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. 2018 Aug 20;178(2):641–653. doi: 10.1104/pp.18.00498

Synaptotagmin-Associated Endoplasmic Reticulum-Plasma Membrane Contact Sites Are Localized to Immobile ER Tubules

Kazuya Ishikawa a,1,2, Kentaro Tamura a,3, Haruko Ueda b, Yoko Ito c,d, Akihiko Nakano c,e, Ikuko Hara-Nishimura b, Tomoo Shimada a,2
PMCID: PMC6181054  PMID: 30126867

SYNAPTOTAGMIN1-associated endoplasmic reticulum (ER)-plasma membrane contact sites are localized to immobile ER tubules in Arabidopsis thaliana and contribute to tubular ER network formation.

Abstract

The plant endoplasmic reticulum (ER), which is morphologically divided into tubules and sheets, seems to flow continuously as a whole, but locally, mobile and immobile regions exist. In eukaryotes, the ER physically and functionally interacts with the plasma membrane (PM) at domains called ER-PM contact sites (EPCSs). Extended synaptotagmin family proteins are concentrated in the cortical ER to form one type of EPCS; however, it is unclear whether the localization of extended synaptotagmin corresponds to the EPCS and where in the cortical ER the EPCSs are formed. Here, we analyzed the spatiotemporal localization of SYNAPTOTAGMIN1 (SYT1), a synaptotagmin in Arabidopsis (Arabidopsis thaliana), to investigate the precise distribution of SYT1-associated EPCSs in the cortical ER. Three-dimensional imaging using superresolution confocal live imaging microscopy demonstrated that SYT1 was specifically localized to the ER-PM boundary. Time-lapse imaging revealed that SYT1 was distributed to immobile ER tubules, but not to mobile tubules. Moreover, SYT1 was frequently localized to the edges of ER sheets that were transformed into immobile ER tubules over time. A lower intracellular calcium ion concentration resulted in an increased EPCS area and disrupted the ER network. Finally, SYT1 deficiency caused a reduction of the immobile tubules and enlargement of the ER meshes. Taken together, our findings show that SYT1-associated EPCS are distributed to immobile tubules and play an important role in the formation of the tubular ER network. This provides important insight into the relationship between the function and dynamics/morphology of the cortical ER.

The endoplasmic reticulum (ER) is a highly dynamic organelle that is distributed throughout the cytoplasm in plant cells. Although the ER as a whole flows continually and rapidly via the acto-myosin system (Ueda et al., 2010; Peremyslov et al., 2013), mobile and persistent regions exist locally (Sparkes et al., 2009a). The ER morphology is commonly divided into a tubular form (hereafter referred to as “tubule”) and a cisternal form (hereafter referred to as “sheet”). Mobile and persistent regions are thought to exist in both the tubule and sheet forms (Sparkes et al., 2009a).

During the continual flow of the ER, it physically and functionally interacts with other organelles, including chloroplasts, peroxisomes, Golgi bodies, and the plasma membrane (PM; Pérez-Sancho et al., 2016). In yeast and mammals, ER-PM contact sites (EPCSs) play important roles in lipid exchange and in the regulation of Ca2+ homeostasis (Prinz, 2014). Many tethering factors have been reported between the cortical ER and the PM, and several types of EPCS have been proposed in eukaryotes (Gallo et al., 2016; Wang et al., 2017).

A type of EPCS consists of extended synaptotagmin (E-Syt) family proteins. In yeast and mammals, three E-Syt family proteins, named E-Syt 1-3 and tricalbin 1-3, respectively (Manford et al., 2012; Giordano et al., 2013), are involved in the formation of EPCS. These E-Syt family proteins are anchored to the ER membrane via an N-terminal hydrophobic hairpin and bind to the PM via C-terminal lipid binding domains. In mammals, the extent to which the C-terminal domains bind to the PM is dependent on the cytosolic Ca2+ concentration and on whether the E-Syts form homo- or heterodimers. The E-Syt1 homodimer and E-Syt1/E-Syt2 heterodimer are predominantly localized throughout the ER at resting Ca2+ levels whereas the E-Syt2 homodimer is concentrated in the cortical ER at resting Ca2+ levels (Giordano et al., 2013; Idevall-Hagren et al., 2015). In plants, only SYNAPTOTAMIN1 (SYT1) has been proposed to be a component of EPCS. Several studies have analyzed the localization of SYT1 in 2D imaging using confocal laser scanning microscopy (CLSM) and have shown that SYT1 is localized to the stationary nodes on the ER (Levy et al., 2015; Pérez-Sancho et al., 2015; Siao et al., 2016).

