Research ArticleResearch Article

CORTICAL MICROTUBULE DISORDERING1 Is Required for Secondary Cell Wall Patterning in Xylem Vessels

Takema Sasaki, Hiroo Fukuda, Yoshihisa Oda

Published December 2017. DOI: https://doi.org/10.1105/tpc.17.00663

Abstract

Proper patterning of the cell wall is essential for plant cell development. Cortical microtubule arrays direct the deposition patterns of cell walls at the plasma membrane. However, the precise mechanism underlying cortical microtubule organization is not well understood. Here, we show that a microtubule-associated protein, CORD1 (CORTICAL MICROTUBULE DISORDERING1), is required for the pitted secondary cell wall pattern of metaxylem vessels in Arabidopsis thaliana. Loss of CORD1 and its paralog, CORD2, led to the formation of irregular secondary cell walls with small pits in metaxylem vessels, while overexpressing CORD1 led to the formation of abnormally enlarged secondary cell wall pits. Ectopic expression of CORD1 disturbed the parallel cortical microtubule array by promoting the detachment of microtubules from the plasma membrane. A reconstructive approach revealed that CORD1-induced disorganization of cortical microtubules impairs the boundaries of plasma membrane domains of active ROP11 GTPase, which govern pit formation. Our data suggest that CORD1 promotes cortical microtubule disorganization to regulate secondary cell wall pit formation. The Arabidopsis genome has six CORD1 paralogs that are expressed in various tissues during plant development, suggesting they are important for regulating cortical microtubules during plant development.

INTRODUCTION

The cell wall is the structural determinant of plant cell morphology. Cellulose microfibrils, the main components of the plant cell wall, physically restrict cell expansion due to their physical strength, causing anisotropic cell growth according to the alignment of cellulose microfibrils. Cellulose microfibers are synthesized at the outer surface of the plasma membrane by the plasma membrane-embedded cellulose synthase (CESA) complex, while other cell wall components such as hemicellulose, pectin, and lignin are synthesized inside the cell and are secreted outside of the cell to be incorporated into the cellulose microfibril matrix. The orientation of the cellulose microfibril is directed by cortical microtubules, which recruit CESA-containing vesicles and guide the trajectory of CESA complexes at the plasma membrane (Paredez et al., 2006; Crowell et al., 2009; Gutierrez et al., 2009). Therefore, the patterning of the cortical microtubule array primarily determines the overall deposition patterns of cellulose microfibrils, which in turn determine plant cell shape.

In most plant tissues, transverse cortical microtubules, which are predominantly aligned perpendicular to the growth axis of the cell, promote anisotropic cell growth, leading to the development of bipolar cylinder-like cells. Live-cell imaging of cortical microtubules revealed the behaviors of cortical microtubules, including treadmilling, branching, severing, and bundling, enabling the cortical microtubules to self-organize through their interactions (Wasteneys and Ambrose, 2009). Microtubule-associated proteins play central roles in regulating the dynamics and interactions of cortical microtubules. Many conserved and plant-specific microtubule-associated proteins help regulate the behaviors of transverse cortical microtubules. MICROTUBULE ORGANIZATION1 (Whittington et al., 2001), KATANIN1 (Burk and Ye, 2002), CLIP-ASSOCIATED PROTEIN (Ambrose and Wasteneys, 2008; Ambrose et al., 2011), and gamma-tubulin complex proteins (Nakamura et al., 2012; Walia et al., 2014), which are conserved in eukaryotes, participate in microtubule dynamics, the severing of microtubules, and microtubule nucleation, all of which are required to maintain the proper arrangement of transverse cortical microtubules. Plant-specific proteins such as ROP-INTERACTIVE CRIB MOTIF-CONTAINING PROTEIN1 (Fu et al., 2009) and SP1-LIKE2 (Shoji et al., 2004; Wightman et al., 2013) also participate in the arrangement of transverse cortical microtubules. Considering the distinct structures and functions of plant cortical microtubules, more plant-specific proteins are likely involved in regulating cortical microtubule organization as well.

In recent years, more complicated behaviors of cortical microtubules during cell differentiation, photosignaling, and hormonal responses have been reported. In pavement cells, cortical microtubules accumulate locally, leading to the development of periodic indentations (Fu et al., 2005; Lin et al., 2013). In the hypocotyl, upon perception of blue light, transverse cortical microtubules are rearranged into longitudinal arrays through the microtubule severing-based amplification of longitudinal microtubules (Lindeboom et al., 2013). Gibberellin and auxin treatment also induces the longitudinal arrangement of cortical microtubules (Vineyard et al., 2013). The molecular mechanisms underlying such rearrangements of cortical microtubules are still not fully understood, and it is reasonable to assume that previously uncharacterized microtubule-associated proteins are also involved in cortical microtubule rearrangement during cell development.

Distinct deposition patterns of secondary cell walls in xylem vessels, such as spiral, reticulate, and pitted patterns, are also governed by cortical microtubule alignment. During xylem vessel cell differentiation, transverse cortical microtubules are gradually rearranged into bundled or pitted patterns to direct the corresponding secondary cell wall patterns (Oda et al., 2005). Increasing evidence suggests that plant-specific microtubule-associated proteins are involved in arranging cortical microtubules in xylem vessel cells. MICROTUBULE-ASSOCIATED PROTEIN70-5 (MAP70-5), encoding a member of a plant-specific microtubule-associated protein family, is believed to control the type of secondary cell wall patterns (Pesquet et al., 2010). MAP65, AUXIN-INDUCED IN ROOT CULTURES9, and CELLULOSE SYNTHASE-INTERACTIVE PROTEIN1 are also thought to be involved in regulating the deposition patterns of secondary cell walls in cultured xylem vessel cells (Mao et al., 2006; Derbyshire et al., 2015).

We previously demonstrated that secondary cell wall pits in metaxylem vessels form through the activity of MICROTUBULE DEPLETION DOMAIN1 (MIDD1), a plant-specific microtubule binding protein. In metaxylem vessel cells, locally activated RHO-RELATED PROTEIN FROM PLANTS11 (ROP11; a small GTPase) recruits MIDD1 to the plasma membrane domains, which in turn promotes disassembly of cortical microtubules through the microtubule depolymerizer Kinesin-13A, resulting in the formation of secondary cell wall pits (Oda et al., 2010; Oda and Fukuda, 2013). Furthermore, we recently reported that the plant-specific scaffold proteins VESICLE TETHERING1 (VETH1) and VETH2 recruit the exocyst complex to cortical microtubules via the tethering protein CONSERVED OLIGOMERIC GOLGI COMPLEX2 to ensure microtubule-dependent cell wall deposition in xylem vessel cells (Oda et al., 2015; Vukašinović et al., 2017). Therefore, further exploring plant-specific proteins that are expressed in xylem vessel cells should lead to the discovery of more microtubule-associated proteins, shedding light on the mechanism underlying cortical microtubule organization.

