Intact microtubules are required for the intercellular movement of the SHORT-ROOT transcription factor

Authors


For correspondence (e-mail gallagkl@sas.upenn.edu).

Summary

In both plants and animals, cell-to-cell signaling controls key aspects of development. In plants, cells communicate through direct transfer of transcription factors between cells. It is thought that most, if not all, mobile transcription factors move via plasmodesmata, membrane-lined channels that connect nearly all cells in the plant. However, the mechanisms by which these proteins access the plasmodesmata are not known. Using four independent assays, we examined the movement of the SHORT-ROOT (SHR) transcription factor under conditions that affect microtubule stability, organization or dynamics. We found that intact microtubules are required for cell-to-cell trafficking of SHR. Either chemical or genetic disruption of microtubules results in a significant reduction in SHR transport. Interestingly, inhibition of microtubules also results in mis-localization of the SHR-INTERACTING EMBRYONIC LETHAL (SIEL) protein, which has been shown to bind directly to SHR and is required for SHR movement. These results show that microtubules facilitate cell-to-cell transport of an endogenous plant protein.

Introduction

Multiple studies have shown that intercellular movement of proteins is widespread during plant development (Lee et al., 2006; Rim et al., 2011). This movement includes both endogenously encoded plant proteins, as well as those encoded by viral genomes. The movement of endogenously encoded proteins (e.g. CAPRICE, KNOTTED and SHORT-ROOT) is essential for the normal growth and development of the plant (Lucas et al., 1995; Nakajima et al., 2001; Kim et al., 2003; Kurata et al., 2005; Gallagher and Benfey, 2009). In the case of viruses, cell-to-cell movement is necessary for spread of the infection. It is therefore important that we understand the mechanisms by which trafficking occurs. For both plant proteins and RNA viruses, the problem of how to get from one cell to another is essentially the same. How does a cytoplasmically localized protein (or protein–RNA complex) move to the plasma membrane, cross the cell wall and then enter the neighboring cell? For movement via plasmodesmata (PD), this problem may be simplified to how do soluble, mobile proteins get to and pass through the PD? Multiple experiments have provided evidence that, for many proteins (and viral RNAs), the endomembrane plays a key role in promoting intercellular movement (Harries et al., 2010). Likewise, there are compelling (although sometimes apparently contradictory) data indicating that the cytoskeleton also promotes the cell-to-cell movement of viral RNAs (Boyko et al., 2007; Brandner et al., 2008; Harries et al., 2010; Ouko et al., 2010; Harries and Ding, 2011; Schoelz et al., 2011); whether the cytoskeleton is also implicated in the movement of endogenously encoded plant proteins is a question that needs to be addressed. In this paper, we examine the role of the microtubule cytoskeleton in the intercellular movement of a well-studied mobile transcriptional factor, SHORT-ROOT (SHR).

In the Arabidopsis root, the formation of two distinct ground tissue layers – the endodermis and cortex – requires cell-to-cell movement of SHR from the stele into the neighboring cell layer, chiefly the endodermis (Helariutta et al., 2000; Nakajima et al., 2001). This movement occurs through PD (Vaten et al., 2011) and is regulated by sequences within the SHR protein and through interactions with the SHR-binding protein, SHR-INTERACTING EMBRYONIC LETHAL (SIEL) (Gallagher et al., 2004; Gallagher and Benfey, 2009; Koizumi et al., 2011). Mutations in SIEL result in a decrease in SHR movement and defective development of the root ground tissue (Gallagher and Benfey, 2009; Gallagher et al., 2004; Koizumi et al., 2011; Vaten et al., 2011). As SIEL associates with endosomes and appears to be cell-autonomous, we proposed that SIEL promotes intercellular movement of SHR, essentially by acting as an intracellular shuttle (Koizumi et al., 2011). Similar models have been proposed for movement of plant viruses (Harries and Ding, 2011; Harries et al., 2009; Oparka, 2004; Wright et al., 2007). In particular, Lewis and Lazarowitz (2010) proposed that movement proteins (MP) encoded by certain species of virus [particularly cauliflower leaf curl virus (CaLCuV) and tobacco mosaic virus (TMV)], which co-opt endogenous pathways to promote movement of the virus, access PD by ‘hitch-hiking’ to the plasma membrane on endocytic vesicles. Other plant viruses and endogenous plant proteins appear to access the PD through associations with the endoplasmic reticulum (ER) and/or the desmotubule (Sambade et al., 2008; Liu et al., 2012).

Some of the viral movement proteins that show associations with the endomembrane also interact with microtubules (Harries et al., 2010). In animal cells, the role for microtubules in promoting viral movement is well established; microtubules promote viral entry and trafficking to the site of replication as well as movement of the virus to the endosome and back to the plasma membrane for viral exit (Brandenburg and Zhuang, 2007). Much of this movement is dependent upon microtubule motor proteins, making it similar to the transport of organelles such as the mitochondria, ER, Golgi apparatus, peroxisomes and secretory vesicles throughout the cell. In contrast, when examined in plants, this type of movement is mediated predominantly by actin-based structures, although there is some evidence for a role of microtubules in the movement and positioning of the Golgi, chloroplasts and P–bodies through interaction with precisely aligned cortical microtubules (Hamada et al., 2012). Examples in which microtubules regulate the trafficking of vesicles or proteins in plants are largely limited to cytokinesis, during which the phragmoplast microtubules direct the deposition of membrane and cell-wall materials to the forming cell plate (Staehelin and Hepler, 1996; Jurgens, 2005), and to the positioning of cellulose synthases during interphase, which track along microtubules in the oriented deposition of cellulose microfibrils (Wasteneys, 2004; Bringmann et al., 2012). Therefore, the primary functions for microtubules in plants appear to be the regulation of cell division and the control of cell morphology.

