E, Grandjean O, Vantard M, Bouchez D et al.: The Arabidopsis TRM1-TON1 interaction reveals a recruitment network common to plant cortical microtubule arrays and eukaryotic centrosomes. Plant Cell 2012, 24:178-191. Current Opinion in Plant Biology 2014, 22:149–158

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81. Maiato H, Khodjakov A, Rieder CL: Drosophila CLASP is required for the incorporation of microtubule subunits into fluxing kinetochore fibres. Nat Cell Biol 2005, 7:42-47.

83. Jaillais Y, Fobis-Loisy I, Mie`ge C, Gaude T: Evidence for a sorting endosome in Arabidopsis root cells. Plant J 2008, 53:237-247.

82. Inoue YH, Savoian MS, Suzuki T, Ma´the´ E, Yamamoto MT, Glover DM: Mutations in orbit/mast reveal that the central spindle is comprised of two microtubule populations, those that initiate cleavage and those that propagate furrow ingression. J Cell Biol 2004, 166:49-60.

84. Al-Bassam J, Chang F: Regulation of microtubule dynamics by TOG-domain proteins XMAP215/Dis1 and CLASP. Trends Cell Biol 2011, 21:604-614.

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