In this study, we identified the microtubule-associated protein CORD1 (CORTICAL MICROTUBULE DISORDERING1), which functions as a regulator of cortical microtubule organization in xylem vessel cells. CORD1 is associated with cortical microtubules and is required for the proper deposition patterns of pitted secondary cell walls in metaxylem vessels. Ectopic expression of this protein caused partial detachment of cortical microtubules from the plasma membrane and disturbed the parallel arrays of cortical microtubules. Arabidopsis thaliana has six CORD1 paralogs, most of which decorate cortical microtubules in vivo. CORD genes are expressed in various tissues during plant development, suggesting that CORD family proteins are broadly involved in cortical microtubule organization.

RESULTS

CORD1 Associates with Cortical Microtubules

To identify microtubule-associated proteins involved in secondary cell wall patterning, we searched microarray and RNA-seq data for developing xylem (Ohashi-Ito et al., 2010; Ko et al., 2012). We selected uncharacterized xylem-expressed genes and fused them with GFP under the control of the estrogen-inducible LexA promoter (Zuo et al., 2000). We then investigated the localization of the GFP-fused gene products using the in vitro xylem vessel cell differentiation system, in which cultured Arabidopsis cells are synchronously differentiated into metaxylem vessel cells (Oda et al., 2010). Of the proteins investigated, an uncharacterized protein, which we designated CORD1, localized to microtubule-like filaments underlying secondary cell wall thickenings (Figure 1A). Indeed, GFP-CORD1 colocalized with cortical microtubules marked with BETA-6 TUBULIN (TUB6) tagged with the red fluorescence protein TagRFP in cultured non-xylem Arabidopsis cells (Figure 1B).

Figure 1.
Figure 1.

CORD1 Localizes along the Microtubule.

(A) Localization of CORD1-GFP in differentiating xylem vessel cells. Secondary cell walls are stained with fluorescent WGA.

(B) Localization of CORD1-GFP in non-xylem vessel cells. CORD1 is colocalized with cortical microtubules labeled with TagRFP-TUB6.

(C) to (E) FRAP analysis of CORD1-GFP. Kymograph generated from the photobleached area of the microtubule labeled with CORD1-GFP (C). Plot and curve fitting of intensity recovery after photobleaching ([D] and [E]).

(F) Localization of CORD1-GFP (proCORD1:CORD1-GFP) in N. benthamiana epidermal cells expressing VND6. Microtubules are marked with TagRFP-TUB6 (35Spro:TagRFP-TUB6).

(G) Time-lapse images of CORD1-GFP (proCORD1:CORD1-GFP) in N. benthamiana epidermal cells expressing VND6. Blue and red arrows indicate elongating and shrinking microtubules, respectively.

Bars = 10 μm in (A) and (B), 20 µm in (F), and 5 µm in (G).

To examine the localization of CORD1 at native expression levels, we introduced proCORD1:CORD1-GFP into the leaf epidermis of Nicotiana benthamiana, together with VASCULAR-RELATED NAC-DOMAIN6 (VND6) under the control of the estradiol-inducible LexA promoter (Zuo et al., 2000). CORD1-GFP was localized uniformly along microtubules, including to their plus ends and minus ends (Figures 1F and 1G; Supplemental Movie 1), suggesting that CORD1 is a microtubule side binding protein.

To examine whether CORD1 directly binds microtubules, we purified glutathione S-transferase (GST)-fused CORD1 protein from Escherichia coli. Unfortunately, the recombinant protein had a strong tendency to aggregate and could therefore not be used in the microtubule binding assay. We also purified GST-fused truncated CORD1 lacking its N-terminal 100 amino acids as well as hexa-histidine-fused CORD1 and GST-fused CORD2. However, these recombinant proteins still showed a tendency to aggregate. Therefore, instead of using recombinant proteins, we used fluorescence recovery after photobleaching (FRAP) analysis. Because the fluorescence recovery rate after photo bleaching indicates the dissociation rate of the protein from the microtubule (Bulinski et al., 2001), we could determine whether CORD1 binds microtubules by comparing the fluorescence recovery rate of CORD1 with that of known MAPs. We detected rapid recovery of CORD1-GFP fluorescence after photobleaching (Figures 1C to 1E); the average 50% recovery time of CORD1-GFP was 7.90 s (n = 23 microtubules). This was similar to the 50% recovery time of MAPs such as GFP-AtMAP65-1 (5.93 s), GFP-NtMAP65-1a (6.95 s), and MIDD1 (8.68 s) (Chang et al., 2005; Oda et al., 2010), suggesting that CORD1 directly binds to microtubules.

CORD Paralogs Associate with Microtubules

CORD1 encodes a 505-amino acid protein of unknown function. The Arabidopsis genome encodes six CORD1 paralogs with an E-value of less than 7e-21 (Supplemental Figure 1A and Supplemental Data Set 1). We designated these paralogs CORD2 to CORD7 (AT1G08760, CORD2; AT4G13370, CORD3; AT1G23790, CORD4; AT1G70340, CORD5; AT3G19610, CORD6; and AT2G31920, CORD7). CORD proteins are composed of a highly conserved N-terminal region and the remaining variable region (Supplemental Figures 1B and 2). Motif search by MEME (http://meme-suite.org/tools/meme) revealed the presence of 12 conserved putative motifs among CORD members (Supplemental Figure 1B). Analysis using the COILS program (Lupas et al., 1991) predicted coiled-coil domains in the C-terminal regions of CORD1, CORD3, CORD6, and CORD7 (Supplemental Figures 1B and 3). A BLAST search using full-length Arabidopsis CORD1 identified orthologs of CORD proteins in various plant species, including Populus trichocarpa, Oryza sativa, Brachypodium distachyon, Physcomitrella patens, and Marchantia polymorpha (Supplemental Figure 4 and Supplemental Data Set 2), but not in animals, yeast, or algae, indicating that the CORD family is conserved among (and unique to) land plants.

To examine whether CORD family proteins other than CORD1 also associate with microtubules, we observed the localization of GFP-tagged CORD proteins expressed under the control of the LexA promoter in N. benthamiana epidermal cells. All of the CORD proteins except CORD5 localized to microtubules (Figure 2). Like CORD1, CORD2-GFP, CORD3-GFP, and CORD7-GFP localized to microtubule arrays in all of 30 observed cells. CORD4-GFP also localized to microtubules in all 30 observed cells. Interestingly, 21 of the 30 observed cells exhibited short fragmented microtubules, suggesting that CORD4 induced microtubule fragmentation. Instead of localizing to microtubule, CORD5-GFP localized to the nucleus. This was unexpected because CORD5 shows high similarity to CORD4. We tested to determine whether the C-terminal fusion of GFP was responsible for the mislocalization CORD5 by examining the localization of an N-terminal fusion of GFP. GFP-CORD5 localized to the nucleus but was also localized to microtubules in 21 of 30 cells. In the remaining nine cells, GFP-CORD5 was localized only to the nucleus. Like CORD4-expressing cells, GFP-CORD5-expressing cells exhibited short fragmented microtubules. These observations suggest that CORD5 also associates with microtubules and has activity similar to that of CORD4. By contrast, CORD6-GFP localized to the cytoplasm in 23 of 30 observed cells. However, in the remaining seven cells, CORD6-GFP localized faintly to microtubules, suggesting that CORD6 also associates with microtubules. Our observations indicated that all CORD proteins have activity to associate with microtubule. Thus, we concluded that CORD proteins represent a previously unknown MAP family.

