Plant Cell Advance Publication. Published on September 25, 2017, doi:10.1105/tpc.17.00309

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RESEARCH ARTICLE

Two Complementary Mechanisms Underpin Cell Wall Patterning during Xylem Vessel Development Rene Schneider1,2, Lu Tang3, Edwin R. Lampugnani1, Sarah Barkwill4, Rahul Lathe2, Yi Zhang2, Heather E. McFarlane1, Edouard Pesquet5,6, Totte Niittyla6, Shawn D. Mansfield4, Yihua Zhou3, Staffan Persson1,2# 1

School of Biosciences, University of Melbourne, Parkville 3010, Melbourne, Australia Max-Planck Institute for Molecular Plant Physiology, Am Muehlenberg 1, 14476 Potsdam, Germany 3 State Key Laboratory of Plant Genomics, Institute of Genetics and Developmental Biology, Chinese Academy of Sciences, Beijing, 100101, China 4 Department of Wood Science, University of British Columbia, Vancouver, BC, Canada 5 Arrhenius laboratories, Department of Ecology, Environment and Plant Sciences (DEEP), Svante Arrhenius väg 20A, Stockholm University, 160 91 Stockholm, Sweden 6 Umeå Plant Science Centre (UPSC), Department of Forest Genetics and Plant Physiology, Swedish University of Agricultural Sciences, 901 87 Umeå, Sweden 2

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Corresponding author: [email protected]

Short title: CSI1/POM2 steers xylem vessel wall patterning One-sentence summary: The CELLULOSE SYNTHASE INTERACTING 1 protein directs secondary wall patterning during the early phases of xylem vessel development. ABSTRACT The evolution of the plant vasculature was essential for the emergence of terrestrial life. Xylem vessels are solute-transporting elements in the vasculature that possess secondary wall thickenings deposited in intricate patterns. Evenly dispersed microtubule (MT) bands support the formation of these wall thickenings, but how the MTs direct cell wall synthesis during this process remains largely unknown. Cellulose is the major secondary wall constituent and is synthesized by plasma membranelocalized cellulose synthases (CesAs) whose catalytic activity propels them through the membrane. We show that the protein CELLULOSE SYNTHASE INTERACTING (CSI)1/POM2 is necessary to align the secondary wall CesAs and MTs during the initial phase of xylem vessel development in Arabidopsis thaliana and Oryza sativa (rice). Surprisingly, these MT-driven patterns successively become imprinted and sufficient to sustain the continued progression of wall thickening in the absence of MTs and CSI1/POM2 function. Hence, two complementary principles underpin wall patterning during xylem vessel development.

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INTRODUCTION

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The plant vasculature is one of the most important evolutionary innovations for terrestrial

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life, as it allowed plants to adapt and grow to significant stature (Myburg et al., 2013). The

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xylem tissue provides essential functions in the vasculature by distributing water throughout

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the plant and providing structural support to the plant body. The xylem cells are encased by

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thickened cell walls that reinforce them and therefore are essential for their function (Turner

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et al., 2007). The organisation of the secondary cell walls differs between xylem vessel cell 1 ©2017 American Society of Plant Biologists. All Rights Reserved

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types and is typically described either as an annular/spiral pattern (called proto-xylem) or a

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reticulate/pitted pattern (called meta-xylem, Pesquet et al., 2011). Before these thickened

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secondary walls are assembled, the xylem cells, like all plant cells, are encased by a flexible

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but strong primary cell wall (Somerville et al. 2004). These walls largely comprise

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polysaccharides, of which cellulose, an unbranched, linear β-1,4-linked glucan, forms a

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significant constituent. Cellulose is synthesized at the plasma membrane by large cellulose

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synthase (CesA) complexes (CSCs; Schneider et al., 2016). The CSCs are composed of a

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heterotrimeric configuration of 18 to 24 CesAs where CesA1, CesA3 and CesA6-like (i.e.

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CesA2, 5, 6 and 9) CesAs produce primary wall cellulose in Arabidopsis thaliana, and

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CesA4, CesA7 and CesA8 comprise the CSCs necessary to make secondary wall cellulose

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(Persson et al., 2007; Desprez et al., 2007; Taylor et al., 2003; Atanassov et al., 2009).

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The CSCs move along linear tracks at the plasma membrane (Paredez et al., 2006),

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likely due to the catalytic activity of the CSCs. Nascent cellulose microfibrils become

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entrapped in the cell wall and further synthesis therefore exerts a force on the CSCs that

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propels them forward through the plasma membrane. The movement of the CSCs is guided

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by cortical microtubules (MTs) during both primary and secondary wall cellulose synthesis

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(Paredez et al., 2006; Watanabe et al., 2015). The protein CELLULOSE SYNTHASE

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INTERACTING 1 (CSI1), also called POM-POM 2 (POM2) is necessary for the MT-based

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guidance of the primary wall CSCs, as lesions in the protein impaired co-alignment between

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tracks of primary wall CSCs and cortical MTs (Bringmann et al., 2012; Li et al., 2012).

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However, reports on the function of CSI1/POM2 during secondary wall cellulose production

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differ. Gu and Somerville (2010) reported no defects on secondary walls nor decreased

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cellulose content in csi1/pom2 mutant Arabidopsis stems. By contrast, Derbyshire et al.

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(2015) showed that induction of tracheary elements in Arabidopsis cell cultures was impaired

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in cells with reduced CSI1/POM2 expression. The role of CSI1/POM2 in secondary wall

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cellulose production therefore remains unclear.

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Secondary walls are typically produced around cells that are situated deep in tissues,

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and that therefore are largely masked by other cells. This location makes it difficult to study

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secondary wall synthesis where it normally occurs. Instead, alternative systems have been

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developed for this purpose, including trans-differentiating cell cultures that can be induced by

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hormone cocktails (e.g. Kubo et al., 2005; Demura et al., 2002; Pesquet et al., 2010) and

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inducible transcription factor-based systems. The latter systems make use of the NAC-related

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transcription factors VASCULAR-RELATED NAC-DOMAIN 6 (VND6) and VND7 that

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promote meta- and proto-xylem-like cell wall structures, respectively (Kubo et al., 2005; 2

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Yamaguchi et al., 2010; Oda et al., 2010). By selectively controlling the activity of the VNDs

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with an inducible promoter system it is possible to induce and explore secondary wall

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formation in cells that normally do not form these structures. VND7-inducible Arabidopsis

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seedlings have been used to evaluate the behavior of the secondary wall CSCs using a

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fluorescently tagged CesA7 (Watanabe et al., 2015), and to assess the coordination between

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transcripts and metabolites during this process (Li et al., 2016a).

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Here, we investigated how proto-xylem vessel wall patterns are controlled by

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analysing the coordination of MTs and cell wall deposition in Arabidopsis and rice. We

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found that CSI1/POM2 orchestrates cell wall synthesis along MTs during the initial

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developmental phase of xylem vessel formation, but that subsequent synthesis occurs via a

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CSI1/POM2 autonomous mechanism. Our results indicate that cell wall patterns are directed

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by two complementary principles during xylem vessel development.

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3

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RESULTS

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CSI1/POM2 influences xylem vessel wall patterning

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To evaluate if defects in CSI1/POM2 function alter cell wall patterning during xylem vessel

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formation in Arabidopsis we examined secondary wall formation in three different systems

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where the function of CSI1/POM2 was impaired. First, we confirmed that the down-

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regulation of CSI1/POM2 caused aberrant secondary wall deposition in proto- and meta-

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xylem trans-differentiating cell suspension cultures (Derbyshire et al., 2015; Supplemental

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Figures 1A, B). Using confocal microscopy, we quantified the occurrence of spiral, reticulate,

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and pitted secondary wall patterns, and the percentage of calcofluor-stained irregular deposits

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in the secondary walls of non-transgenic and CSI1 down-regulated cell lines (Supplemental

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Figures 1C, D). Although it was difficult to assess defects in cell wall patterning in these

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lines, down-regulation of CSI1/POM2 caused a significant increase in irregular deposits

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along the secondary walls (Supplemental Figures 1B, D). This defect was irrespective of the

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patterning of the secondary walls (Supplemental Figure 1D).

