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Plant Science 175 (2008) 747–755 Contents lists available at ScienceDirect Plant Science journal homepage: www.elsevier.com/locate/plantsci Review The roles of microtubules in tropisms Sherryl R. Bisgrove * Department of Biological Sciences, Simon Fraser University, 8888 University Drive, Burnaby, B.C., Canada V5A 1 s6 A R T I C L E I N F O A B S T R A C T Article history: Received 11 March 2008 Received in revised form 19 August 2008 Accepted 19 August 2008 Available online 7 September 2008 Plant tropisms, or growth towards or away from a stimulus, usually involve the bending of shoots or roots which reorient growth in a new direction. Plant responses to tropic cues, especially gravity and light, have been active areas of investigation for many years. Despite all of this attention we still do not understand how these responses are regulated. In this review possible roles for microtubules in tropisms are discussed. Tropisms occur in a series of steps; directional cues are perceived and converted into biochemical signals that induce bending in roots and shoots. One model suggests that microtubules function late in the response pathway, during organ bending. Microtubules have been linked to organ bending by virtue of their role in regulating the direction of cell elongation. In elongating cells microtubules appear to function as guides for the deposition of cellulose microfibrils into the cell wall and the placement of the microfibrils in the wall is thought to constrain the direction of cell elongation. According to the model bending occurs when different microtubule/microfibril alignments across the organ cause cells on the outer flank to elongate more than cells on the inner flank. In support of this idea is the observation that tropic signals can induce the appropriate changes in microtubule orientations across a bending organ. However, attempts to validate the hypothesis have produced conflicting results and the idea that microtubule alignment regulates cell expansion during organ bending is controversial. Microtubules have also been linked to organizational events associated with the plasma membrane. Although speculative, one possibility is that microtubules influence tropisms by positioning regulatory proteins and/or complexes in the plasma membrane. Two possible mechanisms by which microtubules could contribute to organizational events associated with the plasma membrane are outlined. In addition to cell expansion, microtubules are postulated to have roles in the perception of touch and gravity signals. Although microtubules are associated with touch sensing in animals, we know very little about the relevant receptors in plants. One way to assess how microtubules function during tropisms is to identify and study proteins that function in concert with microtubules. In particular, the analysis of microtubuleassociated proteins whose mutant forms confer defects in tropic responses promises to provide additional insights into the roles of microtubules in tropisms. ß 2008 Elsevier Ireland Ltd. All rights reserved. Keywords: Microtubule Cytoskeleton Cell expansion Cell wall Tropism Contents 1. 2. 3. 4. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Signal perception . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Auxin redistribution and differential growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Possible roles for microtubules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cortical microtubule organization and cell elongation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Cellulose deposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Microtubule reorientations and tropisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Beyond cellulose deposition: new roles for microtubules? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Summary and future directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . * Tel.: +1 778 782 5269; fax: +1 778 782 3496. E-mail address: [email protected]. 0168-9452/$ – see front matter ß 2008 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.plantsci.2008.08.009 748 748 748 749 749 750 750 751 753 753 753 748 S.R. Bisgrove / Plant Science 175 (2008) 747–755 1. Introduction Plants are sessile and cannot move when conditions become unfavorable. They can, however, redirect their growth to position stems, roots, leaves, and flowers towards the best possible locations. Shoots grow towards light, maximizing photosynthetic activity. Roots point down along the gravity vector and can be attracted to areas of higher moisture or nutrient content in the soil. When conditions change, plants respond by redirecting their growth in accordance with the new signals. Tropisms, these changes in the direction of growth in response to stimuli, involve the bending of stems or roots that reorient growth in the most favorable direction. Tropisms include responses to a number of cues including gravity (gravitropism), light (phototropism), and touch (thigmotropism), as well as gradients of moisture (hydrotropism), chemicals (chemotropism), and temperature (thermotropism). Although these tropisms have all been documented many have not been investigated in any further detail [1]. Gravitropism and phototropism have received the most attention, although some studies addressing hydrotropism and thigmotropism have been published in recent years [2–10]. Tropisms can be either positive or negative depending on whether growth occurs towards or away from the stimulus. The response occurs through a series of steps. Directional cues are sensed and converted into biochemical signals. Under normal conditions, plants receive many cues at once often from different directions. To cope with all of this information incoming signals must be integrated and transmitted to the responding cells where the changes in growth occur (see ref. [11] for a review). Gravity is a constant stimulus and changes in growth occur only when opposing cues out compete or over-ride gravity [11]. Because of the large amount of incoming information and the requirement for an integrated growth response, the signaling pathways that underlie tropisms are complex and they are not well understood. However, this is an active area of investigation and it is discussed in several recent reviews [1,6,11–19]. 1.1. Signal perception The receptors responsible for initiating phototropism, the phototropins, were the first to be described at the molecular level (see ref. [12] for a review). They are autophosphorylating protein kinases that are activated by blue light [12,17,20]. In addition to phototropins cryptochromes, another class of blue light receptors, and phytochromes, red/far-red reversible photoreceptors, also modulate phototropism [17,19,21]. Once activated, the phototropins transfer the light signal to proteins in downstream signaling pathways, but as of yet only a few of these intermediates have been identified [12,17,22]. Little is known about mechanoperception in plants, although studies in bacteria and animals are providing information about how mechanical stimuli are sensed in these organisms. The relevant receptors appear to be mechanosensitive ion channels [16]. These channels are transmembrane complexes that open in response to mechanical forces exerted on the plasma membrane. In some cases mechanical forces are thought to induce a conformational change in the channel that opens it while in other instances channels are opened indirectly via interactions with signaling molecules or cytoskeletal proteins (see ref. [16] for review). The first molecular characterization of mechanosensitive ion channel proteins in plant membranes has recently been published [7]. These MscS (mechanosensitive channel of small conductance)-like proteins form pressure-sensitive channels in the plasma membrane of protoplasts suggesting that they could be mechanosensors [7]. However, the physiological relevance of these channels in intact plants is unknown. The receptors that mediate gravity perception have not been identified, although two models that describe how gravity is sensed have been put forward in the literature. The starch-statolith hypothesis postulates that gravity-sensitive cells detect the falling of intracellular masses called statoliths (see refs. [18,23] for reviews). The gravitational pressure model proposes that the settling of the protoplast within the cell wall is sensed by receptors at the plasma membrane–extracellular matrix junction [18,24]. These receptors would be capable of detecting differences in tension and compression between the plasma membrane and the extracellular matrix at the top and bottom of the cell. Higher plant cells that can sense gravity commonly contain starch-filled amyloplasts that sediment and it is thought that this is the primary mechanism by which gravity is perceived, although it is possible that protoplast settling is also detected [18,23]. How amyloplast sedimentation is converted into a biochemical signal is not understood, although it has been proposed that mechanosensitive ion channels are involved [11,14,16,25,26]. With the exception of hydrotropism, plant responses to most other directional cues have been described mainly from a phenomenological perspective [1]. Recently genetic approaches have been used in efforts to understand the mechanisms that regulate hydrotropism [3–6,27,28]. Screens for mutants that are defective in hydrotropic responses have been done [29] and the characterization of the corresponding genes promises to yield insight into the molecular mechanisms that regulate hydrotropism [1]. 