Figure 2.
Figure 2.

Subcellular Localization of CORD Proteins.

GFP-tagged CORD proteins (CORD-GFP and GFP-CORD5) and microtubules (TagRFP-TUB6) in leaf epidermal cells of N. benthamiana. Thirty cells were observed for each CORD protein. The numbers indicate the frequency of the cells in which CORD proteins localized to microtubules. Bars = 20 µm.

CORD1 Is Expressed in Xylem Vessel Cells

To elucidate the roles of CORD genes, we investigated their expression patterns in 80 different tissues based on information in the Arabidopsis RNA-seq database (TRAVA, http://travadb.org/) (Klepikova et al., 2016). We also obtained expression data for the well-known xylem vessel-expressed IRREGULAR XYLEM3 (IRX3), VND6, and XYLEM CYSTEINE PEPTIDASE1 (XCP1) genes as a reference for xylem vessel-related expression (Supplemental Data Set 3). We then performed clustering analysis of the genes based on their expression patterns (Supplemental Figure 5). CORD1 was assigned to a clade containing IRX3, VND6, and XCP1. This gene is highly expressed in the axes of the first elongated internode and silique pods, where secondary cell walls are actively formed, suggesting that CORD1 is involved in xylem development. By contrast, CORD3, CORD4, and CORD5 form another clade, with high expression levels in meristematic tissues. CORD6 and CORD7 show more specific expression patterns, while CORD6 is exclusively expressed in anthers and mature flower. Analysis of another gene expression database (Winter et al., 2007) suggested that CORD6 is specifically expressed in pollen. CORD7 is expressed at fairly high levels in young seeds, while CORD2 shows the broadest expression pattern among CORD family members. These expression patterns suggest that CORD family members play widespread roles in plant development, functioning in xylem vessels, meristem, seed, and reproductive tissues. Of the CORD proteins, CORD1 likely functions in xylem vessel-related tissues. Consistent with this, only CORD1 was upregulated in VND6-induced cultured xylem cells, while the other six CORD genes were not upregulated (Supplemental Table 1).

We then performed a promoter-GUS assay to more finely dissect the expression pattern of CORD1. We produced proCORD1:GUS plants harboring the GUS reporter gene under the control of a 2.0-kb fragment of the CORD1 promoter region. GUS signals were preferentially detected in xylem vessel cells in the root and cotyledon (Figures 3A and 3B). In roots, GUS signals were observed in developing protoxylem vessel cells and metaxylem vessel cells (Figure 3B). We also examined the localization of GFP-tagged CORD1 under the control of its own promoter in roots. Consistent with the GUS reporter assay, CORD1-GFP was preferentially detected in developing xylem vessels (Figures 3C and 3D). These results suggest that CORD1 is involved in xylem vessel development.

Figure 3.
Figure 3.

Expression Patterns of CORD Genes.

(A) and (B) GUS staining of ProCORD1:GUS plants. Cotyledon (A) and xylem vessels of roots (B) are shown. In (B), four different stages of roots are shown: differentiating protoxylem (top), differentiating metaxylem (two middle), and mature protoxylems and metaxylems (bottom). P and M indicate mature protoxylem and metaxylem vessel cells, respectively.

(C) and (D) CORD1-GFP at metaxylem (C) and protoxylem vessel cells (D) in the roots of proCORD1:CORD1-GFP plants.

Bars = 200 μm in (A) (top), 30 μm in (A) (bottom), 20 µm in (B), and 10 μm in (C) and (D).

CORD1 Is Required for Secondary Cell Wall Pit Formation

We isolated a T-DNA insertion mutant designated cord1 (SALK_073077) in which T-DNA was inserted in the first exon of the CORD1 locus (Figure 4A). However, the loss of CORD1 in this mutant did not cause any xylem vessel defects, likely due to the redundant functions of CORD family members (Figures 4C and 4D). We therefore crossed cord1 with cord2 (SALK_024770; Figure 4A), as CORD2 displayed the broadest expression pattern among family members, to generate the cord1 cord2 double mutant. Quantitative RT-PCR revealed that cord1 cord2 plants failed to express CORD1 and CORD2 (Figure 4B). Compared with wild-type plants, cord1 cord2 plants developed smaller pits in the secondary cell walls of metaxylem vessels (Figure 4C). The average area of pits in cord1 cord2 plants was ∼20% smaller than that in wild-type plants, while the sizes of pits in cord1 and cord2 plants were comparable to that in the wild type (Figure 4D; Supplemental Data Set 4). Introduction of proCORD1:CORD1-GFP complemented the small pit phenotype of the cord1 cord2 double mutant (Figures 4C and 4D; Supplemental Data Set 4). These data strongly suggest that CORD1 and CORD2 redundantly promote pit formation in the secondary cell walls of metaxylem vessels. Although CORD1 was also expressed in developing protoxylem vessels (Figure 3B), the cord1 cord2 double mutant did not show any visible phenotype in protoxylem vessel cells (Figure 4C). Other CORD members may redundantly function in protoxylem vessel cells.

Figure 4.
Figure 4.

CORD1 and CORD2 Are Required for Secondary Cell Wall Pit Formation.

(A) T-DNA inserts in the cord1 and cord2 mutants.

(B) Expression levels of CORD1 and CORD2 in wild-type and cord1 cord2 plants. Data are means ± sd (n = 3).

(C) Metaxylem vessel cells in the roots of the wild type, cord1 cord2 plants, and cord1 cord2 plants harboring proCORD1:CORD1-GFP. Differential interference contrast images (left) and fluorescence images of secondary cell walls stained with safranin (right) are shown. Arrowheads indicate secondary cell wall pits. Bars = 10 µm.

(D) Pit areas of metaxylem vessel cells in roots. Data are means ± sd (n > 100). *P < 0.01; ANOVA with Scheffe’s test.