The compelling evidence for microtubule-dependent protein movement of a plant virus comes from studies of TMV (Harries et al., 2010; Harries and Ding, 2011; Schoelz et al., 2011; Pena and Heinlein, 2012). Like other RNA viruses, the TMV genome encodes an MP that facilitates intercellular spread of the virus. At the start of TMV infection, the TMV–MP is associated with mobile particles on microtubules. Mutations in TMV–MP that decrease its affinity for microtubules and therefore its localization to these mobile particles inhibit the establishment and intercellular spread of the infection (Boyko et al., 2007). It is therefore likely that microtubules promote intercellular movement of the virus early on at the leading edge of the expanding infection. This is not true late in the infection process, where binding of TMV–MP to microtubules inhibits movement of the virus and targets the MP for degradation (Ashby et al., 2006; Boyko et al., 2007). Likewise, binding of TMV to the movement protein binding protein, MPB2C, which increases the localization of TMV–MP to microtubules, also inhibits movement (Kragler et al., 2003; Curin et al., 2007). Similar results have been shown for the maize homeodomain protein Knotted1, which binds to MPB2C (Winter et al., 2007). Therefore, early in the process of infection, microtubules promote cell-to-cell movement of TMV, whereas late in the course of infection, accumulation of TMV–MP on microtubules sequesters the virus within the cell.

Several models have been suggested to explain the positive role that microtubules play in movement of TMV (Harries et al., 2010; Harries and Ding, 2011; Schoelz et al., 2011; Pena and Heinlein, 2012). The first is similar to what has been seen in animals, i.e. that microtubules serve as tracks for microtubule motor protein-dependent movement of TMV to PD. The second is based in part upon the finding that the sequence of the TMV–MP contains motifs that resemble tubulin (Boyko et al., 2000). This model proposes that TMV–MP is able to bind to microtubules and move within the cell via nucleation-dependent movement. Consistent with this model, microtubules are dynamic under conditions of low levels of MP binding, like those at the leading edge of infection, and stabilized by binding of MP along the length of the microtubule filament, as occurs late in the infection process (Brandner et al., 2008). Moreover, mutations or treatments that decrease microtubule polymerization and dynamics result in decreased susceptibility to TMV (Ouko et al., 2010). The third model suggests a less direct role for microtubules in TMV movement. This model suggests that, via interactions with the ER, microtubules provide a platform for viral assembly and release and may promote movement of viral particles along the ER. Support for this model comes from the finding that stable tubular ER junctions are maintained by intact microtubules, mobile ER compartments frequently pause at microtubules (Hamada et al., 2012), and MP complexes move in a ‘stop and go’ manner when in contact with microtubules (Laliberte and Sanfacon, 2010).

A number of observations suggest that viruses have co-opted for their own transport, pathways that evolved to promote the movement of endogenous plant proteins. Therefore, there may be substantial overlap between the mechanisms of viral movement and the movement of proteins encoded by the plant. Here we examine what role microtubules play in the intercellular transport of SHR. Using a variety of techniques, we find that microtubules play a positive role in the movement of SHR. As microtubules have many functions in the cell, including key roles in cell division and determination of cell shape (Staehelin and Hepler, 1996; Frank and Smith, 2002; Smith, 2003; Wasteneys, 2004; Jurgens, 2005), we further dissect SHR movement and show that neither changes in cell shape nor defects in cytokinesis have any effects on the cell-to-cell transport of SHR. Likewise, we find no evidence for polymerization-dependent movement of SHR or a need for regularly aligned cortical microtubules in SHR movement. Instead, in the absence of intact microtubules, we find that localization of the SHR-binding protein SIEL to endosomes was reduced and SIEL accumulated in perinuclear structures. We therefore propose that intact microtubules support the association of SIEL with endosomes and that this association promotes SHR movement. These results provide evidence for the involvement of interphase microtubules in SIEL localization, and therefore the intercellular movement of a developmentally important transcription factor in root growth.

Results

Disruption of microtubules inhibits movement of SHR into the endodermis

SHR is synthesized in stele cells, where it localizes to both the nucleus and the cytoplasm (Figure 1a). The SHR protein then moves into the endodermis where it is confined to the nucleus (Cui et al., 2007; Gallagher and Benfey, 2009; Gallagher et al., 2004; Nakajima et al., 2001). As the stele is the sole source of the SHR protein in the endodermis, the endodermis to stele (E:S) ratio of the SHR–GFP signal may be used as a marker for SHR movement in live imaging experiments (Figure S1A). In 5-day-old wild-type roots grown on standard Murashige & Skoog (MS) growth medium, the mean E:S ratio for SHR is 1.2 (Figure 1i, bar a). In contrast, in the siel-4 mutant background, where SHR movement is reduced, the mean E:S ratio in normally patterned roots was 0.96 (Koizumi et al., 2011). To determine whether microtubules are involved in the intercellular trafficking of SHR, we tested the effects of the microtubule-depolymerizing drug oryzalin (Morejohn et al., 1987) on the E:S ratio of SHR–GFP fluorescence. For these experiments, we tested a range of oryzalin concentrations from 0.3 to 3.0 μM (Figure 1b–d) that have been shown to disrupt microtubules (Baskin et al., 1994; Sugimoto et al., 2003; Bannigan et al., 2006). We found that treatment of SHR–GFP plants with 0.3 μM oryzalin (the lowest concentration tested) for 12 h reduced the E:S ratio to 0.5, significantly below that of the controls (P = 3.87E-06; Student's t test; Figure 1b,i). Further reductions in the SHR–GFP signal were not seen with higher concentrations of oryzalin, indicating that treatment with 0.3 μM oryzalin for 12 h was sufficient to disrupt microtubules. Likewise, to ensure that the concentration of oryzalin used in these assays disrupts microtubules, we imaged seedlings expressing mEosFP-tagged MICROTUBULE ASSOCIATED PROTEIN 4 (mEosFP–MAP4; Mathur et al. 2010) after 12 h treatment with 0.3 μM oryzalin (Figure S2A–C).