1.2. Auxin redistribution and differential growth Signals controlling tropisms often result in a concentration gradient of auxin across a responding organ and this auxin gradient is responsible for redirecting growth [17,19,30]. Consider, for example, a seedling that has been placed on its side. Auxin accumulates to a higher level on the lower flanks of the hypocotyl and root. According to the Cholodony–Went hypothesis [31], an auxin gradient triggers a differential growth response in which cells on one side of the organ elongate more than cells on the other side. Because the cells are held together and cannot move apart from one another, differential growth results in the formation of a Fig. 1. Bend formation in the hypocotyl and root of a seedling responding to light and gravity. In the seedling on the left, the shoot (shaded in green) grows up towards light; it is negatively gravitropic and positively phototropic. The root, on the other hand, has the opposite response; it grows down and away from the light. Cells in both the hypocotyl and the root expand mainly in the longitudinal direction (indicated by double-headed arrows). The seedling in the middle has been placed on its side and it now perceives a change in the direction of the light and gravity vectors. Cells on the lower flank of the hypocotyl elongate more than the cells on the upper flank (designated by large and small double-headed arrows respectively). This differential growth response produces a bend that reorients the shoot into an upright position (shown in the seedling on the right). A similar differential growth response occurs in the root, but in this case cells on the upper flank elongate more than the cells on the lower flank and the root bends down, towards the gravity vector. S.R. Bisgrove / Plant Science 175 (2008) 747–755 bend (Fig. 1). The Cholodony–Went hypothesis and auxin transport have received a considerable amount of attention in the literature and the reader is referred to several other articles for more detailed discussions [25,30,32–37]. The mechanism by which auxin induces differential cell elongation during organ bending is not understood, but it is thought that auxin alters the rate at which cells elongate by inducing changes in cell wall extensibility [38,39]. Ethylene also influences organ bending by a mechanism that is not well understood but appears to involve the modulation of auxin transport [36,40]. 1.3. Possible roles for microtubules How might microtubules contribute to these growth responses? Microtubules are thought to be involved in the differential growth response that leads to organ bending (see ref. [41] for review). Suggestions have also been made that microtubules function earlier in the gravity and touch pathways, possibly during signal perception or transduction [16,26,41]. These ideas and the evidence that supports them are discussed in this review. In addition, recent analyses that hint at new possibilities for how microtubules might function during tropic responses are presented. 2. Cortical microtubule organization and cell elongation Microtubules are long, tubule-shaped polymers of a- and btubulin. Microtubules are actually highly dynamic, although they appear stable in still photographs. They are almost always growing or shrinking, providing them the flexibility to rearrange into different arrays within the cell. Microtubules function in concert with a large group of microtubule-associated proteins or MAPs. MAPs bind to microtubules and while bound they modify microtubule dynamics or mediate microtubule interactions with other proteins or structures in the cell. The microtubule arrays that appear in higher plants include preprophase bands, spindles, and phragmoplasts in dividing cells, as well as the cortical arrays seen in elongating cells. Plant MAPs are discussed in detail in several recent reviews [42–46]. Microtubules are found in the periphery (or cortex) of elongating cells just below the plasma membrane where they are arranged in parallel hoops that encircle the cell (Fig. 2). Elongation occurs in a direction that is perpendicular to the cortical microtubules and the arrangement of the cortical microtubules is important for controlling the direction in which cells elongate (as 749 discussed below). When microtubules are disrupted by pharmacological or herbicide treatments or by mutation, cells stop elongating and swell by expanding radially (see refs. [47–53] for examples). In addition, there are several mutations that alter cortical microtubule organization and plants with these mutations have defects in the control of directional cell elongation. For example, microtubules become very short and cells expand radially when the temperature-sensitive mor1-1 mutants are grown at the restrictive temperature [51]. The lefty and spiral mutants comprise another class of microtubule mutants that have a twisted pattern of growth [54–57]. These mutant plants can organize their microtubules into parallel arrays, but instead of a transverse alignment, the arrays form shallow helices with either a right- or a left-handed orientation. Theoretically, twisted growth occurs because cells are expanding perpendicularly to microtubules that have a helical rather than a transverse orientation [58,59]. The radially swollen 6 (rsw6) mutants represent a third class of temperature-sensitive microtubule mutants that have defects in cell expansion. At the restrictive temperature rsw6 mutants have microtubules that are well organized into parallel arrays within each cell but the orientations of microtubules among neighboring cells are highly variable [60]. Apparently rsw6 mutants are defective in a mechanism that controls global microtubule array changes across cells. The examples described above all illustrate the importance of cortical microtubules in cell elongation. Although the mechanisms by which microtubules affect cell elongation are not completely understood, it is thought they exert their influence by modifying the cell wall and the wall then acts as a constraint for expansion [61,62,52]. Expansion is driven by water uptake into the vacuole. Water entering the vacuole causes an increase in volume that exerts turgor pressure on the cell wall. When turgor exceeds the resistance imposed by the wall the cell expands (see refs. [62–66] for reviews). Cellulose microfibrils and the linkages that hold them together are thought to be the major load-bearing elements in the wall. Because long and intact microfibrils are resistant to stretching along their lengths, the wall yields to turgor by moving adjacent microfibrils apart from one another (Fig. 2). Cellulose arrays in the wall, therefore, constrain expansion to a direction that is perpendicular to the microfibril orientation. The microfibrils are embedded in a matrix of hemicelluloses, pectins and proteins, and the extensibility of the wall also depends on the level of crosslinking between these molecules and the cellulose microfibrils. When some of the connections that hold adjacent microfibrils Fig. 2. Cells expand in a direction that is perpendicular to cortical microtubules and cellulose microfibrils in the cell wall. (A) A confocal section taken through expanding root cells reveals cortical microtubules (green) in the upper cell that are aligned transverse to the long axis of the cell. The section has passed through more internal regions of the lower cells. (B) Cartoon showing cellulose synthases moving through the plasma membrane. Microtubules below the plasma membrane guide the synthases and the wall microfibrils are positioned parallel with the underlying microtubules. When the cell expands adjacent cellulose microfibrils move apart from one another and elongation occurs in a direction that is perpendicular to the microtubule/microfibril alignment (indicated by double-headed arrows in A and B). Microtubules in (A) were visualized in a transgenic Arabidopsis plant expressing a microtubule binding domain – green fluorescent protein fusion [144]. 750 S.R. Bisgrove / Plant Science 175 (2008) 747–755 together are disrupted, the wall becomes more extensible and cell expansion increases. Thus, the orientation of cellulose microfibrils in the wall and the degree of cross-linking largely determine the direction and the rates at which cells expand [61–68]. 2.1. Cellulose deposition Cellulose microfibrils are produced by cellulose synthase complexes embedded in the plasma membrane. The synthases travel laterally in the membrane, extruding microfibrils to the outside as they move [64–66]. This means that the arrangement of microfibrils in the wall is determined by the paths that are taken by the synthase complexes during cellulose synthesis. Microtubules appear to be involved in guiding synthase movement, but the mechanism is not completely understood. Cortical microtubules in elongating cells are closely associated with the plasma membrane and co-aligned with the microfibrils in the wall (Fig. 2). Based on this observation, the microtubule/microfibril co-alignment model postulates that cortical microtubules act as guides for the cellulose synthase complexes as they move through the plasma membrane (as reviewed in refs. [53,69,70]). This co-alignment model was accepted for many years as the textbook version [71] for how ordered arrays of microfibrils are deposited into the wall, even though direct evidence linking microtubules with cellulose synthase movement was lacking [69]. A direct connection between synthase movements and microtubules in elongating cells was finally established in 2006. Synthase complexes moving in paths that coincided with cortical microtubules were visualized in transgenic Arabidopsis plants expressing cellulose synthase – yellow fluorescent protein fusions [72]. Synthases were seen tracking along curved microtubules and when microtubules were reoriented the synthases altered their movement according to new microtubule positions. Although these observations clearly link synthase movement to microtubules, there is also evidence that suggests this is not the whole story. When microtubules are disassembled either by treatment with pharmacological or herbicidal agents, or by mutation, synthase complexes continue to move in tracks that are roughly parallel with each other and aligned arrays of microfibrils continue to be deposited into the wall [50,72]. How synthase movements might be guided in the absence of microtubules is not clear, but the findings do suggest that there is an alternative or additional guidance mechanism [53,69,73]. Cells with compromised microtubules lose the ability to elongate and they swell by expanding radially even though ordered arrays of cellulose are deposited