Because CORD proteins associate with cortical microtubules, we assumed that loss of CORD1 and CORD2 would affect the cortical microtubule organization in metaxylem vessel cells. We introduced proIRX3:EYFP-TUB6 into wild-type plants and cord1 cord2 plants. However, we could not acquire images of sufficient quality to compare the cortical microtubule organization because the metaxylem vessel cells were located deep inside the roots. Instead, we developed an experimental system to ectopically induce metaxylem vessel differentiation in the excised hypocotyl tissues. We cultured excised hypocotyls of etiolated seedlings with 2,4-D and kinetin together with bikinin, which was recently found to promote xylem cell differentiation (Kondo et al., 2015). Six days after the onset of culture, metaxylem vessel-like cells with pitted cell walls appeared at the cortex and endodermis of the hypocotyl (Supplemental Figures 6A and 6B). The pitted cell walls were stained with safranin, indicating that the pitted cell walls were secondary cell walls (Supplemental Figure 6C). Consistent with this observation, xylem vessel-related genes, such as VND6, VND7, MIDD1, IRX3, XCP1, and CORD1, were more than 10-fold upregulated, while an phloem-related gene, ALTERED PHLOEM DEVELOPMENT, was upregulated but not to the same extent as the xylem vessel-related genes (Supplemental Figure 6D), suggesting that the culture induced metaxylem vessel differentiation (Supplemental Figure 6D). Next, we cultured the excised hypocotyls of wild-type plants and cord1 cord2 plants expressing proIRX3:EYFP-TUB6. The cord1 cord2 plants formed smaller pits of secondary cell walls than wild-type plants (Figures 5A and 5B), suggesting that CORD1 and CORD2 function similarly in the ectopic metaxylem vessels. We then observed the cortical microtubules in the ectopic metaxylem vessel cells at the earlier stage (Figures 5C and 5E) and later stage (Figures 5F and 5H) of secondary cell wall pit formation. The cortical microtubules in cord1 cord2 cells appeared to be more parallel than in wild-type cells at both stages (Figures 5C and 5F). The histograms of microtubule orientation frequency showed that the cortical microtubules tend to be more parallel in the cord1 cord2 cells that in wild-type cells (Figures 5D and 5G). Indeed, the mean standard deviations of the microtubule orientation in cord1 cord2 cells were lower than that of wild-type cells (Figures 5E and 5H). Importantly, the difference in the mean sd between cord1 cord2 cells and wild-type cells was more remarkable at the earlier stage (Figure 5E) than at the later stage (Figure 5H). These data suggest that CORD1 and CORD2 regulate cortical microtubule alignment to determine the size of secondary cell wall pits.

Figure 5.
Figure 5.

CORD1 and CORD2 Are Required for Cortical Microtubule Arrangement in Metaxylem Vessel Cells.

(A) Ectopic metaxylem vessel cells in wild-type and cord1 cord2 hypocotyls.

(B) Pit areas of ectopic metaxylem vessel cells in wild-type and cord1 cord2 hypocotyls. Data are means ± sd (n > 100), **P < 0.01 (Student’s t test).

(C) and (F) Cortical microtubules (proIRX3:EYFP-TUB6) in ectopic metaxylem vessel cells at early (C) and late (F) differentiation stages in wild type and cord1 cord2 hypocotyls.

(D) and (G) Histograms of the frequency of microtubule orientation in wild-type and cord1 cord2 cells of (D) and (G). Lines are curves fitted to the histogram by a Gaussian function.

(E) and (H) Standard deviation of the microtubule orientation in ectopic metaxylem vessel cells at early (E) and late (H) differentiation stages in wild-type and cord1 cord2 hypocotyls. Data are means ± sd (n > 21 cells), *P < 0.05 and **P < 0.01 (Student’s t test).

Bars = 20 µm in (A) and 10 µm in (C) and (F).

To investigate the role of CORD1 further in metaxylem vessel development, we examined the effects of CORD1 overexpression. We introduced proLexA:CORD1-GFP into transgenic plants harboring 35Spro:TagRFP-TUB6. After 2 d of CORD1 overexpression, the transgenic plants displayed enlarged pits in their metaxylem vessels (Figure 6C), as well as epidermal cell swelling in roots (Figures 6A and 6B). Upon overexpression of CORD1, the transverse cortical microtubule arrays in root epidermal cells were destroyed. Instead, fragmented, short cortical microtubules were randomly distributed in these cells (Figure 6D). Given that disrupting intact cortical microtubules causes cell swelling (Baskin et al., 1994), we reasoned that the swelling of epidermal cells in CORD1-overexpressing plants was indirectly caused by the disorganization of transverse cortical microtubules. Therefore, overexpression of CORD1 disturbed the parallel array of cortical microtubules, which in turn affected cell morphology.

Figure 6.
Figure 6.

Overexpression of CORD1 Causes Cell Swelling and Deformation of Secondary Cell Wall Patterns.

(A) to (C) Phenotypes of control plants (Control, 35Spro:TagRFP-TUB6) and CORD1-GFP-overexpressing plants (CORD1ox, 35Spro:TagRFP-TUB6/proLexA:CORD1-GFP). Roots (A), root epidermis (B), and xylem vessels (C) are shown. The epidermal cell walls were stained with propidium iodide.

(D) Cortical microtubules in root epidermal cells of control plants (Control) and CORD1-GFP-overexpressing plants (CORD1ox). Images were acquired 24 h after estradiol treatment. Red and green signals indicate TagRFP-TUB6 and CORD1-GFP, respectively.

Bars = 200 μm in (A), 20 μm in (B) and (D), and 5 µm in (C).

Exogenous Expression of CORD1 Disturbs the Cortical Microtubule Array

CORD1 has eight motifs that are conserved among CORD family members, four of which are located close together in the N-terminal region of CORD1, giving rise to a high-similarity domain in this family. We therefore suspected that the N-terminal region of CORD1 has microtubule binding activity. To investigate the functions of the conserved motifs in CORD1, we introduced GFP-fused truncated CORD1 fragments (Figure 6A) together with 35Spro:TagRFP-TUB6 into cultured non-xylem Arabidopsis cells. Unexpectedly, the N-terminal fragment of CORD1 (amino acids 1–130: CORD1N-terminal) did not localize to microtubules but instead localized to the cytoplasm (Figure 6A). However, truncated CORD1 (amino acids 131–505: CORD1∆N) lacking the N-terminal region localized to microtubules. Another truncated version of CORD1 (amino acids 131–400: CORD1∆N∆C) lacking both the C-terminal and N-terminal regions did not localize to microtubules (Figure 7A). CORD1∆1-300 (amino acids 301–505) did not localize to microtubules, whereas CORD1∆1-200 (amino acids 201–505) did (Figure 7A). These observations suggest that the region between amino acids 201 and 505 is necessary and sufficient for microtubule localization. This region contains motifs that are conserved among all CORD family members (Supplemental Figure 1B), suggesting that other CORD proteins also interact with microtubules through these conserved motifs.

Figure 7.
Figure 7.

CORD1 Overexpression Disturbs the Parallel Alignment of the Cortical Microtubule Array.

(A) GFP-tagged full-length CORD1 (proLexA:CORD1FL-GFP) or truncated CORD1 (proLexA:CORD1N-terminal-GFP, proLexA:CORD1∆N–GFP, proLexA:CORD1∆N∆C–GFP, proLexA:CORD1∆1-200-GFP, and proLexA:CORD1∆1-200-GFP) was introduced into cultured non-xylem vessel cells. Microtubules were marked with TagRFP-TUB6 (35Spro:TagRFP-TUB6). Note that cells expressing CORD1Nter or CORD1∆N exhibit randomized cortical microtubule networks (arrowheads). r indicates Pearson’s coefficient between red and green signals in the white box in the Merged channel. Note the low (r < 0.2) and high (r > 0.7) values for non-microtubule and microtubule localization, respectively. Bars = 10 µm.