Figure 1.

Oryzalin inhibits SHR movement. (a–d) Confocal micrographs of wild-type Col roots expressing SHR–GFP treated with the indicated concentrations of oryzalin. (a) Wild-type root expressing SHR–GFP. SHR moves from the stele tissue (outlined with a dotted white line) into the neighboring endodermis (E). (b–d) Results after 12 h treatment with various concentrations of oryzalin. (e–h) Wild-type transgenic roots expressing both SHR–GFP and the pWol:icalsm transgene. Roots in (f)–(h) were treated with estradiol for 8 h and then allowed to recover for 6 h on estradiol-free medium either without (f) any inhibitor or (g) in the presence of oryzalin, or (h) in the presence of CD. E, endodermis. (i) Quantification of the E:S ratio of SHR:GFP. The letters on the bars correspond to the confocal micrographs above, which are representative images. Values are means ± SD of at least eight plants for each indicated treatment (see Figure S1B–D for more information).

As long-term incubation with oryzalin may significantly affect root growth (Baskin et al., 1994; Sugimoto et al., 2003), we examined the E:S ratio in roots treated for 6 h with the intermediate concentration of oryzalin tested (1.0 μM). The 6 h timeframe was chosen as previous experiments indicated that turnover of SHR–GFP takes approximately 6 h in the endodermis (Vaten et al., 2011). Consistent with the results of the 12 h treatment, incubation of roots expressing SHR–GFP with 1.0 μM oryzalin for 6 h reduced the E:S ratio to 0.65 (P = 0.00017; Student's t test relative to untreated controls), similar to the levels seen with the other treatments (Figure 1i), suggesting that microtubules are involved in SHR movement. To ensure that disruption of microtubules did not generally inhibit movement via PD, we examined the effects of oryzalin on movement of unmodified (‘free’) GFP driven by the phloem-specific SUC2 promoter. GFP made in the phloem is free to diffuse via PD throughout the root tip (Figure S2E,F) (Stadler et al., 2005). Treatment of the pSUC2-GFP roots with 1.0 μM oryzalin for 12 h revealed no change in the distribution of fluorescence, indicating that passive diffusion is not affected by depolymerization of microtubules (Figure S2E,F).

To further analyze the effects of oryzalin on SHR movement, we took advantage of two additional, independent methods for assaying SHR movement that allow us to decrease SHR levels in the endodermis and then examine recovery under short-term treatment with or without oryzalin. As these assays start by reducing SHR levels in the endodermis, they reduce some of the confounding effects of protein perdurance of SHR in the endodermis on measurements of movement, and may therefore provide more accurate results. The first assay takes advantage of a semi-dominant, synthetic allele of the gene encoding CALLOSE SYNTHASE 3 (icalsm) that is inducibly expressed in the stele from the stele-specific WOODENLEG promoter (pWOL, also known as the CRE1 promoter) (Vaten et al., 2011). Activation of the transgenic pWOL:icalsm system by application of estradiol results in the transient accumulation of callose at PD and a decrease in the PD aperture (Figure S1B,C) (Vaten et al., 2011). In response to 8 h of treatment with estradiol, there was a dramatic decrease in SHR–GFP signal in the endodermis of pWOL:icalsm-expressing plants (Figure 1e). However, this block was reversible. Upon removal from estradiol, SHR–GFP was again able to move from the stele into the endodermis (Figure 1f and Figure S1D). Using this system, we blocked movement of SHR–GFP and then monitored its recovery in the absence or presence of inhibitor. In the absence of oryzalin, the E:S ratio of SHR–GFP increased from 0.48 to 1.02 after 6 h on estradiol-free medium (Figure 1f,i); this represents a 71% recovery in movement. In contrast, when roots were removed from estradiol treatment and allowed to recover on medium that contained 1.0 μM oryzalin, we observed only a 14% recovery (from an E:S ratio of 0.48 to 0.59) in the same timeframe (Figure 1g,i); this represents 20% of that observed in untreated roots (P = 1.25E-05; Student's t test), suggesting that a loss of intact microtubules significantly reduces SHR movement.

The second method that we used to examine movement of SHR–GFP in the presence of oryzalin was fluorescent recovery after photobleaching (FRAP) (Figure S1E–G). In assays to examine the recovery of SHR movement, the SHR–GFP signal was sequentially bleached in the endodermis using a 488 nm laser. Recovery was then monitored over 120 min, during which time images were captured every 40 min to reduce the possibility of additional bleaching (Figure 2). As SHR is not directly expressed in the endodermis, any recovery in fluorescent signal in the endodermis represents movement of the SHR–GFP protein from the stele. In the absence of oryzalin, we observed a 60% recovery in the level of SHR–GFP in the endodermis (Figure 2b) 120 min after photobleaching. In the presence of oryzalin, there was a 22% recovery in SHR–GFP, significantly lower than the untreated controls (P = 1.01E-08; Student's t test; Figure 2). Taken together, the results of both assays (Figure S2B–E) indicate that oryzalin causes a 65–80% reduction in overall SHR movement into the endodermis.