into their walls [50]. This suggests that organized cellulose arrays alone are not sufficient for directional control of cell expansion and that microtubules may be influencing the wall in additional ways [50,53,73]. One explanation is that microtubule disruption leads to the deposition of short cellulose microfibrils [53]. According to the microfibril length regulation model, walls with shorter microfibrils will be less able to restrict cell expansion to one direction [53]. One could also speculate that microtubules control the spatial arrangement of additional molecules involved in cell wall modification and/or synthesis. If so, microtubule disruption could result in the loss or aberrant localization of molecules that stabilize or loosen the wall leading to altered cell expansion. Microtubules have been implicated in the localization of two other proteins involved in cellulose synthesis, KORRIGAN (an endo-1,4-b-D-glucanase involved in cellulose synthesis [74]) and COBRA (a glycosylphosphatidyl inositol (GPI) containing protein [75]). Both KORRIGAN and COBRA are distributed in linear arrays along the plasma membrane and treatment with the microtubule depolymerizing herbicide oryzalin disrupts this localization pattern [74,75]. 2.2. Microtubule reorientations and tropisms The concept that microtubules determine the orientation of cellulose microfibrils in the wall has been put forth as an explanation for how bend formation might be regulated [76,77]. Microtubule reorientations have been observed in the cells that are involved in bend formation in plants that are responding to tropic cues. Microtubules on the inner flanks of bending organs become oriented parallel with the long axis of the organ while microtubules on the outer sides are transversely oriented [76–81]. If cellulose synthase complexes follow microtubule tracks, then microfibrils will be deposited in the same direction as the reoriented microtubules. Since cells with transverse microfibrils elongate more than cells with longitudinal microfibrils, the end result will be organ bending [41,76,77,79,80]. In line with this idea is the observation that microtubules reorient in response to auxin [82–88]. Hence, the formation of an auxin gradient across a tropically stimulated root or stem could trigger microtubule/ microfibril reorientations that lead to bending [41,77,80]. These microtubule reorientations must also be coordinated across all of the cells in the bend. Although a model in which auxin-induced microtubule reorientations are responsible for organ bending has been suggested (see ref. [41] for a review) attempts to validate this hypothesis have produced conflicting results [41]. One set of experiments involves treating seedlings with agents that disrupt microtubule organization or function. In some cases tropic bending occurred while in other instances it was inhibited [84,89,90]. For example, in some experiments maize roots pretreated with microtubule depolymerizing agents were able to bend in response to a gravity stimulus [89,84], although the total amount of curvature was less than in untreated controls [84]. However, in other experiments treating maize coleoptiles with herbicides that depolymerized microtubules inhibited gravitropic bending even at low concentrations that only partially eliminated microtubules [90]. In contrast, phototropism proceeded even after complete removal of cortical microtubules [90]. Imposing mechanical strain on an organ can also cause microtubules to reorient [91]. This raises the possibility that the reorientations during tropic responses result from the mechanical strain produced when the organ bends. When mechanical counterforces were used to prevent or reverse tropic curvatures in maize coleoptiles and bean epicotyls, microtubules reoriented in response to the mechanical strain rather than the tropic stimuli [91–93]. On the other hand, microtubules in maize coleoptiles did reorient in response to gravity when bending was physically prevented, suggesting that tropic stimuli can induce microtubule reorientations in the absence of a bend [80]. Issues have also been raised regarding the timing of the microtubule reorientations that are proposed to result in differential growth and organ bending. In some cases microtubule reorientations that preceded organ bending were observed. For example, in gravity stimulated maize coleoptiles [80] and cut snapdragon spikes [94] microtubule reorientations occurred before the organs bent. However, microtubule reorientations do not always precede organ bending. For example, in gravity stimulated maize roots organ bending was initiated before microtubule reorientations occurred [78]. In addition, microtubule reorientations that preceded organ bending were not consistently observed when maize coleoptiles responding to light and gravity stimuli were analyzed [95]. Another issue that has been raised is based on the idea that changes in cell elongation rates due to microtubule reorientations are likely to be time-consuming as they involve reinforcing the cell wall through cellulose