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We next investigated if the xylem of mature stems of Arabidopsis plants showed

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structural defects when CSI1/POM2 function was impaired. We made longitudinal sections

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of the first internodes allowing structural characterization of intact and transected xylem

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vessels in the previously described csi1/pom2 mutants pom2-4 and csi1-1/pom2-8

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(Bringmann et al., 2012) as well as wild-type plants (Figures 1A, B). We found that the

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secondary wall bands were significantly more disordered in the pom2-4 and csi1-1/pom2-8

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mutants, as evident from measuring the spread in orientation angles of neighboring wall

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bands (Figure 1C).

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We next used a VND7-inducible Arabidopsis line (Yamaguchi et al., 2010) to study

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proto-xylem vessel secondary wall patterning. Here, we observed xylem-related wall

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synthesis as indicated by well-organized band patterns that were transversely and evenly

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distributed around induced hypocotyl cells (Figure 1D). We quantified the geometry of the

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bands and found that they were aligned tightly around an average angle of 0.6 ± 3.8° (mean

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± S.D., 132 cells from 5 seedlings) against the horizontal axis (Figures 1D, F).

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To assess if the CSI1/POM2 function influenced the wall patterns, we introgressed the

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pom2-4 mutant into the VND7-inducible line. The xylem vessel wall patterns were less well

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aligned in the pom2-4 background (Figures 1E, F). Here, the bands displayed significantly

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wider and less uniform angles as compared to control (-2.4 ± 22.7°; 136 cells from 5

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seedlings, Figures 1E to G). In addition, the band spacing was substantially altered in the

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pom2-4 mutant as compared to the control VND7-inducible line (Figure 1H). These results

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indicate that while CSI1/POM2 is not essential for the formation of secondary wall bands,

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confirming that xylem vessels are intact (Gu and Somerville, 2010), the protein influences the

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geometry and relative position of the deposition of the bands.

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To investigate if the defects in secondary wall patterning were associated with

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changes in cell wall architecture and ultrastructure, we measured the microfibril angle

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(MFA), cell wall crystallinity, degree of cellulose polymerization (DP), and cellulose content

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in pom2-4 mutant stems and compared the results with wild-type stems (Figures 1I to L). We

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found that the cellulose showed differences in both MFAs and crystallinity (Figures 1I, J),

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corroborating defects in cellulose synthesis. We also found a slight increase in glucose

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content, most likely due to an increase in amorphous cellulose due to the decreased levels of

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crystalline cellulose (Figure 1L). These data indicate that CSI1/POM2 influence the quality

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of secondary wall cellulose synthesis.

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CSI1/POM2 mimics the behavior of, and can interact with, the secondary wall CesA

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proteins

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To investigate how the CSI1/POM2 behaves during the transition from primary to secondary

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wall synthesis, we crossed plants expressing a functional, native promoter-driven triple (3x)

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YFP translational fusion with CSI1/POM2 (Worden et al., 2015) into the VND7-inducible

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Arabidopsis line. The 3xYFP-CSI1/POM2 can be seen as fluorescent foci that track together

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with the CSCs at the cell cortex along linear trajectories during primary wall synthesis

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(Worden et al., 2015). After induction of VND7, we observed a clear change in the cellular

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distribution of the 3xYFP-CSI1/POM2. Although the 3xYFP-CSI1/POM2 foci maintained

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linear movement, the pattern of movement changed following induction. The foci were

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initially evenly distributed across the plasma membrane; however, this pattern changed in

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favor of dense and regularly spaced banded patterns (Supplemental Movie 1).

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Cortical MTs change their distribution during the progression of xylem vessel

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production and form distinct banded or helical arrays (Watanabe et al., 2015), similar to what

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we observed for the 3xYFP-CSI1/POM2 foci. To see if the 3xYFP-CSI1/POM2 and MT re-

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distributions co-occurred during xylem vessel development we crossed a mCherry-TUA5

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expressing plant with the 3xYFP-CSI1/POM2 VND7-inducible plant, and analyzed the

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progeny. We found that the changes in the 3xYFP-CSI1/POM2 patterns co-occurred with the

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re-arrangement of the MT array during the transition from primary to secondary wall

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synthesis (Figures 2A to E), indicating that the CSI1/POM2 likely tracks with both primary

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and secondary wall CSCs.

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The primary wall CSCs typically track with a speed of about 250 nm/min (Paredez et

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al., 2006), and CSI1/POM2 proteins track together with these CSCs (Bringmann et al., 2012).

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However, the secondary wall CSCs track significantly faster than the primary wall CSCs

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(Watanabe et al., 2015), and if the CSI1/POM2 proteins are associated with the secondary

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wall CSCs one would anticipate an increase in speed of the CSI1/POM2s over time after

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VND7 induction. To test this, we measured the speed of the 3xYFP-CSI1/POM2 at early-,

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mid-, and late time points after VND7 induction. We selected these time points based on the

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MT re-organization status after induction (Supplemental Figure 2), and they roughly coincide

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with time points used in Watanabe et al. (2015). We found that the CSI1/POM2 proteins

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moved with a speed of approx. 266 ± 35 nm/min (1015 foci in 12 cells from 3 seedlings) in

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DMSO-treated cells and at 293 ± 98 nm/min (905 foci in 17 cells in 15 seedlings) in cells in

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the early stages of the secondary wall synthesis, i.e. where MTs still exhibited a primary wall-

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like pattern (Figures 2F to H). However, during the middle stages of secondary wall

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synthesis, i.e. where MTs formed diffuse bands, we observed a significant increase in

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CSI1/POM2 speed (422 ± 72 nm/min; 1391 foci in 18 cells from 15 seedlings). Once

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secondary cell wall synthesis had progressed to late stages, we found that the speed of the

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CSI1/POM2 proteins declined (211 ± 75 nm/min; 234 foci in 7 cells from 15 seedlings),

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possibly related to the initiation of programmed cell death. Notably, the secondary wall CSCs

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underwent a very similar transition in speeds during early-, mid-, and late secondary wall

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stages (Watanabe et al., 2015). These data indicate that the CSI1/POM2 may track with the

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secondary wall CesA proteins, similar to what has been shown for the primary wall CesAs

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(Gu et al., 2010). To test whether CSI1/POM2 can interact with secondary wall CesAs, we

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performed Bimolecular Fluorescence Complementation (BiFC) assays between the three

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secondary wall Arabidopsis CesAs and CSI1/POM2. We found that the proteins can interact

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when transiently expressed in tobacco epidermal leaf cells (Supplemental Figure 3). Hence,

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the CSI1/POM2 proteins behave similarly to the secondary wall CesAs and can interact with

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them.

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The CSI1/POM2s track with the secondary wall CesAs and are rapidly recruited to

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their sites of action

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The CSI1/POM2 and secondary wall CesAs behave similarly and can interact, suggesting

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that the proteins also track together during xylem vessel development. To test this, we 6

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generated plants expressing a mCherry-tagged CSI1/POM2 fusion protein under control of

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the CSI1/POM2 promoter and introgressed these plants with VND7-inducible lines

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expressing the YFP-CesA7 construct. The two fluorescent proteins showed similar behavior

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and closely co-localized throughout the different stages of VND7 induction (Figures 3A to

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D). These observations are supported by close inspections of kymographs from movies of the

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fluorescent proteins, where the tracking of the proteins coincided (Figure 3B).