Figure 2.

Oryzalin treatment significantly inhibits FRAP of SHR. (a) SHR–GFP was photobleached, and fluorescence recovery was monitored for 120 min with or without inhibitors (as indicated). (b) Fluorescence recovery in the absence (closed squares) of either drug, the presence of oryzalin (closed triangles) or the presence of CD (open circles). The numbers on the y axis represent percentage recovery, and are × 100. Percentage recovery was calculated as (FA – FB)/(FP – FB).

As disruption of microtubules may cause defects in the organization or assembly of actin filaments (Collings et al., 2006; Collings, 2008), we examined the effects that treatment of roots with the actin-depolymerizing agents cytochalasin D (CD) or latrunculin B (LatB) had on the movement of SHR from the stele into the endodermis (Figure S2G–I). In contrast to the oryzalin treatments, the effects of 20 μM CD and 50 nM LatB treatment (for 24 h) on SHR movement were relatively mild, with a 10% and 13% decrease, respectively, in the E:S ratios (= 0.06 and 0.03, respectively; Student's t test relative to untreated controls). Likewise, when estradiol-treated seedlings expressing the pWOL:icalsm transgene were recovered on medium containing 20.0 μM CD, recovery was approximately 75% of that observed in untreated control roots (Figure 1h,i); this is almost four times what was observed in the oryzalin-treated roots (Figure 1g,i). Similarly, treatment of roots with oryzalin had a more dramatic effect on FRAP of SHR than treatment with CD (Figure 2). Nonetheless, the difference in recovery of SHR-GFP fluorescence between the control roots and the CD-treated roots was significant (= 0.000515; Student's t test relative to untreated controls), indicating that actin may play a role in promoting SHR transport.

As oryzalin has a much greater effect on SHR movement than either LatB or CD, these results show that the effect of oryzalin on SHR movement is not via actin. Control experiments were performed to examine root elongation after treatment with 20 μM CD or 50 nM LatB. After 24 h on either 20 μM CD or 50 nM LatB, the rate of root elongation was reduced by over 70% (Figure S2J), suggesting that, at the concentrations used in these assays, both CD and LatB are effective in inhibiting actin function (Rahman et al., 2007).

To further examine the role that microtubules play in SHR movement, SHR–GFP was analyzed in two conditional mutants, microtubule organization 1 (mor1) (Kawamura et al., 2006; Whittington et al., 2001; Lechner et al., 2012) and radially swollen 7 (rsw7) (Figure 3), which fail to maintain intact microtubules under restrictive temperatures (Bannigan et al., 2007). MOR1 is an Arabidopsis homolog of the highly conserved microtubule-associated protein MAP215 (Kawamura and Wasteneys, 2008). MAP215 homologs are important for the formation and maintenance of intact microtubule arrays. RSW7 encodes AtKRP125c, a kinesin-5 motor protein that is required for microtubule organization (Bannigan et al., 2007). Microtubules in both mor1-1 and rsw7 are relatively normal at the permissive temperature of 19°C and disrupted at 30°C (Kawamura et al., 2006; Kawamura and Wasteneys, 2008; Whittington et al., 2001; Bannigan et al., 2007; Winter et al., 2007). The E:S ratios of SHR–GFP in the mor1-1 and rsw7 mutants at 19°C were approximately 1.2 (Figure 3a,b,e), which is not significantly different than the wild-type controls in these experiments (Figure S3A,C). In contrast, incubation at 30°C for 12 h resulted in a 30% and 43% decrease in the E:S ratios of mor1-1 and rsw7, respectively (P = 3.25E-09 and 2.37E-08, respectively; Student's t test) with no effect on wild-type (Figure 3c–e and Figure S3). To verify these results, FRAP of SHR–GFP was assessed in the mor1-1 and rsw7 mutants (Figure 3f). We observed a decrease in the recovery of SHR–GFP at the restrictive temperature that was similar to the effects of oryzalin on SHR movement (Figure 2b and Figure 3f). These data suggest a microtubule-dependent pathway for intercellular movement of SHR.

Figure 3.

Intact microtubules promote SHR movement. (a–d) mor1-1 and rsw7 grown at 19°C prior to transfer to 30°C (as indicated) for 8 h. E, endodermis. (e) Quantification of the E:S ratio under the conditions indicated. The letters on the bars correspond to the confocal micrographs on the left, which are representative images. Values are means ± SD of at least eight plants for each indicated treatment. (f) FRAP assays performed as described in Figure S1 at the non-permissive temperature. The numbers on the y axis represent percentage recovery, and are × 100.