(B) Box plot of anisotropic microtubule orientation. Anisotropy indicates the degree of randomization of the cortical microtubule array. Each box indicates the lower and upper quartiles (n > 80). The line in the box indicates the median. Cells were observed 24 h after treatment with estradiol (Est+) or DMSO (Est-). *P < 0.01; ANOVA with Scheffe’s test.

To investigate the functions of the conserved CORD1 motifs in microtubule organization, we made detailed observations of cortical microtubule organization in cultured cells expressing full-length or truncated CORD1. The majority of cortical microtubules were oriented transversely in control cells in which exogenous CORD1-GFP was not expressed. However, 24 h after the induction of CORD1-GFP expression, the parallel arrays of cortical microtubules were disturbed and randomized microtubule networks formed (Figure 7B). To evaluate the effects of CORD1 on cortical microtubule organization, we quantified the anisotropy of cortical microtubules. Cells expressing full-length CORD1-GFP, CORD1∆N-GFP, or CORD1∆1-200, which localized to microtubules, exhibited increased anisotropy of cortical microtubules (Figure 7B). By contrast, cells expressing CORD1N terminus, CORD1∆N∆C-GFP, or CORD1∆1-300, which did not localize to microtubules, failed to exhibit increased anisotropy of cortical microtubules (Figure 7B). These results indicate that microtubule binding of CORD1 is tightly coupled with its activity to disturb cortical microtubule arrays.

To explore how CORD1 affects cortical microtubule organization, we performed time-lapse observations of cortical microtubules in cultured non-xylem vessel cells expressing GFP-TUB6 together with proLexA:CORD1. To visualize the behavior of microtubules, we created overlapping images from frames taken at 0, 15, and 30 s from the time-lapse image sets, each of which was set on the red, green, and blue channels, respectively. This enabled us to easily recognize the growth, shrinkage, and sliding of microtubules. In the control cells, in which proLexA:CORD1 was not introduced, transverse cortical microtubules exhibited typical growth and shrinkage, with very little lateral sliding of cortical microtubules due to their tight attachment to the plasma membrane (Figure 8A; Supplemental Movie 2). By contrast, in cells expressing CORD1, sliding and wavering of microtubules were frequently observed in addition to growth and shrinkage (Figures 8B and 8C; Supplemental Movie 3). Such cortical microtubule behavior suggests that the cortical microtubules were partially detached from the plasma membrane. Supporting this idea, significant amounts of microtubules were observed in the cytoplasm (Figure 8E), while almost all of the microtubules were located at the peripheries of control cells (Figure 8D). Indeed, in CORD1-GFP-expressing cells, the amount of cytoplasmic microtubules was more than three times greater than in control cells (Figure 8F). Unlike typical cortical microtubules (Figure 8D; Supplemental Movie 4), the cytoplasmic microtubules in CORD1-expressing cells were not static but instead were quite dynamic (Figure 8E; Supplemental Movie 5). These observations suggest that CORD1 promotes partial detachment of cortical microtubules from the plasma membrane, which in turn causes sliding and wavering behavior in cortical microtubules.

Figure 8.
Figure 8.

CORD1 Promotes Mobility of Cortical Microtubules.

(A), (B), (D), and (E) Microtubules (GFP-TUB6) in cultured non-xylem vessel cells expressing 35Spro:GFP-TUB6 without ([A] and [D]) or with proLexA:TagRFP-CORD1 ([B] and [E]). The cortex ([A] and [B]) and mid-plane ([D] and [E]) are shown. Right panels show overlay images of the 0 s (red), 15 s (green), and 30 s (blue) period from time-lapse image sets. White signals indicate immobile microtubules. Colored microtubules indicate that the microtubules are growing, shrinking, or sliding.

(C) Magnified image of the white box in (B) and schematic illustration. Arrows indicate a wavering microtubule.

(F) Number of cytoplasmic microtubules in control cells and CORD1-expressing cells. Data are means ± sd (n = 15). *P < 0.01; Student’s t test.

(G) Range of angles of microtubule branches during a 30-s period after branch initiation. θ0sec indicates the angle of the microtubule branch at the initiation of branching. θ30sec indicates the angle of the microtubule branch 30 s after the θ0sec time point. Range of the angle = absolute value of (θ0sec − θ30sec).

Bars = 10 μm in (A), (B), (D), and (E) and 2 μm in (C).

Cortical microtubules undergo branch nucleation on preexisting cortical microtubules. The branching angle, which is typically 40 degrees (Murata et al., 2005), is thought to be important for the transverse array of cortical microtubules. Therefore, we investigated branching of the cortical microtubules. We recorded the angles of microtubule branches over a 30-s period after the initiation of branching. The average branching angle at branch initiation was 41.6 ± 7.1 degrees in CORD1-expressing cells and 42.2 ± 7.2 degrees in control cells (Figure 8G), both of which are typical branching angles, suggesting that CORD1 does not affect the average branching angle of cortical microtubules. However, the range of microtubule branching angles during a 30-s period after branch initiation significantly increased in response to CORD1 expression. In non-CORD1-expressing cells, the angle range of the branched microtubules during the 30-s period was <10 degrees (Figure 8G), with over 80% of the branched microtubules exhibiting angle ranges within 5 degrees. By contrast, in CORD1-expressing cells, only 50% of microtubules exhibited angle ranges within 5 degrees. Over 50% of the branched microtubules exhibited a broad angle range of 5 to 25 degrees (Figure 8G). These results indicate that CORD1 destabilizes microtubule angles after branching, probably by promoting the partial detachment of cortical microtubules from the plasma membrane. The unstable branching angles of the cortical microtubules after branch initiation, as well as their wavering and detachment from the plasma membrane, are likely responsible for the disorganization of cortical microtubules in response to CORD1 expression.

In this context, we hypothesized that CORD1 might interact with the plasma membrane to disturb the interaction between cortical microtubules and the plasma membrane. To test this possibility, we observed CORD1-GFP localization in z-stack images of cultured cells (Supplemental Figure 7). CORD1-GFP was localized not only to cortical microtubules but also to the cytoplasmic microtubules (Supplemental Figure 7), suggesting that CORD1 does not specifically localize to cortical microtubules. In the epidermis of N. benthamiana leaves expressing CORD1-GFP and the plasma membrane marker, PLDalpha4-TagRFP, CORD1-GFP did not localize to the plasma membrane after treatment with 30 μM oryzalin, which entirely depolymerizes cortical and cytoplasmic microtubules (Supplemental Figure 8), indicating that CORD1 does not interact with the plasma membrane.