synthesis after the microtubule reorientations occur [96]. In support of this idea, S.R. Bisgrove / Plant Science 175 (2008) 747–755 correlations of cell elongation rates and microtubule/microfibril alignments in elongating Arabidopsis roots indicate that microfibrils remain transversely aligned until well after microtubules have reoriented [97]. Since the changes in cell elongation rates that occur during plant tropisms are initiated quickly, it has been argued that microtubule reorientation is unlikely to play a role in the early stages of bend formation even if the reorientations occur before the bend forms [96]. However, another suggestion is that microtubule reorientations play a role later in bend formation, perhaps as a mechanism to reinforce bending once it is initiated [78]. Given all of the results discussed above, a simple model in which auxin-induced microtubule reorientations lead to differential growth and organ bending is unlikely, although microtubules could influence cell expansion and organ bending in other ways. In addition to affecting microtubule organization, auxin also alters gene expression and ion homeostasis in cells [33–36,38,98– 102]. According to the ‘‘Acid Growth Theory’’ [38,98] auxin stimulates cell expansion by triggering the extrusion of protons into the wall. Acidification activates pH-sensitive enzymes and these proteins increase cell expansion by cleaving the linkages between polysaccharide components in the wall [38,39,98]. Another well-known effect of auxin is its ability to alter gene transcription and it follows that a gradient of auxin across a stem or root would result in the expression of different sets of genes on each side of the organ. Proteins that enhance cell expansion are synthesized preferentially on the faster growing flank where they can stimulate growth. The cell wall loosening proteins known as expansins localize preferentially to the more rapidly expanding outer flank of gravistimulated maize roots [103]. Expression analyses in Brassica oleracea seedlings have identified a set of tropically stimulated genes that are preferentially expressed in the cells where auxin levels are the highest. Among the up-regulated genes were enzymes that have been associated with roles in cell wall expansion [33,68]. These newly synthesized proteins must be transported to their sites of action in the plasma membrane or cell wall. Although speculative, it is possible that microtubules are involved in positioning these molecules in the proper places at the appropriate times (see Fig. 3 and discussion below). Note, in this scenario microtubule disruption would not necessarily prevent localization to the plasma membrane or cell wall, although it could affect placements with respect to space and/or time. 3. Beyond cellulose deposition: new roles for microtubules? Several lines of evidence suggest that microtubules could be involved in organizational events associated with the plasma membrane [104]. As mentioned above, the localization of both COBRA and KORRIGAN, two proteins with roles in cell wall biosynthesis, are microtubule-dependent [74,75]. Although these proteins could be part of cellulose synthase complexes, their localization in the plasma membrane is microtubule-dependent. A link between microtubules and the localization of an arabinogalactan cell surface GPI-linked protein from tomato has also been reported [105]. Microtubule disruption caused a relocalization of this protein [105]. GPI-anchored proteins have been associated with protein-containing membrane microdomains or lipid rafts in the plasma membrane [106–108]. Evidence from the animal literature suggests that lipid rafts may have roles in signaling processes and it is thought that they could serve as centers for signaling cascades [109,110]. It has been proposed that similar membrane microdomains may also be present in plant cells [111,112]. Although speculative, one possibility is that microtubules play a role in positioning signaling proteins and/or complexes in the plasma membrane in a manner similar to the 751 mechanism by which cellulose synthase complexes are guided during cellulose deposition (Fig. 3A and B). In such a scenario, microtubules would influence cell elongation and organ bending by coordinating the positions of signaling molecules in the plasma membrane with the structural components of the wall that are modified during the response (Fig. 3B). In animal and fungal cells the rapidly growing or plus-ends of microtubules also interact with proteins localized in the cortex of the cell next to the plasma membrane (reviewed in refs. [113– 115]) and there is evidence that suggests microtubule plus-ends have roles in plant tropisms. Arabidopsis plants carrying mutations in the microtubule plus-end binding protein END BINDING 1 (EB1) are slower to initiate bends after touch and/or gravity stimuli [116]. The delay appears to be specific for the initiation of differential growth as mutants grow at the same rate as wild type plants. Mutant roots have a tendency to grow in loops on tilted agar surfaces, indicating