As microtubules are required for both mitosis and cytokinesis, disruption of microtubules results in a reduced rate of cell division and root growth (Staehelin and Hepler, 1996; Jurgens, 2005). To determine whether a reduced rate of root growth or incomplete cytokinesis directly affects SHR movement, we treated roots with the DNA synthesis inhibitor hydroxyurea (HU). HU treatment slows down progression through the cell cycle by inhibiting the ribonucleotide reductase enzyme and slowing dNTP production (Collins and Oates, 1987). HU does not inhibit microtubules (Balczon et al., 1999). When roots were treated with 3.0 mM HU for 12 h, there was a slight reduction in cell elongation, and the rate of cell production dropped to half that of the control (Figure S4). However, we saw no change in the SHR–GFP E:S ratio relative to the controls (Figure 4a,d), indicating that a reduced rate of growth does not inhibit SHR movement. Likewise, even at the permissive temperature, rsw7 roots had a reduced rate of cell production (approximately 50% of wild-type; Figure S4) and cell division defects that included cell-wall stubs and incomplete cytokinesis; however, movement of SHR–GFP was relatively normal at 19°C (Figure 3b,e).

Figure 4.

SHR movement persists in the presence of abnormal cell divisions, stabilized and mis-oriented microtubules. (a) SHR–GFP movement in the presence of HU. (b) SHR–GFP movement recovers after 6 h treatment with 1.0 μM oryzalin followed by incubation for 3 h on medium lacking oryzalin. (c,e) Recovery of SHR–GFP movement in rsw7 and mor1-1 after transfer from 30°C (8 h) to 19°C for 4 h. (d) E:S ratios after recovery for (a–c,e). The letters on the bars correspond to the confocal micrographs. (f,g) Taxol at the concentrations indicated had no effect on SHR–GFP movement. (h) E:S ratios after recovery for (f,g). The letters on the bars correspond to the confocal micrographs. (i–k) Movement of SHR–GFP is relatively normal in (i) lefty-2 and (j,k) rsw6 mutants. (l) Quantification of the E:S ratios of SHR–GFP in lefty-2 and rsw6 mutants. The letters on the bars correspond to the confocal micrographs.

The strongest argument against defective cell division or cell morphology playing a primary role in the inhibition of SHR movement is the ability of SHR movement to recover after oryzalin treatment or inhibition of RSW7 or MOR1 (Figure 4b–e), which all have significant effects on cell shape and cytokinesis. When roots were treated with 1.0 μM oryzalin for 6 h followed by a 3 h recovery on oryzalin-free medium, the E:S ratio of SHR–GFP returned to approximately 85% of pre-treatment levels (Figure 4b,d). Similarly, after 8 h at restrictive temperature followed by 4 h at 19°C, SHR–GFP movement recovered to over 80% of the pre-treatment levels in rsw7 and mor1-1 (Figure 4c–e). Therefore, reorganization of microtubules may promote intercellular transport of SHR even in the presence of defective cell divisions and abnormally shaped cells. Therefore, defective cell division and cell morphology are not the major factors contributing to a decrease in the intercellular movement of SHR.

Neither dynamic nor regularly aligned microtubules are essential for SHR movement

One of the models for how TMV–MP moves is through intercalation with polymerizing microtubules (nucleation-dependent movement; Ouko et al., 2010). To explore how microtubules mediate SHR trafficking, we tested whether dynamic microtubules (those that are capable of polymerization and depolymerization) are important for SHR movement. To test this, we treated SHR–GFP-expressing roots with taxol, which stabilizes polymerized microtubules (Baskin et al., 1994). As shown in Figure S5(A–C), when treated for 24 h with 1.0 μM taxol, fluorescence recovery of mEosFP–MAP4 after 80 sec photobleaching is only half (51%) that of the control, indicating that 1.0 μM taxol reduces microtubule dynamics. To analyze the effects of taxol on SHR movement, two treatments were examined: 1.0 μM taxol for 12 h and 10.0 μM taxol for 6 h (Figure 4f–h). Neither treatment significantly affected movement of SHR–GFP, indicating that dynamic microtubules are not required for SHR movement. This suggests that nucleation-dependent movement, which requires dynamic microtubules, is not required for SHR movement.

Another possibility is that SHR moves intracellularly along regularly aligned stable microtubules in association with a microtubule-associated motor protein. As cortical microtubules in the Arabidopsis root are arranged in parallel arrays that are perpendicular to the axis of growth and coordinated between neighboring cells (Baskin et al., 1994; Overall et al., 2001), they may provide a route for trafficking radially along the plasma membrane to the PD. To test this, we examined SHR movement in mutants that fail to maintain regularly aligned microtubules. In contrast to mutants such as mor1-1 and rsw7, which exhibit a general disruption of microtubules at the non-permissive temperature (Kawamura et al., 2006; Kawamura and Wasteneys, 2008; Whittington et al., 2001; Bannigan et al., 2007; Lechner et al., 2012), mutants such as lefty-2 and radially swollen 6 (rsw6) have intact microtubules that are inconsistently organized between cells and generally mis-aligned (Bannigan et al., 2006; Thitamadee et al., 2002). The lefty-2 allele is not a temperature-sensitive allele, whereas rsw6 shows the mutant phenotype preferentially at 30°C. To test whether a loss in the coordinated patterning of microtubules affects the intercellular movement of SHR, we examined SHR–GFP ratios in both lefty-2 and rsw6 roots. lefty-2 mutants have a mutation in α–tubulin 4 that results in formation of right-handed obliquely oriented cortical arrays of microtubules that cause a left-handed spiral growth habit in the root (Thitamadee et al., 2002). The E:S ratio of SHR–GFP was slightly reduced in the lefty-2 mutants compared to the control, but was not outside of the range observed in wild-type roots (Figure 4i,l). Similar results were seen for rsw6. At 19°C, rsw6 mutants have intact microtubules that are organized into uniform coordinated transverse arrays that are consistent between cells. At 30°C, this organization is lost (Bannigan et al., 2006). However, after 12 h at 30°C, we saw no defects in SHR movement in the rsw6 roots (Figure 4j–l). These results suggest that microtubules need not be perpendicular to the axis of root growth or coordinated between cells in order to promote movement of SHR.