CORD1 Impairs the Lateral Restriction of Active ROP11 Domains by Cortical Microtubules

We previously showed that the sizes of secondary cell wall pits are determined by Kinesin-13A. Kinesin-13A is recruited to the plasma membrane domains of activated ROP11 GTPases via MIDD1 to locally disrupt cortical microtubules (Oda and Fukuda, 2012a, 2013). Loss of Kinesin-13A leads to the formation of small secondary wall pits (Oda and Fukuda, 2012a, 2013), which resembles the phenotype of cord1 cord2 plants. We also found that cortical microtubules surrounding the active ROP domains laterally inhibit the diffusion of active ROP11 domains, resulting in a mutual inhibitory interaction between the active ROP domains and cortical microtubules (Oda and Fukuda, 2012a). Considering the small secondary cell wall pits in cord1 cord2 plants, we reasoned that CORD1 might influence the active ROP11 domains that govern the behavior of Kinesin-13A. However, CORD1 did not directly interact with ROP11 in a bimolecular fluorescence complementation assay (Supplemental Figure 9). Thus, we postulated that CORD1 might indirectly influence ROP11 domains by altering cortical microtubule organization. To test this idea, we reconstructed the active ROP domains in N. benthamiana non-xylem leaf epidermal cells (Oda and Fukuda, 2012a) and examined the effects of CORD1 on the spatial interactions between the ROP domains and cortical microtubules. We introduced three constructs, proLexA:ROP11, proLexA:ROPGAP3, and proLexA:ROPGEF4PRONE, into N. benthamiana leaves to reconstruct the active ROP11 domains, together with proLexA:TagRFP-MIDD1 and proUBQ10:EYFP-TUB6, which served as markers for active ROP11 and microtubules, respectively. As previously shown, active ROP11 domains marked with TagRFP-MIDD1 were detected on the plasma membrane. Cortical microtubules defined the boundaries of the activated ROP domains, confining the localization of the domains (Figures 9A and 9C). Cortical microtubules were rarely observed within the active ROP domains. By contrast, when CORD1-ECFP was introduced into the cells together with the five constructs, microtubules no longer determined the boundaries of ROP domain localization. Microtubule fragments were frequently observed within active ROP domains, as well as outside of active ROP11 domains (Figures 9B and 9D). Indeed, the number of microtubules that overlapped with active ROP domains increased 2.5-fold in response to CORD1 expression (Figure 9E). This observation suggests that CORD1 disrupts the boundaries of active ROP domains as well as the mutual inhibitory interactions between ROP11 domains and cortical microtubules.

Figure 9.
Figure 9.

CORD1 Abolishes the Restricted Localization of Active ROP Domains by Cortical Microtubules.

(A) and (B) Active ROP domains reconstituted in N. benthamiana epidermal cells. ROPGEF4PRONE, ROP11, ROPGAP3, GFP-TUB6 (green), and TagRFP-MIDD1∆N (red) were co-introduced without (A) or with (B) CORD1-ECFP. Bars = 10 µm.

(C) Magnified image of the white box in (A).

(D) Magnified image of the white box in (B). Arrowheads indicate cortical microtubules overlapping with active ROP domains.

(E) Number of cortical microtubules overlapping with active ROP domains. Data are means ± sd (n > 50). *P < 0.01; Student’s t test.

DISCUSSION

CORD1 Is a Microtubule-Associated Protein

In this study, we identified CORD1 as a previously unknown protein associated with microtubules. CORD1 localized to cortical microtubules and exhibited rapid turnover on microtubules, similar to microtubule side binding proteins. Although we were unable to test the direct binding of CORD1 with microtubules in vitro, our data strongly suggest that CORD1 is a microtubule-associated protein. Overexpression of CORD1 led to disorganization of the cortical microtubules in roots and cultured cells. Conversely, loss of CORD1 and CORD2 led to excess parallel cortical microtubules in metaxylem vessel cells. In cultured cells, only 24-h CORD1 induction was sufficient to induce microtubule disorganization without any defects in cell morphology. Thus, it is likely that CORD1 directly affects microtubule organization. Close observation of cortical microtubules in CORD1-expressing cells revealed that CORD1 causes atypical wavering behavior in cortical microtubules. Furthermore, the branching angles of cortical microtubules, which normally remain at 40 degrees from the mother cortical microtubules, became unstable in the presence of CORD1. The amount of cytoplasmic microtubules increased with the increase in CORD1 expression. Collectively, our data suggest that CORD1 promotes disorganization of the cortical microtubule array by weakening the attachment of cortical microtubules to the plasma membrane.

The mechanism by which CORD1 weakens the attachment of microtubules to the plasma membrane is unknown. The small compound morlin (DeBolt et al., 2007), as well as n-butanol (Gardiner et al., 2003), can induce the detachment of cortical microtubules from the plasma membrane and disorganize the parallel cortical microtubule array, as observed in CORD1-expressing cells. Morlin also affects the motility of CESA complexes, which suggests that morlin may inhibit the interaction between cortical microtubules and CESA complexes (DeBolt et al., 2007). Like morlin, CORD1 might also inhibit the interaction between cortical microtubules and the cellulose synthase complex. On the other hand, n-butanol antagonizes phosphatidic acid production by phospholipase D. N-acylethanolamine, which specifically inhibits the phospholipase Dα isoform, also disrupts cortical microtubules (Motes et al., 2005). Phosphatidic acids activate various types of signaling to microtubules and actin microfilaments (Pleskot et al., 2013). Interestingly, phosphatidic acids interact with MAP65-1 to activate the tubulin binding activity of MAP65-1 (Zhang et al., 2012). Thus, it is also possible that CORD1 inhibits phosphatidic acid signaling to microtubules. Because n-butanol can directly disrupt microtubules (Hirase et al., 2006), we cannot exclude the possibility that CORD1 directly affects the microtubule structure.

CORD1 Regulates the Sizes of Secondary Cell Wall Pits in Metaxylem Vessels

Our in silico expression analysis, promoter-GUS analysis, and analysis of proCORD1:CORD1-GFP plants suggested that CORD1 is expressed in the xylem vessels. Consistent with this notion, cord1 cord2 plants, which lack functional expression of CORD1 together with its closest paralog, CORD2, displayed abnormally small pits in the secondary cell walls of metaxylem vessels, strongly suggesting that CORD1 is required for normal secondary cell wall pit formation in xylem vessels. Conversely, overexpressing CORD1 led to the production of enlarged secondary cell wall pits, suggesting that CORD1 regulates the sizes of secondary cell wall pits. We previously showed that the loss of Kinesin-13A causes a similar phenotype. Kinesin-13A is a microtubule depolymerizer that preferentially localizes to secondary cell wall pits and induces depolymerization of cortical microtubules. Unlike Kinesin-13A, however, CORD1 was not specifically localized to secondary cell wall pits but was broadly localized to the cortical microtubules surrounding secondary cell wall pits to promote disordering of cortical microtubules. Thus, the role of CORD1 in regulating secondary cell wall pit size is different from that of Kinesin-13A.

We previously demonstrated that, in developing metaxylem vessel cells, locally activated ROP11 recruits the MIDD1-Kinesin-13A complex to induce microtubule disruption, resulting in the formation of secondary cell wall pits (Oda and Fukuda, 2013). We also showed that cortical microtubules surrounding active ROP11 domains inhibit the outward diffusion of active ROP11 GTPases to restrict the area occupied by active ROP11 domains, causing a mutual inhibitory interaction between cortical microtubules and ROP domains (Oda and Fukuda, 2012b). In this study, we reconstructed active ROP domains in N. benthamiana leaf epidermal cells and confirmed that cortical microtubules spatially restrict ROP domains. We then determined that CORD1 abolishes the spatial restriction of reconstructed ROP domains by cortical microtubules. This effect is likely due to the partial detachment of cortical microtubules as well as the disorganized alignment of cortical microtubules, which collectively reduce the frequency of physical interactions between cortical microtubules and ROP11 on the plasma membrane.