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To assess the recruitment of CSI1/POM2 and CesA7 to their sites during secondary

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wall synthesis, we performed fluorescence recovery after photo-bleaching (FRAP)

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experiments. We first used the VND7-lines expressing mCherry-CSI1/POM2 and YFP-

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CesA7; however, the mCherry signal proved too weak to accurately assess fluorescence

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recovery. Instead, we used VND7-induced lines expressing either 3xYFP-CSI1/POM2 or

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YFP-CesA7, and counted the number of insertion events over time, which permitted

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measurement of the average insertion, or delivery, times (Supplemental Figures 4A to D).

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The 3xYFP-CSI1/POM2 signal rapidly re-populated the bleached area after FRAP

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(Supplemental Figures 4A, D; recovery time 32 ± 13 s, 46 bands in 7 cells in 3 seedlings),

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whereas the recovery of the YFP-CesA7 was significantly slower (106 ± 68 s, 32 bands in 6

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cells in 4 seedlings). We calculated the ratio of the recovery of the two fluorescently-labelled

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proteins to be 3.3 ± 2.5. As the secondary wall CSCs contain CesA4, CesA7 and CesA8,

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possibly in equal stoichiometry (Gonneau et al., 2014; Hill et al., 2014), it is likely that each

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CesA in the secondary wall CSC is associated with one CSI1/POM2 protein. However, it is

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important to note two things; firstly, the pom2-4 and irx3-4 mutations were not homozygous

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in the 3xYFP-CSI1/POM2 and the YFP-CesA7 lines, respectively. While one might assume

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that each CSC will contain both labelled and un-labelled CesA7, and thus that each CSC is

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tracked in our image analysis, it is possible that we under-estimate the numbers of

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CSI1/POM2s associated with each CSC. Secondly, the analyses were done on seedlings with

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either the 3xYFP-CSI1/POM2 or YFP-CesA7, which alone may introduce experimental

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differences. While we therefore favor a ratio between the CSI1/POM2 and CesA as 1:1 at a

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given secondary wall CSC, further experiments are needed to firmly corroborate this

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hypothesis.

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Optical flow analyses support global bi-directionality, but local uni-directionality, of the

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CSI1/POM2

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To assess, in more detail, the migratory patterns of CSI1/POM2 during xylem vessel

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development we analyzed the behavior of the 3xYFP-CSI1/POM2 using optical flow 7

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analyses (Supplemental Figures 5A to C). This analysis can examine the patterns of apparent

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motion and size of fluorescent objects. We false-colored the motion of fluorescent objects

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based on direction, i.e. movement to the left or right were colored purple and green,

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respectively (Figure 4A). We detected clear bi-directional movement of the 3xYFP-

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CSI1/POM2 objects in both DMSO and VND7-induced cells, i.e. the YFP-CSI1/POM2

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trajectories clearly overlapped along kymograph sections (Figure 4B), and the average optical

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flow images contained a significant number of white pixels (Figure 4B, C). However,

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domains of apparent uni-directional movement were significantly larger in the cells

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undergoing xylem differentiation (Figure 4D). These data indicate that the flow of the

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CSI1/POM2, and therefore most likely also the CSCs, is preferentially bi-directional on a

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cellular scale, but uni-directional on a local scale. Hence, in regions where one CSI1/POM2

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migrated in a defined direction, the majority of the associated CSI1/POM2s were likely to

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follow the same direction (Figures 4B, C). To see if the movement of the 3xYFP-CSI1/POM2

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foci depended on whether they were part of uni- or bi-directional domains, we measured the

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speeds of the foci from the different domains. We found that the CSI1/POM2 migrated with

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similar speeds independent of being part of a bi- or uni-directionally moving domain (Figure

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4E). These data indicate that the secondary wall cellulose is preferentially produced in one

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direction at any given sub-region of the cell wall bands.

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We observed many 3xYFP-CSI1/POM2 foci that first moved in one direction but

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suddenly stopped and changed direction (white arrows in Figure 4B). Such events were

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detected only in xylem vessel-differentiating cells and not in the DMSO-treated cells. These

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observations suggest that CSI1/POM2 can be transferred between CSCs moving in opposite

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directions during secondary wall synthesis, perhaps to support tight associations between the

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secondary wall CSCs and underlying MTs.

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Mutations in CSI1/POM2 cause mis-alignments of secondary wall CesAs and

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microtubules

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Lesions in CSI1/POM2 caused defects in the alignment of primary wall CesA trajectories and

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cortical MTs (Bringmann et al., 2012; Li et al., 2012). To investigate whether defects in

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CSI1/POM2 also affected the alignment of the secondary wall CesAs and the MTs, we

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generated YFP-CesA7 mCh-TUA5 dual-labelled VND7-inducible lines in wild-type or

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pom2-4 mutant backgrounds. The YFP-CesA7 trajectories co-aligned with cortical MTs

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during all stages of xylem vessel development in the wild-type background (Supplemental

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Figure 6). However, we observed clear defects in the alignment of CesA7 trajectories and 8

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MTs in pom2-4 mutant cells (Figures 5A to F; Supplemental Movie 2). Notably, substantial

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mis-alignment was observed only during early stages of secondary wall synthesis (Figure

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5D). These observations were corroborated by quantification of YFP-CesA7 and mCh-TUA5

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co-localization, revealing a significant reduction in CesA7 overlap with MTs during the early

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developmental stage, but not during subsequent stages, in pom2-4 as compared to wild-type

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(Figures 5E, F).

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To further assess whether defects in CSI1/POM2 influenced the behavior of the

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secondary wall CSC, we measured insertion rates and speeds of YFP-CesA7 in either VND7-

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induced wild-type or pom2-4 mutant backgrounds. We found that the insertion times of YFP-

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CesA7 in pom2-4 were not significantly different from wild-type (Supplemental Figures 4C,

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D; 84 ± 27 s, 39 bands in 6 cells in 5 seedlings). By contrast, we observed changes in the

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distribution of YFP-CesA7 speeds during the progression of secondary cell wall synthesis

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(Figure 5G). In the pom2-4 mutant background, YFP-CesA7 moved with higher speeds

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during early stages of secondary wall synthesis, but showed significantly slower speeds than

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wild-type during mid stages. In addition, during late secondary wall synthesis stages, the

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YFP-CesA7 speeds appeared less tightly controlled in the pom2-4 mutant as compared to

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wild-type. These findings indicate that CSI1/POM2 is involved in regulating the speed of

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secondary wall CesAs, possibly by maintaining the CesAs in close vicinity of the MTs.

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Xylem vessel cell wall patterns can be maintained in the absence of microtubules

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CSI1/POM2 is regarded as the component that guides CSCs along cortical MTs during

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primary wall synthesis (Bringmann et al., 2012; Li et al., 2012). The observation that the

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CSI1/POM2 is not essential for alignment of the CSCs and MTs during the mid- and late-

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stages of xylem vessel development indicated that these stages do not depend on MT-based

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guidance to maintain cell wall patterning. To test this conclusion, we first established time

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points when the MT array was re-organized during VND7-induced xylem vessel formation.