Intact microtubules promote the localization of SIEL to endosomes

If neither dynamic nor regularly aligned microtubules are required for SHR movement, then how do microtubules participate in cell-to-cell transport? The only factor known to promote SHR movement is SIEL. We identified SIEL as part of a yeast two-hybrid screen in which SHR was used as the bait. SIEL is an endosome-associated protein that promotes movement of SHR from the stele into the endodermis. Null alleles of siel are embryo-lethal; whereas hypomorphs have reduced movement of SHR and cause defects in root patterning (Koizumi et al., 2011). The reduction in SHR movement in siel is similar in degree to the reduction caused by microtubule disruption. In experiments shown here, in which SHR movement is examined in the siel-4 mutant background without regard to root morphology, movement was approximately 74% of wildtype. Treatment of siel-4 roots with oryzalin further reduced SHR movement by an additional 21% (to approximately 53% of the untreated wild-type controls), but not below what is seen in wild-type roots treated with oryzalin (P = 0.38, Student's t test; Figure S6A–E), indicating a lack of additive effects. As siel-4 is a weak hypomorph, these results suggest that oryzalin-induced inhibition of SHR movement acts via SIEL.

To determine how microtubule-dependent trafficking of SHR functions through the SIEL pathway, we examined SIEL localization in taxol-, CD-, HU- and oryzalin-treated roots. In the control roots, YFP–SIEL is distributed throughout the cell, with a relatively high proportion of signal close to the cell cortex (Figure 5a) and in association with endosome markers, particularly markers of the recycling endosomes (over 60% of punctate YFP–SIEL co-localized with markers of the endosome; Koizumi et al., 2011). The localization of YFP–SIEL was not noticeably altered by application of taxol, CD or HU (Figure S7A–C). However, treatment of YFP–SIEL plants with 1.0 μM oryzalin for 8 h resulted in a redistribution of YFP–SIEL away from endosomes (Figure 5c–h) and the cell cortex to punctate structures along the periphery of the nucleus (Figure 5b). Similar localization was seen in the rsw7 and mor1-1 mutants (Figure S7D,E). Consistent with the recovery of SHR movement upon withdrawal from oryzalin or the return of rsw7 and mor1-1 to the permissive temperature, YFP–SIEL redistributes to the cell cortex after 3 h recovery from 1.0 μM oryzalin treatment (10 h) or return of rsw7 and mor1-1 to 19°C (Figure S7F–H).

Figure 5.

Oryzalin affects localization of YFP–SIEL. (a,b) YFP–SIEL localization in wild-type roots without oryzalin treatment (a) and roots treated with oryzalin (b). Oryzalin causes YFP–SIEL to localize around the nucleus (arrows). (c–e) Co-localization of YFP–SIEL with the endosome/recycling endosome marker RabA5d. Arrows in (e) show areas of YFP–SIEL and RabA5d–mCherry overlap. (f–h) Co-localization of YFP–SIEL with ARA7–mCherry. Arrows in (h) show regions of overlap between the YFP and mCherry signals.

The perinuclear localization of SIEL in the presence of oryzalin prompted us to study the localization of endosome markers, particularly components of the recycling endosome, as they are often described as showing perinuclear localization (Guilherme et al., 2000; Hickson et al., 2003; Ullrich et al., 1996). To do this, we used the so-called ‘Wave’ markers lines that reliably mark various subcellular compartments (Geldner et al., 2009). Individual Wave marker lines that are annotated as showing associations with the endosome/recycling endosome (at least in part) were examined before and after 8 h treatment with 1.0 μM oryzalin, or with HU or taxol (used as controls). There were no obvious changes in their cellular distribution or relocalization to a perinuclear region in the cell for any of the endosome markers (Figure S8). These results suggest that the perinuclear localization of SIEL does not represent a redistribution of the endosome to a region immediately outside of the nucleus. However, we cannot rule out the possibility that our assays missed an endosomal compartment that is not represented in the Wave collection of markers (Geldner et al., 2009). Collectively, our results show that intact microtubules promote localization of SIEL to the endosome and the cell cortex, and therefore suggest a mechanism by which microtubules promote SHR movement.

Discussion

The intercellular movement of transcription factors is prevalent in plants. It has been estimated that more than 15% of root-expressed transcription factors move from the cells in which they are synthesized (Lee et al., 2006). Likewise, a recent report by Rim et al. (2011) suggests that the capacity for cell-to-cell trafficking is a common feature of multiple families of transcription factors. Despite this widespread movement, the mechanisms that regulate intercellular transport are largely unknown. Here we show that movement of the SHR protein from the stele to the endodermis is moderately decreased by disruption of actin and is significantly inhibited by short-term treatment with oryzalin or mutations in MAP215 (MOR1) or AtKRP125c (RSW7). As oryzalin and the mor1-1 and rsw7 mutations all disrupt microtubules, these results indicate that intact microtubules promote intercellular transport of SHR. However, as neither taxol treatment nor the rsw6 and lefty mutations significantly disrupted movement of SHR, it is unlikely that microtubules serve as tracks upon which SHR moves to access the neighboring cell or the PD. As PD largely lack microtubules, it is also unlikely that microtubules promote SHR movement through the PD. Instead, the reduction in SHR movement in roots that lack intact microtubules is probably the result of mis-localization of the SIEL protein.