that they also have difficulty turning off differential growth [116]. How might EB1 influence differential growth during tropic responses? Although the mechanism is unknown, EB1 proteins have been the object of intense scrutiny in animal and fungal cells and results from these studies could provide insight into the roles of EB1 in plants. EB1 accumulates preferentially on the rapidly growing (or plus) ends of microtubules in all of the organisms in which it has been studied [113,115,117–120]. While bound to the microtubule end EB1 proteins usually make microtubules more dynamic [113]. The increase in dynamics is thought to help microtubules search the cytoplasm for ‘‘capture’’ sites. When an appropriate site is encountered EB1 mediates interactions between the microtubule end and other proteins localized at the site [113,121]. Capture sites often contain actin and EB1 is known to mediate interactions between microtubules and actin in some cell types (reviewed in ref. [113]). The interactions of microtubule ends with capture sites serves several functions including the targeted delivery of vesicles, signaling molecules, and ion channels to specific places in the cell [113,114,122–131]. Whether a similar system operates in plant cells is not known. However, one could speculate that the interaction of microtubule ends with specific sites in the cortex of the cell plays a role in tropic responses (see Fig. 3C). A captured microtubule could direct the insertion, removal, or position of proteins and/or complexes at specific sites in the cortex of the cell by serving as a track along which cargo-bearing microtubule-based motors like kinesins would travel. Alternatively, interactions between microtubule ends and capture sites could also be transient in nature. Microtubule plus-end binding proteins like EB1 appear to serve as platforms for recruiting signaling molecules to the active ends of microtubules (reviewed in ref. [113]). In this case a brief encounter at an appropriate capture site would be sufficient to deliver a specific factor. The idea that microtubules might be involved in the spatial integration of endo- and exocytotic events has been proposed [104]. In addition to a role in the localization of the secreted proteins COBRA, KORRIGAN, and the arabinogalactan protein discussed above, microtubules also appear to have roles in endocytosis in plant cells [132,133]. During preprophase band formation in cultured tobacco BY-2 cells endocytotic vesicles are internalized in trans-vacuolar cytoplasmic strands that form connections between the nucleus and the cortex of the cell. These cytoplasmic strands contain microtubules and treatment of cells with an herbicide that depolymerizes microtubules disrupts vesicle internalization [132]. Ligand-mediated endocytosis of a receptor involved in plant defense responses (the flagellin receptor FLS2) also utilizes microtubules [133]. Endocytosis of FLS2 is thought to be a regulatory step in the signaling pathway [133]. One could speculate that these microtubule-dependant endocytotic 752 S.R. Bisgrove / Plant Science 175 (2008) 747–755 Fig. 3. Possible roles for microtubules in plant tropisms. (A) According to the microtubule/microfibril co-alignment model microtubules serve as guides that influence the movement of cellulose synthase complexes during cellulose deposition. In response to tropic cues, there is a microtubule reorientation that has been proposed to change the direction of cellulose deposition and reduce the amount of cell elongation on the inner flank of a bending organ. Although the change in cell elongation rate may be too slow to play a role in bend initiation, the reorientations could be involved at later stages, perhaps to reinforce the bend once it is initiated. (B) Although speculative, it is possible that microtubules could constrain the positions of signaling proteins and/or complexes in the plasma membrane in a way that is similar to the mechanism postulated in the microtubule/microfibril co-alignment model. This would enable the plant to spatially coordinate structural components of the wall with the signaling molecules that trigger changes in wall extensibility during tropic responses. (C) Another way that microtubules could influence tropism is through interactions of their ends with specific sites in the cortex of the cell. Proteins that preferentially bind the more rapidly growing or plus-ends of microtubules, such as EB1, can mediate both microtubule dynamics and interactions of the microtubule end with other proteins or structures in the cell. EB1 tends to make microtubules more dynamic and this increase in dynamics is thought to facilitate microtubule searching of the cytoplasm for specific ‘‘capture’’ sites. In animal and fungal cells EB1 is also known to interact with several other proteins including signaling molecules. When the microtubule end encounters a suitable site proteins associated with the microtubule end (yellow balls) and proteins localized at the capture site (blue balls) can interact with one another. Capture sites often contain