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In our hands, diffuse bands of MTs were not established until around 16 h after the VND7

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induction (Figure 6A), and the bands became progressively more condensed during the

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subsequent eight hours. To assess the influence of MTs on cell wall pattern maintenance we

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treated VND7-induced seedlings with the MT-depolymerizing drug oryzalin (Morejohn et al.,

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1987; 20 µM) at different time points after induction, and then investigated the ensuing wall

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patterns 48 h after VND7 induction. Seedlings treated with oryzalin eight hours after

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induction lacked cell wall bands entirely. By contrast, cell wall bands were evident in

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seedlings treated with oryzalin 16 and 24 h after VND7 induction (Figure 6B, third and fourth 9

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image from left). These wall bands were not as well defined and evenly spaced as the control

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seedlings (DMSO-treated; Figure 6B, left image). However, when comparing the wall

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patterns with the typical MT array organization after 16 and 24 h VND7 induction (Figure

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6A, third and fourth image), the 16 and 24 h wall bands showed very similar distributions

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(Figures 6A, B). In addition, the wall bands in the oryzalin-treated seedlings (treated 16 and

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24 h, and imaged at 48 h, after VND7 induction) were substantially more pronounced as

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compared to the wall patterns in seedlings at 24 h after VND7 induction (Figure 6B). These

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data indicate that the xylem vessel wall patterns become reinforced despite removal of the

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MT array.

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Secondary wall CesAs remain preferentially delivered to sites of secondary cell wall

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bands in absence of microtubules

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To determine the behavior of the CSCs and the CSI1/POM2 in the absence of MTs during the

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VND7 induction, we used the dual-labelled YFP-CesA7 mCherry-TUA5 and 3xYFP-

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CSI1/POM2 mCherry-TUA5 lines. We treated the seedlings with oryzalin for 4 h after 24 h

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VND7 induction, confirming effective MT de-polymerization, and assessed the behavior of

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the YFP-tagged proteins. While some YFP-CesA7 puncta clearly were not associated with

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distinct bands, many were, despite complete de-polymerization of MTs (Figure 6C). These

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observations were confirmed with fluorescence intensity values along transects from time

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average images (Figures 6C, D). Similar observations were made using the 3xYFP-

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CSI1/POM2 (Supplemental Figure 7), indicating that the proto-xylem vessels need MTs to

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establish the wall patterns, but that the patterns can be maintained in the absence of MTs.

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To investigate the dynamic behavior of the secondary wall CSCs in more detail, we

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first looked at the behavior of YFP-CesA7-containing Golgi bodies. We observed that the

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Golgi moved erratically at the cell cortex and that they preferentially associated with regions

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that coincided with microtubule bands (Supplemental Movie 3). We next investigated the

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behavior of the Golgi in cells where microtubules had been depolymerized by oryzalin

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treatment. Golgi followed very similar patterns of movement, i.e. they preferentially

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populated regions where microtubule bands had been before the oryzalin treatment (Figures

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6E to G; Supplemental Movie 4). To confirm these observations, we analysed the number of

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Golgi localised to wall bands vs. gaps using TrackMate. We found that 85% ± 16% (mean +/-

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S.D., N = 7 bands in 3 cells) of the Golgi were localised beneath wall bands, with the rest

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localised to gaps, in the presence of microtubules. After oryzalin treatment 75% ± 18% of the

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Golgi were localised beneath wall bands. This difference was not significant (p=0.39, 10

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Welch’s unpaired t-test) indicating that Golgi movement is independent of microtubule

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bands. Golgi positions typically correspond to sites of delivery of CesAs (Crowell et al.,

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2009; Sampathkumar et al., 2013). To see if CesAs were inserted to the plasma membrane

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mainly above areas with Golgi movement, we applied FRAP and assessed how the YFP-

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CesA7 signal re-populated the bleached area. Indeed, the CesAs were preferentially inserted

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at regions where the bulk of Golgi was evident and thus in proximity of the wall bands both

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in presence and absence of microtubules (Figures 6E to G). To investigate if the delivered

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CesAs moved in any direction after delivery, or if they followed the tracks of previous CesAs

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we analyzed the CesA behavior at the plasma membrane. While we found that the YFP-

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CesA7 foci moved slower in the absence of microtubules as compared to cells with

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microtubules (Figures 6H, I), the majority of CesAs moved parallel to the cell wall bands. In

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all, 88% ± 4% and 93% ± 8% of the newly inserted CesAs moved parallel to cell wall bands

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before and after oryzalin treatment, respectively (means ± SD, 101 and 35 CesAs in 3 cells;

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Figure 6F). Given the Golgi behavior, our data indicate that the cellular regions where cell

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wall bands are made are different in their molecular composition as compared to inter-band

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regions.

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Defects in CSI1/POM2 affect patterning of cell walls in rice xylem vessels

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The importance of CSI1/POM2 in cellulose synthesis has been supported by data from only

353

Arabidopsis. To see if the protein is also important for secondary wall synthesis in other plant

354

species, we investigated the role of CSI1/POM2 in xylem vessel wall formation in rice.

355

CSI1/POM2 was in part discovered based on co-expression of the corresponding gene with

356

the primary wall CesA genes in Arabidopsis (Gu et al., 2010). We therefore explored what

357

rice CSI1/POM2 homolog displayed the closest co-expression with the rice primary and

358

secondary wall CesAs using FamNet (Ruprecht et al., 2016). We found that the most likely

359

candidate for this function was Os06g11990, which we referred to as CSI-like 1 (CSIL1;

360

Supplemental Figure 8A). These data were corroborated by phylogenetic analyses, which

361

revealed that the rice CSIL1 was closely related to the Arabidopsis CSI1/POM2, and through

362

expression analyses that showed ubiquitous expression of the gene and very low expression

363

of the other CSIL genes (Supplemental Figures 8B to D, Supplemental Data set 1). To assess

364

whether this protein affects rice growth, we generated RNAi-mediated suppression constructs

365

to down-regulate CSIL1. Several independent homozygous T3 progeny of the transformants

366

had substantially decreased CSI1L transcript abundance, as estimated by quantitative RT-

367

PCR (Figure 7A) and showed stunted growth with reduced cellulose content (Figures 7B, C). 11

368

Notably, when estimating the secondary cell wall (SCW) thickness, we found that the CSIL1

369

RNAi plants had considerably thinner walls as compared to control plants (Figures 7D, E).

370

While these effects were more pronounced than what we observed in Arabidopsis, they

371

clearly support a function of CSIL1 in secondary wall synthesis in rice. In addition, when we

372

assessed the xylem vessel wall patterns, we found that the spacing between the bands was

373

significantly changed (Figures 7F, G). These changes were in close agreement with the

374

phenotypes we observed in the Arabidopsis pom2-4 and pom2-8 mutants (Figure 1).

375

To assess if the CSIL1 can also interact with the rice secondary wall CesAs we

376

performed split-luciferase assays of the rice secondary wall CesAs, i.e. CesA4, CesA7 and

377

CesA9, and the CSIL1. All the rice secondary wall CesAs could interact with the CSIL1

378

(Figure 7J, Supplemental Figure 8E), corroborating a function of CSIL1 in rice secondary

379

wall cellulose production. In addition, transient co-infiltration of mRFP-CSIL1 and GFP-

380

CesA4 into N. benthamiana leaves revealed tight co-localization of the proteins at plasma

381

membrane focal planes (Figures 7H, I). The patterns of co-localization were reminiscent of

382

cortical MTs, supporting a related function of the rice CSIL1 and the Arabidopsis

383

CSI1/POM2. Hence, we conclude that CSI1/POM2 also contributes to xylem vessel patterns

384

in rice.

385 386

12

387

DISCUSSION

388

Cell wall patterning has been attributed to MT-based guidance of CSCs (Oda and Fukuda,

389

2013; Schneider et al., 2016). While the guiding principles have been largely resolved for

390

primary cell wall cellulose synthesis, the corresponding mechanisms for secondary wall

391

deposition have remained ill defined. We show that secondary wall patterning depends on

392

MT-based cell wall deposition. However, once the wall patterns are established, they can also

393

be sustained in the absence of MTs, as hypothesized in Zinnia elegans cell suspensions

394

(Roberts et al., 2004). The re-organization of the MT array therefore represents a critical

395

initial establishment phase for the xylem vessel bands to form, whereas the patterns can be

396

maintained in the absence of MTs during the subsequent phases.