The only protein that has been shown to promote SHR movement is SIEL. To determine whether microtubules and SIEL function in the same pathway to promote SHR movement, we treated siel mutant roots with oryzalin. Ideally this experiment would have been performed on siel null mutants, with the expectation being that oryzalin will result in no further enhancement of the null phenotype if SIEL and microtubules act together to promote SHR movement. However, loss of SIEL function is lethal, and strong hypomorphic alleles produce a very short and disorganized root in which quantitative analysis of SHR movement is impossible. We therefore treated siel-4 roots (weak hypomorphs) with oryzalin. If SIEL and microtubules act in different pathways, we anticipate either additive or synergistic effects on SHR movement. However, if the reduction in SHR movement in the oryzalin-treated roots was the result of mis-localization of SIEL, we would expect the effects of oryzalin to be essentially epistatic to siel-4. Indeed, this is what we observed. The reduction in SHR movement in the oryzalin-treated roots was not significantly different to the reduction in SHR movement in the wild-type roots treated with oryzalin. These results are consistent with microtubules and SIEL acting in the same pathway.

Based upon the localization of TMV–MP and KN1 and their associations with MPB2C and microtubules, both positive and negative roles have been postulated for microtubules in the movement of viruses and endogenous plant proteins between cells (Kragler et al., 2003; Curin et al., 2007; Winter et al., 2007; Ruggenthaler et al., 2009; Harries et al., 2010; Harries and Ding, 2011). Consistent with a positive role for microtubules in the spread of TMV, mutations in the TMV–MP that abolish microtubule localization inhibit viral spread, and plants with reduced microtubule turnover have increased viral resistance (Boyko et al., 2007; Ouko et al., 2010). At least three models have been proposed to explain the TMV–MP results. The first is that association of TMV–MP with molecular motors allows movement along microtubule tracks, facilitating the localization of TMV–MP to the PD. The second is that TMV–MP relies upon microtubule polymerization to move, essentially through interaction with tubulin and incorporation into microtubules (Ouko et al., 2010). However, most data support a third model, which suggests that microtubules play an indirect role in the movement of TMV by providing a docking site for viral assembly and through associations with the ER. Sambade and Heinlein proposed that association of TMV–MP with microtubules in the cellular cortex promotes the formation of focused ribonucleoprotein complexes, which become associated with the ER and undergo actin-dependent movement to the PD (2009). In support of this hypothesis is the finding that, when microtubules are disrupted or the binding of TMV–MP to microtubules is inhibited, TMV–MP is still localized to PD (Boyko et al., 2007; Ouko et al., 2010). Therefore, microtubules are not required to access the PD. Instead it is more likely that microtubules provide a platform for the formation of viral replication complexes.

In the Arabidopsis root, interphase microtubules are present predominantly in the cell cortex, where they are arranged into regular transverse arrays. In the stele tissue, the cortical microtubules are less regularly patterned, and there is less consistency in the overall organization between cells. In addition, in the cells in the root mersitem that have not undergone vacuolation, microtubules are found in association with the nucleus. These microtubule arrays are distinct from those that form the pre-prophase bands or the spindle in that they are less densely associated with the nucleus and they extend well into the cell cortex (Baskin et al., 1992). Therefore, in theory, microtubules may support movement of proteins such as SHR from the nucleus to the cell cortex, or along the plasma membrane to the PD; they may also serve as platforms for protein–protein interactions in either of these two domains.

Previously we showed that SHR localizes to both the nucleus and cytoplasm, and that localization to both of these compartments is required for cell-to-cell movement (Gallagher et al., 2004; Gallagher and Benfey, 2009). Likewise, we showed that, in root cells, the SHR-interacting protein SIEL is found both in the nucleus and the cytoplasm, where it associates with endosomes and promotes intercellular movement of SHR into the endodermis (Koizumi et al., 2011). In the presence of oryzalin, there was a decrease in the endosomal localization of SIEL and a reduction in SHR movement. As SHR and SIEL both show cytoplasmic and nuclear localization, cytoplasmic microtubules extending from the nucleus may provide a pathway for localization of SIEL to the endosome. However, as the orientation of microtubules appears unimportant for SHR movement, it seems more likely that microtubules strengthen SIEL's ability to associate with endosomes and/or that they serve as a platform for interaction between SHR and SIEL, similar to the model proposed for the ER and microtubules in viral movement (Sambade and Heinlein, 2009; Sambade et al., 2008). We suggest that, once localized to the endosome, SHR and SIEL may traffic within the cell as an endosome-associated complex. As PD are regions of increased membrane recycling, this may bias movement of SHR to the PD (Oparka, 2004).

In plants, most intracellular movement of vesicles and organelles is thought to occur through associations with actin. Roles have been suggested for microtubules in movement of the Golgi and for localization of components of the cellulose synthase complex to the plasma membrane (Crowell et al., 2009). In fungi, the microtubule cytoskeleton has been shown to affect movement of endosomes, including the recycling endosome, in animals (Apodaca, 2001; Lapierre et al., 2012). In fact, most long-range movement of vesicles in animal cells is thought to occur through associations with polarized microtubules (Hirokawa et al., 2009), whereas localized movement in the cortex is mediated by actin. Very interesting results were obtained in Ustilago maydis, in which microtubule motors shuttle large ribonucleoprotein complexes in association with endosomes. Both intact microtubules and endosomes are required for this movement (Baumann et al., 2012). We examined localization of endosome-associated markers fused to mCherry or YFP. There was no obvious change in the cellular distribution or localization of any of these markers in oryzalin-treated versus untreated roots, suggesting that disruption of microtubules does not cause a change in the cellular distribution of endosomes. However, we cannot rule out the possibility that particular classes of endosomes that are not represented by the Wave set of markers show microtubule-dependent localization.