We previously showed that disrupting cortical microtubules in differentiating xylem vessel cells causes outward diffusion of active ROP11 domains (Oda and Fukuda, 2012b). Thus, in differentiating xylem vessel cells, the partial detachment and disorganization of cortical microtubules likely promote the outward diffusion of active ROP domains, resulting in larger areas occupied by active ROP11 domains, which in turn gives rise to larger secondary cell wall pits. Under this scenario, the high levels of CORD1 lead to random cortical microtubules that are partially detached from the plasma membrane, which allows ROPs to spread even more widely along the plasma membrane, resulting in larger secondary cell wall pits (Figure 10). Conversely, the loss of CORD1 maximizes the attachment of cortical microtubules to the plasma membrane and induces the formation of a highly ordered microtubule array, which strongly confines the area occupied by ROPs on the plasma membrane, resulting in smaller secondary cell wall pits (Figure 10). Thus, CORD1 likely determines the sizes of secondary cell wall pits by regulating the restriction of ROP by cortical microtubules.

Figure 10.
Figure 10.

Schematic Model of the Role of CORD1 in Secondary Cell Wall Pit Formation.

(A) During xylem vessel differentiation, CORD1 induces detachment of microtubules from the plasma membrane, resulting in disordering of cortical microtubule arrays.

(B) In metaxylem vessel cells, active ROP domains disrupt the cortical microtubules through the activity of the MIDD1-Kinesin-13A complex. Cortical microtubules surrounding the active ROP domains laterally restrict the distribution of active ROP domains. In the absence of CORD1 (cord1 cord2 mutant), the cortical microtubule array may be hyperparallel and tightly attached to the plasma membrane, which strongly restricts the active ROP domains, resulting in smaller secondary cell wall pits. In the presence of CORD1 (wild-type and CORD1-overexpressing plants), the arrangement of the cortical microtubule array becomes random, allowing broad diffusion of active ROP11, thereby resulting in larger secondary cell wall pits than those of cord1 cord2 plants.

The CORD Family Is Conserved in Land Plants

We identified six CORD1 paralogs in the Arabidopsis genome. Further analysis revealed 12 conserved motifs in this protein family, as well as coiled-coil domains in the C-terminal regions in some family members. Truncation analysis revealed that the motif located at the C-terminal region is required for the association of CORD1 with the microtubule. This motif is conserved in all seven CORD proteins in Arabidopsis, suggesting that all CORDs are microtubule-associated proteins. Indeed, CORD proteins localized to the cortical microtubules in N. benthamiana leaf epidermal cells.

Interestingly, CORD5, and its closest homolog, CORD4, induced short fragmented cortical microtubules in N. benthamiana leaf epidermal cells. Thus, the functions of CORD5, as well as CORD4, might be different from those of other CORD proteins. Indeed, CORD5 also localized to the nucleus. Some microtubule-associated proteins are known to localize to the nucleus. For example, END BINDING PROTEIN 1c is localized to the nucleus during interphase, depending on its C-terminal nuclear localization signal, but decollates spindle microtubules during mitosis (Komaki et al., 2010). Similarly, CORD5, and possibly CORD4, may associate with microtubules during mitosis.

The CORD genes displayed various expression patterns, including expression in xylem vessels, meristem, and reproductive tissues, suggesting that they play diverse roles in plant development. Among the CORD genes, CORD2 showed relatively broad expression, indicating that CORD2 redundantly functions with other CORD proteins. Indeed, our mutant analysis demonstrated that CORD2 functions redundantly with CORD1 in metaxylem vessels. Our BLAST analysis revealed that CORD genes are conserved among diverse plant species, including mosses, monocots, dicots, and trees, but are not found in algae. Thus, CORD proteins may play fundamental roles in microtubule organization and possibly cell morphogenesis in land plants.

During plant development, CORD proteins might facilitate the arrangement of cortical microtubules. Partial detachment of cortical microtubules was observed in maturing leaf epidermal cells, which is thought to promote the orientation of cortical microtubules from transverse to longitudinal (Sainsbury et al., 2008). Disordered cortical microtubules have also been observed during various events such as root hair development (Van Bruaene et al., 2004), trichome initiation (Tian et al., 2015), and early embryo development (Kimata et al., 2016). CORD proteins might contribute to the rearrangement of cortical microtubules by promoting the disorganization of cortical microtubules during these events. Further analysis of CORD family members should shed light on the mechanism by which cortical microtubules are actively disordered, which may facilitate the rearrangement of cortical microtubules to ensure proper plant cell morphogenesis.

METHODS

Plant Materials and Cell Culture

The mutant and transgenic Arabidopsis thaliana plants analyzed in this study were in the Columbia-0 (Col-0) background. T-DNA insertion mutants cord1 (SALK_073077) and cord2 (SALK_024770) were obtained from the ABRC (https://abrc.osu.edu/). To generate proLexA:CORD1-GFP/35Spro:TagRFP-TUB6 transgenic plants, the 35Spro:TagRFP-TUB6 plants (Oda and Fukuda, 2013) were transformed with proLexA:CORD1-GFP. Cell culture and transformation of Arabidopsis Col-0 suspension cells were performed as described previously (Oda and Fukuda, 2013).

Microscopy

Plants and cultured cells were observed under an inverted fluorescence microscope (IX83-ZDC; Olympus) fitted with a confocal unit (CSU-W1; Yokogawa), an UPLSAPO 60xW water immersion lens (NA 1.2), and laser lines set at 445, 488, and 561 nm. The acquired images were analyzed using MetaMorph (Molecular Devices) and ImageJ (http://rsbweb.nih.gov/ij/).

To evaluate colocalization with microtubules, Pearson’s coefficient between the green channel and red channel was determined using ImageJ software. The 100 × 100-pixel regions were cropped from the acquired images, and the background of the cropped regions was removed using the rolling ball method (20 pixels radius). Then, Pearson’s coefficient was calculated using JACoP (Just Another Colocalization Plug-in).

Microtubule orientation was quantified using ImageJ software. The acquired images were filtered through a fast Fourier transformation (FFT) band-pass filter (filter large 10, filter small 5, and tolerance 0) and processed with the threshold command Default B&D. The processed images were cropped to 100 × 100 pixels, and texture analysis was performed via FFT using the FFT command. FFT-treated images were filtered through a Gaussian blur filter (sigma = 5) and processed with the threshold command Otsu. Finally, the circularity of the processed images was calculated using the analyze particle command. Circularity was defined as the anisotropy of microtubule orientation.

For time-lapse movies, images were acquired every 3 s. The time-lapse image stacks were converted to AVI files (10 frames per second) using ImageJ software.