397

Several MT-associated proteins have been implicated in the MT re-organization

398

during xylem vessel development, including MAP65-1, AIR9, MAP70-1 and MAP70-5

399

(Pesquet et al., 2010; Derbyshire et al., 2015). These proteins contribute to the MT-

400

bundling/stabilization, and are important to achieve the MT array re-arrangements during

401

secondary wall synthesis. In addition, several small GTPases, MICROTUBULE

402

DEPLETION DOMAIN 1 (MIDD1), a KINESIN 13A, and a recently described member of

403

the IQD family (IQD13) are involved in depleting MTs from the areas between thickenings

404

(Oda and Fukuda, 2012, Sugiyama et al., 2017). Nevertheless, the mechanism for how the

405

MTs guide cellulose synthesis during this important developmental process has remained

406

elusive. Our results indicate that CSI1/POM2 is necessary for MT-based CSC guidance

407

during the initial phase of xylem vessel development, but that it is not needed during the

408

subsequent stages. Time-course experiments using oryzalin corroborate that cell wall

409

patterns, and the tracking of CSCs along defined bands, can be maintained also in the absence

410

of MTs. These data indicate that other mechanisms, perhaps cell wall-mediated CSC

411

guidance, may play significant roles during these stages. It is plausible that xylans and/or

412

other cell wall constituents that are deposited along the MT bands may serve this function in

413

the absence of CSI1/POM2. Computational modelling and NMR experiments suggest a tight

414

interplay between xylans and cellulose microfibrils (Busse-Wicher et al., 2014; Simmons et

415

al., 2016). Such interactions could influence the direction of the CSCs and therefore cause

416

them to successively align along the secondary wall bands. Apart from potential cell wall

417

polymers directing cellulose synthesis, the observation that Golgi movement and CesA

418

delivery are different at regions that underlie cell wall bands indicate that other cellular

419

features may also influence the cell wall band progression. We speculate that the membrane

420

environment is different in these regions as compared to membrane regions that lie between 13

421

the bands, and that these differences influence the movement of the Golgi bodies and thus the

422

delivery of CesAs. Another possibility is that the movement of the Golgi is restricted due to

423

physical constraints. The secondary wall bands lead to deformation of the plasma membrane,

424

i.e. the membrane is slightly indented below the bands. These deformations could make it

425

difficult for Golgi to pass through these regions and perhaps could trap Golgi beneath the

426

bands. While there is much left to explore about this process, we find it unlikely that the

427

maintenance of the band patterning is solely due to the cell wall polymers directing cellulose

428

synthesis.

429

The primary wall CSCs typically track with a uniform speed and bi-directionality

430

along cortical MTs (Paredez et al., 2006). The speed of the primary wall CSCs is reliant on

431

CSI1/POM2 function, as CSC speeds were significantly reduced in csi1/pom2 mutants (Gu et

432

al., 2010). During the secondary wall synthesis, we also see clear alterations in CesA7 speeds

433

in pom2-4; however, these changes appear to be largely due to a wider spread of speeds

434

rather than a uniform reduction, and are tied to particular developmental stages of xylem

435

vessel development. The increased variance in the CSC speeds was primarily observed

436

during the mid-stages of secondary wall synthesis (Figure 5G). While we observed major

437

mis-alignment between the MTs and CesA7 trajectories only during the early stages, it is

438

possible that the lack of direct engagement of the CSCs with the MTs causes difficulties in

439

maintaining the speeds. It is worth highlighting that although we observed clear secondary

440

wall bands in the csi1/pom2 lines, these bands were less well ordered as compared to the wild

441

type. We speculate that CSI1/POM2 proteins provide a feedback function to the formation of

442

MT bands and that this may be compromised in the csi1/pom2 lines, which in turn may affect

443

the final cell wall band patterns. This is in line with observations during primary wall

444

synthesis where MT organization is perturbed when CSI1/POM2 is mutated (Bringmann et

445

al., 2012; Landrein et al., 2013). If the CSCs are indeed also guided by cell wall components

446

and membrane environment during this development stage, as discussed above, it is plausible

447

that such guidance is not optimal and that it can manifest in changes in the quality and

448

quantity of cellulose that is produced. These data are in agreement with our recorded changes

449

in the MFA, cellulose microfibril crystallinity and amorphous cellulose in the pom2-4 mutant.

450

The CSI1/POM2 speeds changed during the different stages of xylem vessel

451

development. For example, the speeds significantly increased during the mid-stages of

452

transition (Figure 2H). These data are very similar to what has independently been reported

453

for the secondary wall CesA7 (Watanabe et al., 2015), but contrast those of Li et al. (2016b).

454

Assuming that the speed of the CSCs represents catalytic activity, these findings support a 14

455

scenario in which an increase in speed of tracking and CesA abundance leads to a major

456

boost in cellulose synthesis, which is compatible with the rapid development and subsequent

457

death of the xylem vessels. Li et al. (2016b) also studied secondary wall CesA behavior in the

458

VND7-inducible system and concluded that the CSCs moved uni-directionally as “swarms”

459

(referred to as “directionally coherent movement”; Li et al., 2016b) during xylem vessel

460

development. Our data support this report, but it is important to note that the uni-directional

461

movement apparent in CSI1/POM2 was observed both during primary wall synthesis

462

(DMSO-treatment; Figure 4) and xylem vessel development, and appears to depend on local

463

vs. global cell wall synthesis.

464

In summary, CSI1/POM2 directs xylem vessel patterning by coordinating the

465

secondary wall CSCs and MTs during the transition from primary to secondary wall

466

synthesis. However, the banding patterns can largely be maintained in absence of

467

CSI1/POM2 and MTs during later stages of development. We therefore conclude that the

468

wall patterning during proto-xylem development is initiated and more importantly sustained

469

by two complementary mechanisms.

470 471 472

MATERIALS AND METHODS

473

More detailed descriptions of some procedures are provided in Supplemental Methods.

474 475

Plant Material

476

We used the previously described Arabidopsis thaliana lines pom2-4 and pom2-8/csi1-1

477

(SALK_136239; Bringmann et al., 2012). To generate multiple marker lines in the VND7

478

background, we crossed seeds of pom2-4 and native promoter-driven triple (3x)YFP-

479

CSI1/POM2 (Worden et al., 2015) into the VND7-inducible Arabidopsis line

480

proCaMV35s::VND7::VP16::GR (Yamaguchi et al., 2010). The F3 progeny was used for

481

analysis. The pom2-4 mutant was used as the main allele, as it produces seeds more readily

482

than some of the more severe csi1/pom2 lines. Note that the pom2-4 was generated from a T-

483

DNA population between Nossen and Columbia. We have back-crossed the pom2-4

484

extensively to Col-0 and we used segregating progeny from crosses of pom2-4 and different

485

markers to assure the best possible genetic homogeneity between samples. To generate dual-

486

labelled plants for MTs we crossed 3xYFP-CSI1/POM2 in the VND7 background into

487

mCherry-TUA5 (Gutierrez et al., 2009) and used the F2 progeny of that cross. To visualize

488

secondary wall CesAs we crossed YFP-CesA7 in the irx3-4 background (Watanabe et al., 15

489

2015) into the wild-type and pom2-4-mutated mCherry-TUA5-VND7 background. We used

490

the F2 generation for experiments. We confirmed the homozygosity of the pom2-4 mutation

491

by growing seedlings on solid (0.8% agar) half-strength Murashige and Skoog (MS) medium

492

(pH 5.7) supplemented with 5% sucrose allowing the identification of the obvious stunted

493

root phenotype, and confirmed via PCR (Supplemental Table 1). We further checked for the

494

presence of YFP- and mCherry-markers using a fluorescence stereomicroscope prior to

495

further treatment.