In plants, it is not known how soluble cytoplasmic proteins become associated with vesicles. Nor is it known what contributions microtubules make to protein targeting. There are many proteins in animals that localize to either endosomes (e.g. the sorting nexins, SXNs) or secretory vesicles (e.g. SEC5, SEC6, SEC8, SEC10 and SEC15). The SXNs bind directly to phosphatidylinositol 3–phosphate in the endosomal membrane (Ullrich et al., 1996). In contrast, components of the exocyst complex tend to bind to activated Rab proteins (He and Guo, 2009). For neither the SXNs or the SECs is it known how these proteins are transported to their targets. One possibility is that they simply diffuse in the cytoplasm until they contact their binding partner. However, another is that they are actively transported via either the actin or microtubule cytoskeleton. Similar suggestions have been made for the movement of proteins to the PD, i.e. either diffusion in the cytoplasm or specific targeting. Recently, it was shown that a protein involved in chloroplast function, SNOWY COTYLEDON 3 (SCO3) requires intact microtubules for localization to peroxisomes, and that this localization is required for SCO3 function (Albrecht et al., 2010).

We propose a mechanism for SHR movement in which microtubules promote the localization of SIEL to endosomes. The finding that SIEL accumulates around the nucleus in the absence of intact microtubules suggests that microtubules promote localization of SIEL to the endosome and the cell cortex. Together with the previous finding that both nuclear and cytoplasmic localization of SHR are required for its intercellular transport (Gallagher et al., 2004; Gallagher and Benfey, 2009), we propose a model for SHR movement in which, upon nuclear export, some proportion of the SHR pool of protein becomes bound to SIEL, which itself associates with the endosome; SHR then undergoes SIEL-associated movement to the cell cortex where the SHR protein is transported out of the cell. As SIEL has been shown to interact with several other mobile proteins, this may be a general mechanism for protein transport.

Experimental procedures

Plant material and growth conditions

Arabidopsis thaliana Col–0 was used as the wild-type. Plants were germinated and grown vertically on 1.0 x Murashige & Skoog (MS) medium (Caisson, www.caissonlabs.com) containing 0.05% w/v MES (pH 5.7), 1.0% w/v sucrose and 1.0% granulated agar (DIFCO,www.bd.com) in a growth chamber at 19°C under a 16 h light/8 h dark cycle. The 30°C treatments were performed in the same chamber except the temperature setting was increased to 30°C. Plants were used 6 days after plating unless otherwise stated. The pSHR:SHR:GFP line was crossed into the mor1-1, rsw7, rsw6 and lefty-2 backgrounds. Homozygous mutants expressing the SHR–GFP reporter were selected based upon fluorescence and the root phenotype at 30°C. The SUC2–GFP lines have been described by Stadler et al. (2005).

Treatment with inhibitors

Stock solutions of 20 mM oryzalin (Sigma, www.sigmaaldrich.com) 10 mM cytochalasin D (Sigma), 1.0 mM Latrunculin B (Sigma) and 10 mM taxol (Acros Organics, www.acros.be) all in dimethylsulfoxide and 3.0 M hydroxyurea (Sigma) in ddH2O were prepared and stored at −20°C. Plates containing the specified inhibitors were prepared by adding the appropriate stock solution to autoclaved and cooled MS agar. Controls received the same amount of water or dimethylsulfoxide. For callose induction, 1.0 μM 17–β–estradiol (Sigma) was used as described previously (Vaten et al., 2011). For all of the recovery studies, the seedlings were gently washed briefly (less than 1 min at room temperature) in liquid MS medium (as above without agar) and then transferred to fresh MS agar plates for growth under the same conditions.

Microscopy and imaging

Root elongation rates were measured by marking the tip of the root on the plate once per day and then photographing the plate. The images were analyzed by ImageJ software (rsbweb.nih.gov/ij/) to calculate the root length between two markers on successive days. The length of the cortex cells were measured from confocal micrographs of medial longitudinal sections through roots stained with propidium iodide (0.01 μg/ml in water). The cell flux was calculated as described by Baskin and Wilson (1997). Confocal images were captured on a Leica (www.leica.com) TCS SL microscope with the appropriate filter sets for visualizing GFP and propidium iodide. The measurement of SHR–GFP intensity, the calculation of the E:S ratio, FRAP of SHR–GFP and co-localization between YFP–SIEL and the endosome markers were performed as described by Koizumi et al. (2011). FRAP of mEosFP–MAP4 was performed using a Zeiss (www.zeiss.com) LSM 710 laser scanning confocal microscope using a Zeiss LD C–Apochromat 40 ×/1.1 NA water immersion objective. Photobleaching was achieved using six iterations of the 488 nm laser at 100% power. Recovery was followed by image acquisition using 10% laser power at 10 sec intervals.

Acknowledgements

We thank T. Baskin for seeds and comments, D. Wagner for comments on the manuscript, and D. Wagner, R.S. Poethig, B.D. Gregory and members of their laboratories for technical assistance. S.W. is supported by US National Science Foundation grant 0920327 awarded to K.L.G.

Ancillary