Constructs

To generate the expression vectors, DNA fragments from CORD1, CORD2, CORD3, CORD4, CORD5, CORD6, CORD7, and truncated CORD1 were amplified from Arabidopsis Col-0 cDNA using the appropriate primers (Supplemental Table 2) and cloned into the pENTR/D-TOPO entry vector (Invitrogen). These entry clones were recombined with the pER-XG expression vector for the C-terminal GFP fusion or the pER-XR expression vector for the C-terminal tag-RFP fusion (Oda and Fukuda, 2012a) using LR Clonase II mix (Invitrogen). To analyze the localization of CORD1 and to perform complementation of the cord1 cord2 double mutant, fragments of the 2.0-kb CORD1 promoter region and a genomic region of CORD1 containing a 2-kb promoter fragment were PCR amplified and cloned into the pENTR/D-TOPO entry vector, respectively. The promoter fragment and genomic region fragment of CORD1 with the 2-kb promoter region were recombined in the pGWB533 expression vector for GUS analysis and in the pGWB501 expression vector for expression of the C-terminal GFP fusion (Nakagawa et al., 2007). To make 35Spro:PLDalpha4-TagRFP, the coding sequence of Arabidopsis PLDalpha4 was PCR amplified and cloned into the pENTR/D-TOPO vector, and the resulting clone was recombined with the pGWB460 vector. To generate constructs for the bimolecular fluorescence complementation assay, CORD1 and ROP11 (Oda and Fukuda, 2012a) entry clones were recombined with pER-X-YN and pER-X-YC vectors (Oda et al., 2015). To make proIRX3:EYFP-TUB6, the promoter region of IRX3 was PCR amplified from the Arabidopsis genome and cloned into the pENTR/D-TOPO vector. The EYFP-TUB6 fragment was PCR amplified from proUBQ10:EYFP-TUB6 (Sugiyama et al., 2017) and inserted into the AscI site of the pENTR/D-TOPO vector harboring proIRX3, and the resulting clone was recombined with the pGWB501 vector.

Transformation of Nicotiana benthamiana

Transient transformation of N. benthamiana leaves was performed as described previously (Oda and Fukuda, 2012a). For reconstruction of active ROP domains, proLexA:ROP11, proLexA:ROPGAP3, proLexA:ROPGEF4PRONE, and proLexA:TagRFP-MIDD1 (Oda and Fukuda, 2012a) were used. 35Spro:TagRFP-TUB6 (Oda et al., 2010) was used for labeling microtubules.

FRAP Analysis

FRAP was performed under an Olympus FV-1200 confocal microscope equipped with a 488-nm laser. FRAP was observed in leaf epidermal cells of N. benthamiana harboring CORD1-GFP, which was expressed for 24 h following induction with 0.2 µM estradiol. In the FRAP experiments, 640 × 56-pixel areas including microtubules were bleached for 1 s, and the images of fluorescence recovery were collected every 0.5 s. The fitting curve and t1/2 were calculated as described previously (Oda et al., 2010).

GUS Assay

Cotyledons and roots were incubated in GUS-staining buffer (0.5 mg mL−1 X-gluc and 100 mM phosphate buffer, pH 7.0) for 12 h at room temperature, and the cotyledons were subsequently cleared with 70% ethanol for 12 h at room temperature.

Quantitative RT-PCR

Total RNA was prepared from Arabidopsis cells using the SDS-phenol method, treated with DNase, and purified using an RNeasy Plant Mini Kit (Qiagen) as described previously (Oda and Fukuda, 2013). After reverse transcription with the oligo(dT20) primer and SuperScript III reverse transcriptase (Thermo Scientific), quantitative RT-PCR was performed using a LightCycler 96 instrument (Roche Diagnostics) with FastStart Essential DNA Green Master (Roche Diagnostics).

Expression Data Analysis

The expression levels of genes in 80 different tissues were obtained from the TRAVA database (Klepikova et al., 2016). Heat maps were produced to visualize the clustering results using the Heatplus R package from Bioconductor version 3.4 (http://www.bioconductor.org/).

Hypocotyl Culture

Hypocotyls of etiolated seedlings 5 d after germination were excised and cultured in liquid MS medium containing 2.3 g/L MS salt mix, 10 g/L sucrose, and B5 vitamins including 0.5 mg/L nicotinic acid, 0.1 mg/L thiamine HCl, 0.5 mg/L pyridoxine HCl, 0.1 g/L myo-Inositol, and 2 mg/L glycine (pH 5.8), supplemented with 1 mg/L kinetin, 0.1 mg/L 2,4-D, and 10 µM bikinin in 12-well plates at 22°C and 60 rpm under continuous light.

For quantitative RT-PCR, total RNA was extracted from ∼100 hypocotyls at each time point and analyzed as described above. Three replicates were performed.

The microtubule orientation histograms were determined using ImageJ software. Three 200 × 200-pixel regions were collected from each cell and denoised using a Gaussian filter (sigma = 1) and the subtract background function (10-pixel radius). Histogram and curve fitting were performed using the “Directionality” plug-in (https://imagej.net/Directionality). Local orientation of microtubules was determined using a 5×5 sobel filter.

Accession Numbers

Sequence data from this article can be found in the GenBank/EMBL libraries under the following accession numbers: AT3G14170, CORD1; AT1G08760, CORD2; AT4G13370, CORD3; AT1G23790, CORD4; AT1G70340, CORD5; AT3G19610, CORD6; AT2G31920, CORD7; XP_001785292, jgi|Phypa1_1|101176; XP_001772202, jgi|Phypa1_1|234615; XP_001752982, jgi|Phypa1_1|65610; and OAE28537, Mapoly0073s0066.

Supplemental Data

Acknowledgments

We thank N. Chua (Rockefeller University) for the pER8 vector, U. Grossniklaus (University of Zurich) for the pMDC7 vector, and T. Nakagawa (Shimane University) for the pGWB vectors. We also thank T. Higaki (Kumamoto University) for technical advice and Y. Noguchi (National Institute of Genetics) and F. Hasegawa (National Institute of Genetics) for technical assistance. This work was supported by Grants-in-Aid from the Ministry of Ministry of Education, Culture, Sports, Science, and Technology of Japan (Grants 16H01247 and 15H01243 to Y.O. and 15H05958 to H.F.), the Japan Society for the Promotion of Science (Grants 16H06172 to Y.O. and 16H06377 to H.F.), the Japan Science and Technology Agency (Precursory Research for Embryonic Science and Technology project to Y.O.; Grant JPMJPR11B3), the Mitsubishi Foundation to Y.O., and the Naito Foundation to H.F.

AUTHOR CONTRIBUTIONS

Y.O. and H.F. designed the research. T.S. and Y.O. performed experiments. Y.O. developed the hypocotyl system. T.S., Y.O., and H.F. wrote the article.

Footnotes

  • Received August 22, 2017.
  • Revised October 10, 2017.
  • Accepted November 8, 2017.
  • Published November 13, 2017.

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CORTICAL MICROTUBULE DISORDERING1 Is Required for Secondary Cell Wall Patterning in Xylem Vessels
Takema Sasaki, Hiroo Fukuda, Yoshihisa Oda
The Plant Cell Dec 2017, 29 (12) 3123-3139; DOI: 10.1105/tpc.17.00663
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