496

For generation of CSIL1-RNAi plants, the targeted fragments were amplified from the

497

cDNA of rice CSIL1, and inserted into PKANNIBAL vectors (see below). The construct was

498

transfected into Agrobacterium tumefaciens EHA105 and introduced into the wild-type

499

variety Nipponbare.

500 501

Plant Growth Conditions and Treatments

502

Arabidopsis plants were germinated and grown essentially as described by Liu et al. (2016).

503

More specifically, seeds of the VND7-inducible Arabidopsis lines described above were

504

surface-sterilized by washing for 10 minutes in 1.25% sodium-hypochlorite solution

505

supplemented with 0.05% Tween20. Sterilized seeds were washed excessively with sterile

506

water. Seeds were plated on solid (0.8% agar) half-strength MS medium (pH 5.7)

507

supplemented with 1% sucrose for normal growth and 5% sucrose for pom2-4 genotyping

508

purposes, respectively. Plates were stratified for at least two days in a dark cold room (4 °C).

509

Germination was triggered by exposing plates for eight hours to light (100 µE m-2 s-1).

510

Subsequently, the plates were wrapped with aluminum foil and placed vertically in a growth

511

room air-conditioned to 60% relative humidity, and 21 °C temperatures. Seedlings used for

512

spinning disc confocal microscopy were grown for three days in the dark, transferred to 24-

513

well plates containing DMSO (control) or 10 to 100 µM dexamethasone (induction) to induce

514

VND7. Subsequently, plates were wrapped with aluminum foil and placed on a slowly

515

rotating orbital shaker in the growth room.

516

The plants used for determination of cell wall alterations were sterilized for 3 minutes

517

in 70 % ethanol followed by 10 minutes in 10 % bleach, then rinsed six times in sterile

518

distilled water and plated on solid (0.8% agar) half-strength MS plates and stratified for two

519

days in a dark cold room (4 °C) before being incubated at 21˚C at a 16/8 hour light/dark

520

cycle. Ten-day-old seedlings were transferred to soil and grown under the same conditions

521

for about 9 weeks through maturity to full senescence.

16

522 523

Arabidopsis cell suspension cultures were generated, and tracheary element formation induced, as previously described (Pesquet et al., 2010).

524

Rice plants (Oryza sativa L.), including the wild-type plants and CSIL1-RNAi plants,

525

were grown in experimental fields at the Institute of Genetics and Developmental Biology in

526

Beijing and in Linshui, Hainan province during the natural growing seasons.

527

Nicotiana benthamiana plants were grown in soil in a glasshouse with continuous

528

cool white fluorescent lights (100 µE m-2 s-1) and natural daylight at 20–26°C, as previously

529

described (Lampugnani et al., 2016).

530 531

Live Cell Imaging

532

Imaging was done essentially as described in Liu et al. (2016). Seedlings were observed

533

under the microscope between 10 and 30 hours after VND7 induction. Induced 3xYFP-

534

CSI1/POM2 seedlings were imaged using the CSU-X1 spinning disk head (Yokogawa)

535

mounted to an inverted Nikon Ti-E microscope equipped with a 100x oil-immersion

536

objective (Plan Apo TIRF, NA 1.45). Fluorescence detection was achieved using an Evolve

537

EM-CCD camera (Photometrics Technology, USA). Induced YFP-CesA7 seedlings were

538

imaged using the CSU-W1 spinning disk head (Yokogawa) mounted to an inverted Nikon Ti-

539

E microscope equipped with a 100x oil-immersion objective (Apo TIRF, NA 1.49).

540

Fluorescence detection was achieved using a deep-cooled iXon Ultra 888 EM-CCD (Andor

541

Technology, Northern Ireland). Both setups were controlled via PC using MetaMorph

542

(Molecular Devices, USA). Photo-bleaching was achieved using either the iLas laser

543

illumination system (Roper Scientific, France) or the Andor FRAPPA scanning instrument.

544

Seedlings were mounted on 1.5 grade glass coverslips and covered by 1 mm thick

545

agarose pads made from water supplemented with 1% agarose. We imaged 3xYFP-

546

CSI1/POM2, mCherry-TUA5, and YFP-CesA7 using time-lapse recordings with typical

547

exposure times between 200-400 ms, time-intervals of 10 seconds and total durations

548

between 5-10 minutes. Fluorescence recovery was recorded in intervals of 2 to 5 seconds for

549

3xYFP-CSI1/POM2 and 10 seconds for YFP-CesA7.

550

For analysis of co-localization of rice CesA4-CSIL1, Agrobacterium tumefaciens

551

EHA105 harboring GFP-CesA4 and mRFP-CSIL1 were co-injected into the lower epidermis

552

of 4-week-old Nicotiana benthamiana leaves. After cultivation for two more days, the leaves

553

were observed with oil immersed objective on the spinning-disc confocal microscope

554

(PerkinElmer UltarVIEW VoX). To obtain the GFP and mRFP fluorescence images, the 488

17

555

nm and 561 nm lines of laser were used for excitation, and emission was detected at 500–540

556

nm and 600–640 nm, separately.

557 558

Scanning Electron Microscopy

559

For xylem defect analysis, the first internodes of 8-week-old wild-type, pom2-4, and pom2-8

560

(csi1-1) mutant plants, were cut into longitudinal sections and immediately fixed in 2.5 %

561

glutaraldehyde in PBS buffer for 30 minutes. Sections were washed three times in PBS and

562

subsequently three times in water. Dehydration was achieved by washing the sections for

563

minimum 1 hour each in an ethanol series from 10 % to 100 % in 10 % steps. After several

564

washes with 100 % ethanol, critical point drying was performed and the dried samples were

565

gold-coated. Examination of the samples was performed using an XL30 field-emission SEM

566

from Phillips.

567

The 2nd internodes of mature wild-type and CSIL1-RNAi plants were fixed in

568

4 % paraformaldehyde (Sigma). To view the wall thickness of sclerenchyma cells, the

569

internodes were transversely cut to expose the epidermal sclerenchyma cells. To observe the

570

secondary pattern of vessel cells, the internodes were longitudinally cut under the stereoscope.

571

After critical-point drying, the samples were sprayed with gold particles and observed with a

572

scanning electron microscope (S-3000N, Hitachi).

573 574

Cell Wall Staining

575

To label the secondary walls in VND7-induced wild-type and pom2-4 mutants, DirectRed23

576

(Anderson et al., 2010; Sigma) was added to a final concentration of 0.06% to 6-well plates

577

containing 3-day-old seedlings 24 hours after induction. The samples were washed with

578

ultrapure water to reduce the amount of unbound dye. Subsequently, samples were observed

579

under the Spinning Disk microscope by recording z-stacks using a 561 nm laser and

580

610/40 nm emission filters.

581 582

Image Analyses

583

The velocity of CSI1 foci was measured using the open-source software FIESTA (Ruhnow et

584

al., 2011). Briefly, the velocity of moving foci is determined by measuring their slope in

585

kymograph projections. We measured 1015 trajectories for non-induced cells and 905, 1391,

586

and 367 trajectories for induced cells in early, mid, and late stages of the secondary wall

587

program, respectively. Co-localization of CSI1 with MTs and CesA7, respectively, was 18

588

measured using the JaCoP plugin of Fiji. To increase the reliability of the co-localization

589

measurements, we used a dual approach of measuring Pearson’s and Mander’s coefficients.

590

Furthermore, we used Costes randomization to validate the significance of the determined

591

Pearson’s coefficients. Costes-randomized image series always had a Pearson’s coefficient of

592

at least a factor of 50 lower than the original image series.