actin and EB1 is sometimes involved in crosstalk between the two cytoskeletal structures (for simplicity actin is not shown in A or B). Transient encounters would be sufficient for the release of factors associated with the microtubule end while stably connected microtubules could serve as tracks that direct the movement of cargo-bearing motor proteins (not shown) to and from the capture site. These interactions could play a role in tropisms by delivering signals or materials that are needed to regulate organ bending to the appropriate sites in the cortex of the cell. They could also be involved in signal perception either via connections to mechanosensory ion channels in the plasma membrane or by mediating the removal or insertion of signaling molecules such as receptors into or out of the plasma membrane. events involve the interactions of microtubule ends with cortical capture sites. Other plasma membrane proteins, including hormone receptors and ion channels, are also internalized in plant cells, although the cytoskeletal requirements for many of these endocytotic events have not been determined [134–136]. Microtubules have been implicated with roles in the perception of touch and gravity signals [16,26,41,90,95,137–139]. There are reports that microtubule inhibitors modify the rate of amyloplast sedimentation and inhibit auxin transport during gravitropism (reviewed in ref. [41]). The mechanism by which gravity is converted S.R. Bisgrove / Plant Science 175 (2008) 747–755 into a biochemical signal is unknown, although one possibility is that falling amyloplasts could be sensed by mechanosensitive ion channels [26]. Touch sensing is also thought to be mediated by mechanosensitive ion channels, a hypothesis that is based in part on analyses conducted on touch receptor neurons in the nematode worm Caenorhabditis elegans [16,26]. Genetic analyses indicate that mutations in tubulins associated with these neurons cause loss of mechanoresponse [140]. These microtubules are thought to form tethers that are linked, either directly or indirectly, to ion channels in the plasma membrane [141,142]. Force from a touch stimulus is thought to displace the microtubule tethers and this induces a conformational change in the channel that activates it [26,140,141]. Ca2+ channels have been implicated in touch responses in plants [9,16,26] and patch-clamping analyses have detected mechanosensitive calcium channels that are affected by microtubule inhibitors [143]. However, the molecular identity of these channels is unknown. Proteins that function as mechanosensitive ion channels in plants have recently been identified [7,10]. Whether these channels are involved in touch or gravity sensing and how their activities are regulated is unknown. 4. Summary and future directions Although microtubules have been associated with plant tropisms for many years our understanding of how they participate in these growth responses is far from complete. One proposal is that microtubules function during the differential growth response that leads to organ bending [41,76,77,79,80]. This idea is based on the concept that microtubules determine the direction of cell elongation by guiding the deposition of cellulose microfibrils into the cell wall. However, the hypothesis is controversial. In particular there are questions surrounding the timing of microtubule/ microfibril reorientations with respect to the onset of differential growth [78,80,94,96,97]. Several recent reports indicate that microtubules also have roles in organizational events associated with the plasma membrane [74,75,105,132,133]. In addition, plants carrying mutations in the microtubule plus-end binding protein EB1 exhibit delayed responses to touch/gravity signals [116]. Although speculative, these studies raise the possibility that microtubules could influence the wall in ways that are independent of a role in guiding cellulose deposition. Another possibility is that microtubules function in the perception of touch and gravity signals [16,26,41,90,95,137– 139]. There is evidence that indicates microtubules are involved in the activation of mechanosensory ion channels in animal cells [140,141,142] and microtubules appear to affect the activity of mechanosensitive Ca2+ channels in plant membranes [143]. However, mechanistic links between microtubules and the mechanoreceptors that might be involved in touch/gravity sensing are speculative. One way to assess how microtubules function during tropisms is to analyze proteins that interact with microtubules. Several microtubule-associated proteins have been identified in plants (see refs. [43,46] for reviews). Many of these proteins affect the stability or organization of microtubule arrays [46] and the corresponding mutants often have cell expansion defects that make it difficult to measure tropic responses. Because plants carrying mutations in EB1 genes have defects in touch/gravity responses they hold promise for providing further insights into the role of microtubules in tropisms. 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