593

To quantify the insertion rate of 3xYFP-CSI1 and YFP-CesA7 after photo bleaching,

594

we used the ThunderSTORM plugin of Fiji to detect the appearance of foci in the bleached

595

areas. We analyzed the recovery in areas slightly smaller than the bleached area to avoid

596

migrating complexes in the plasma membrane to be included in the recovery signal. We

597

plotted the number of detected foci over time and analyzed the recovery using a mono-

598

exponential growth model (reaction-limited case).

599

The misalignment between CesA trajectories and MTs was measured in dual-color

600

average projections of the time series using Fiji. We measured the angle of short stretches of

601

clearly visible CesA7 trajectories and compared them to the angle of the underlying MTs. For

602

each cell at least 10 trajectory-MT pairs were measured.

603

Orientation and Spacing of Secondary Wall Bands in VND7-Induced Seedlings were

604

measured using Fiji. Z-stacks were smoothed and average-projected using inbuilt Fiji plugins.

605

Subsequently, individual cells were cropped and aligned with the growth axis of the seedling.

606

A total of 136 and 132 VND7-induced cells were captured for wild-type and the pom2-4

607

mutant, respectively. We quantified the average orientation of secondary wall bands and the

608

variability of band orientations within each cell, termed dispersion, using the Fiji plugin

609

‘Directionality’ with default settings. Band spacing was analyzed using a custom-made

610

Matlab (Mathworks, USA) program. Briefly, the program displayed the intensity profile

611

along the long axis of the cell and a graphical user-interface subsequently allowed for the

612

determination of band positions in a point-and-click manner.

613 614

Optical Flow Analysis

615

The optical flow was analyzed using the Fiji plugin PIV analyzer using 4-by-4 pixel

616

averaging, interpolation and a mask of 0.1. The image series were pre-processed using

617

subtract background (50 pixel sliding paraboloid) and 4-frame walking averaging. The

618

resulting optical flow image series was average projected to obtain images displaying the

619

mean optical flow of intensity. The direction of the optical flow was determined using Fiji by

620

decomposing the mean optical flow images into Hue (H), Saturation (S) and Brightness (B)

621

with the following thresholds: for movement to the right (H between 34 and 94, S between 50 19

622

and 255, B between 1 and 255), for movement to the left (H between 161 and 221, S between

623

50 and 255, B between 1 and 255), for movement into both directions (H between 0 and 255,

624

S between 0 and 50, B between 1 and 255).

625 626

Biochemical Analyses

627

The microfibril angle (MFA) of at least 18 Arabidopsis stems from VND7 and the pom2-4

628

mutant in VND7 were measured using an X-ray diffraction technique (Ukrainetz et al., 2008).

629

The bottom 3 cm of mature, senesced plant stems were used for analysis. The 002 diffraction

630

spectra of each stem were screened for T-value distribution and symmetry on a Bruker D8

631

discover X-ray diffraction unit equipped with an area array detector (GADDS). Wide-angle

632

diffraction was used in the transmission mode, and measurements were made with CuKα1

633

radiation (λ = 1.54 Å). The X-ray source was fit with a 0.5 mm collimator and a GADDS

634

detector collected the scattered photons. The X-ray source and the detector were both set at a

635

theta angle of 0°. The diffraction data was integrated using GADDS software and further

636

analyzed to estimate MFA values.

637

Cell wall crystallinity was determined on the same stems used for measuring MFA,

638

using the same X-ray unit and parameters as the MFA measurements, except the source theta

639

was set at 17°. The diffraction data were integrated using GADDS software and the output

640

data further analyzed using a crystallinity calculation program based on the Vonk method

641

(Vonk, 1973).

642

Cellulose content: After X-ray data collection was complete the same stems were then

643

pooled by genotype and ground on a Thomas Wiley Mini Mill to pass through a #60 mesh

644

(250 µm). The powdered sample was then dried for 24 hours at 50 °C and 15 mg of tissue

645

was weighed into each pre-weighed tube. At least three technical replicates were done on

646

each pooled genotype. First, the Alcohol Insoluble Residue (AIR) was prepared as described

647

by Pattathil et al. (2012). The AIR was then subjected to a series of extractions in a procedure

648

modified from the AIR fractionation method also described by Pattathil et al. (2012). The

649

modifications involved completing the chlorite extraction first as well as the removal of both

650

the 1 M and the post-chlorite 4 M potassium hydroxide extractions. The resulting cellulose

651

residue was then pre-dried in a vacuum centrifuge and finished in a 50 °C oven for 48 hours

652

before the final weights were measured.

653

Degree of polymerization: The resulting cellulose was dissolved at 5 mg/mL in 9 %

654

Lithium Chloride (LiCl)/ N,N-Dimethylacetamide (DMAc) through a 4-step solvent

655

exchange of nanopure water, anhydrous ethanol, DMAc and 9 % LiCl/DMAc. Once 20

656

dissolved, the samples were diluted to 0.5 mg/mL cellulose in 0.9 % LiCl/DMAc and each

657

was separated on an Agilent 1100 SEC system containing Waters Styragel HR4 and HR6

658

columns coupled to a Wyatt Dawn Heleos II Light Scattering Detector. The average

659

molecular weight was calculated from the output using Wyatt’s Astra 6 software before

660

converting to degree of polymerization.

661 662 663

Expression and Phylogenetic Analyses

664

Rice CSI1 homologous genes were identified based on the annotation of the rice genome

665

database (Rice Genome Annotation Project, http://rice.plantbiology.msu.edu/). The

666

phylogenetic tree of CSI1 and its like proteins in rice and Arabidopsis was generated using

667

Maximum Likelihood with the MEGA5 software with 1000 bootstrap replications

668

(Supplemental Data File 1; Tamura et al., 2011). The spatio-temporal expression profiles of

669

rice

670

(http://ricexpro.dna.affrc.go.jp/).

CSIL1

was

from

the

expression

data

in

RiceXPro

database

671

To examine the expression of CSIL1 in the wild-type and transgenic plants, total RNA

672

was extracted from young internodes using the Plant RNA Purification Reagent (Invitrogen),

673

complementary DNA was synthesized from RNA using the Reverse Transcription system kit

674

(Takara). The expression level of CSIL1 was examined by qPCR with a CFX96 Real-time

675

System (Bio-Rad) using rice HNR as internal control. The primers for the RNAi construct

676

and qPCR analyses are listed in Supplemental Table 1.

677 678

Constructs

679

CSIL1-RNAi construct was generated by amplifying CSIL1 from a rice cDNA library, and the

680

cDNA was inserted into a PKANNIBAL vector (Wesley et al., 2001) using BamHI and XbaI.

681

The construct was transformed into the wild-type varieties Nipponbare ecotype using

682

Agrobacterium tumefaciens. The expression level of CSIL1 was quantified by qPCR with a

683

CFX96 Real-time System (Bio-Rad). Coding sequences of Arabidopsis EH1, CSI1/POM2,

684

CesA4, CesA7, CesA8 were amplified from cDNA and cloned into pAMON and pSUR using

685

the Gibson assembly method to generate N-terminal fusions to VN155 (I152L) and VC155,

686

respectively (Lee et al., 2014), for BiFC analyses (see below). All primers are listed in

687

Supplemental Table 1.

688

Transgenic cell suspensions were produced by co-culture with Agrobacterium

689

transformed with 35S:CSI1-RNAi construct (Derbyshire et al., 2015) as previously described 21

690

by Pesquet et al. (2010). Expression levels of CSI1 were measured using RT-qPCR on 5

691

independent biological repeats (primer sequences and method described in Derbyshire et al.,

692

2015) and expressed as percentage of CSI1-to-UBIQUITIN gene ratio. Down-regulated lines

693

showed a residual CSI1 expression of 57 ± 17 % (mean ± S.D.) as compared to wild-type

694

cells (100 ± 13 %, p < 0.005, Welch’s unpaired t-test). The expression of the different CSIs,

695

primary, and secondary CesAs during the TE differentiation time-course have been checked

696

using macro-array data (GEO GSE73146, Derbyshire et al., 2015).

697 698

Interaction Analyses

699

Coding sequences of EH1, CSI1/POM2, CesA4, CesA7, and CesA8 were amplified from

700

cDNA using the primers defined in Supplemental Table 1 and cloned into linearized BiFC

701

vectors pURIL (GENE-V(I152L)N), pDOX (GENE-VC), pAMON (V(I152L)N-GENE) or

702

pSUR (VC-GENE) using the Gibson assembly method as previously described (Lampugnani

703

et al, 2016). The pURIL and pDOX were linearized using KpnI and SfoI, while pAMON and

704

pSUR were linearized using BamHI and SfoI. EH1 was cloned into pURIL and pDOX while

705

CSI1/POM2, CesA4, CesA7, and CesA8 were cloned into pAMON and pSUR to generate C-

706

terminal and N-terminal fusions respectively. Constructs were introduced into Agrobacterium

707

and combinations of BiFC constructs, together with Agrobacterium strains carrying

708

35S::CFP-N7 (Kaplan-Levy et al, 2014) and P19 (Voinnet et al, 2003), were introduced into

709

N. benthamiana leaves by infiltration following the procedure described in Zhang et al.,

710

(2016). Leaves were examined for fluorescence 3 days post-infiltration, on an inverted Nikon

711

Ti-E microscope equipped with a CSU-W1 spinning disk head (Yokogawa). Detection

712

occurred using a 100x oil-immersion objective (Apo TIRF, NA 1.49) and an iXon Ultra 888

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EM-CCD (Andor Technology, Northern Ireland). All BIFC combinations were imaged under

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the same conditions. Specifically, a 445 nm laser line was used to excite CFP, while a 515 nm

715

laser line was used to excite YFP. Emissions were detected with 470/40 and 535/30 band pass

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filters. Z-stacks of images were collected using exposure times of 100 ms.

717

The split-luciferase complementation assay was performed as described (Chen et al.,

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2008). In brief, the cDNA of CSIL1, CESA4, CESA7 and CESA9 were amplified

719

(Supplemental Table 1) and inserted into the binary vectors for expression fused with N- or

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C-terminal luciferase. The resulting constructs were transfected into Agrobacterium

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tumefaciens strain C58 and infiltrated with the leaves of 4-week-old Nicotiana benthamiana.

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Interaction was determined based on the fluorescent signal intensity harvested by IndiGO

723

software. 22

724 725 726

Accession Numbers

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CSI1/POM2: At2g22125, CesA4: At5g44030, CesA6: At5g64740, CesA7: At5g17420,

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CesA8: At4g18780, VND7: At1g71930, TUA5 At5g19780, EH1: At1g20760, OsCSIL1:

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Os06g11990, OsCesA4: Os01g54620, OsCesA7: Os10g32980, OsCesA9, Os09g25490.

730

Sequence data from this article can be found in the Arabidopsis Genome Initiative or

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GenBank/EMBL databases under the following accession numbers: GEO GSE73146.

732 733

Supplemental Data

734

Supplemental Figure 1. Defects in CSI1/POM2 Cause Aberrant Secondary Wall Patterns.

735

Supplemental Figure 2. Representative Images of Primary Wall Synthesis and Different

736

Stages (early, mid, and late) of Xylem Vessel Development.

737

Supplemental Figure 3. BiFC Assay Demonstrating Interactions between CSI1/POM2 and

738

Secondary wall CesA4, CesA7, and CesA8 Transiently Expressed in Epidermal Cells of N.

739

benthamiana Leaves.

740

Supplemental Figure 4. CSI1/POM2 Recovers More Quickly after Photobleaching than

741

CesA7 during Xylem Vessel Formation.

742

Supplemental Figure 5. Schematic Workflow of Optical Flow Analyses.

743

Supplemental Figure 6. Secondary Wall CesA7 Tracks along Microtubules throughout all

744

Stages of Xylem Vessel Development in Wild-type Background, but not in the pom2-4

745

Mutant.

746

Supplemental Figure 7. YFP-CSI1/POM2 Can Maintain Tracks along Bands in the Absence

747

of MTs.

748

Supplemental Figure 8. Rice CSIL1 Is a Homolog of CSI1/POM2 and Can Interact with

749

Secondary Wall Rice CesAs.

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Supplemental Table 1. Primers used for BiFC constructs.

751

Supplemental Movie 1. Cellular distribution of 3xYFP-CSI1/POM2 in non-induced cells

752

and early, mid, and late stages of secondary wall formation.

753

Supplemental Movie 2. YFP-CesA7 trajectories are not aligned with cortical MTs in the

754

pom2-4 mutant during early stages of secondary wall formation.

755

Supplemental Movie 3. YFP-CesA7 quickly recycles at MT bands after fluorescence photo

756

bleaching (FRAP).

23

757

Supplemental Movie 4. YFP-CesA7 quickly recycles to sites of secondary wall formation

758

also in the absence of MTs.

759

Supplemental Data set 1. Multiple protein sequence alignment of CSI proteins in

760

Arabidopsis and rice.

761

Supplemental Data Set 2. ANOVA Tables.

762 763 764

ACKNOWLEDGEMENTS

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S.P was funded by a R@MAP Professorship at University of Melbourne. This work was in

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part supported by an ARC Discovery grant (DP150103495), a Future Fellowship grant

767

(FT160100218), and the National Natural Science Foundation of China (Grants 31530051).

768

SDM acknowledges funding from the NSERC Discovery program. We thank Prof. Taku

769

Demura for sharing the VND7-line.

770 771

24

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FIGURE LEGENDS

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Figure 1 | Defects in CSI1/POM2 cause aberrant xylem vessel patterns. (A and B) SEM

775

graphs of longitudinal sections of mature wild-type stems. Exposed (A) and transected (B)

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xylem vessels of wild-type plants, and pom2-4 and pom2-8 (csi1-1) mutants. Scale bar =

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10 µm. (C) Band-to-band orientations in pom2-4 (16 cells in 6 seedlings) and pom2-8 (44

778

cells in 6 seedlings) compared to wild-type xylem (27 cells in 6 seedlings) obtained from the

779

images in (A) and (B). (D and E) S4B-staining of cellulose in VND7-induced hypocotyls.

780

Dotted lines indicate highly ordered bands in the secondary walls of wild-type cells (D) and

781

irregular bands in pom2-4 mutant cells (E). Scale bar = 5 μm. (F) Distribution of the average

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band orientations (yellow lines in A, B). (G) The spread of band orientations within

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individual cells of induced pom2-4 cells (602 bands in 115 cells in 5 seedlings) compared to

784

wild-type cells (824 bands in 132 cells in 5 seedlings). (H) Secondary wall band spacing in

785

the pom2-4 mutant compared to wild-type cells. (I) Microfibril angle (MFA, relative to

786

growth axis) in the pom2-4 mutant compared to wild-type. (J) Cell wall crystallinity in the

787

pom2-4 mutant compared to wild-type. (K) Degree of polymerization (DP) in the pom2-4

788

mutant compared to wild-type. (L) Cellulose content (%-fraction of dry weight) in the pom2-

789

4 mutant compared to wild-type. All measurements (I to L) were done on ground stems of

790

10-week-old, fully-senesced plants grown in 16-hour light / 8-hour dark conditions. Data are

791

means ± S.D. Statistical significance was tested by Welch’s unpaired t-test, * p < 0.05, ** p