E-Syt family proteins have not been shown to be localized to the boundary between the ER and PM in living cells using 3D imaging in plants, yeast, or mammals; therefore, it was not clear in which region of the ER the EPCS reside in the context of ER morphology and dynamics. This is mainly owing to technical limitations. The closest distance between the ER and PM in EPCS is expected to be less than 20 nm (Fernández-Busnadiego et al., 2015), and the ER rapidly flows in plant cells. These factors have made it difficult to investigate the EPCS distribution in detail by using common CLSM. In this study, we showed specific localization of SYT1 to EPCS in Arabidopsis (Arabidopsis thaliana) in 3D using superresolution confocal live imaging microscopy (SCLIM) equipped with a high-speed spinning-disk confocal scanner and a high-sensitivity camera system to achieve superresolution (Matsuura-Tokita et al., 2006; Kurokawa et al., 2013). We further revealed the EPCS properties and distribution in the context of ER morphology and dynamics using time-lapse imaging and treatment with several chemical inhibitors. Our findings provide detailed information on EPCS distribution in the ER, which provide a molecular basis for understanding the relationship between the function and dynamics/morphology of the cortical ER.

RESULTS

SYT1 Is Specifically Localized to the Boundary between the ER and the PM

To investigate SYT1 localization, we first generated transgenic Arabidopsis coexpressing mCherry fused with an ER retention signal as an ER marker (mCherry-HDEL) and SYT1 fused with a monomeric GFP (SYT1-mGFP) driven by the SYT1 promoter in the syt1 background. This syt1 mutant is a T-DNA insertion line used in the previous studies and exhibits a modest growth defect under normal growth conditions (Schapire et al., 2008; Levy et al., 2015; Pérez-Sancho et al., 2015; Siao et al., 2016). Since previous studies have shown that the subcellular localization of membrane proteins can be artificially altered by dimerization of fluorescence proteins (Segami et al., 2014), we used mGFP, which is a GFP variant with an amino acid substitution that prevents artificial GFP dimerization (Zacharias et al., 2002). Under CLSM, SYT1-mGFP was seen as discontinuous structures resembling small beads on strings and partially colocalized with the mCherry-HDEL (Fig. 1A).

Figure 1.

Figure 1.

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SYT1 is specifically localized to the boundary between the ER and PM. A, CLSM visualization of transgenic Arabidopsis expressing mCherry-HDEL (pseudocolor magenta) and ProSYT1::SYT1-mGFP. B and C, SCLIM visualization. The cell cortex of transgenic lines expressing SYT1-mGFP in the syt1 background was observed. The ER and PM were labeled with mCherry-HDEL (B; pseudocolor magenta) and FM4-64 (C; pseudocolor magenta). Z-stack images were captured at 0.1-µm intervals. xy (i) and xz (ii) views. The dotted line in (i) indicates the y axis of (ii). The signal intensity along the dotted line in (ii) is presented in (iii). The coordinate of the peak of SYT1 signal intensity was set to 0. The z-stack images were deconvolved and reconstructed into 3D images (iv). Arrowheads indicate SYT1-mGFP that was localized to the ER tubules. Epidermal cells of true leaves were observed 13 d after sowing. Bars = 5 µm in A and 1 µm in B and C.

Next, to investigate the extent to which SYT1 was localized to the ER-PM boundary, we analyzed SYT1 localization in 3D. The closest distance between the ER and PM in EPCS is expected to be less than 20 nm (Fernández-Busnadiego et al., 2015), and the ER rapidly flows in plant cells. Therefore, we used SCLIM equipped with a high-speed spinning-disk confocal scanner and a high-sensitivity camera system to achieve superresolution beyond the diffraction limit by mathematical data processing (Matsuura-Tokita et al., 2006; Kurokawa et al., 2013). In syt1 mutant plants expressing SYT1-mGFP and mCherry-HDEL, SYT1-mGFP was primarily localized to the ER tubules (Fig. 1Bi). A horizontal view of the z-stack images showed that SYT1-mGFP was specifically localized on the PM side of the ER (Fig. 1, Bii and Biii; Supplemental Video 1). The distance between SYT1-mGFP and the ER was estimated to be less than 0.3 µm by calculating the distance between the peaks of signal intensities. We note that it is the estimation of distance between the centers rather than the closest distance. The 3D reconstruction with deconvolution more clearly separated SYT1-mGFP and mCherry-HDEL and showed that SYT1-mGFP was predominantly localized along the PM side of ER tubules (Fig. 1Biv). In SYT1-mGFP-expressing syt1 plants stained with FM4-64, a PM-staining dye, the z-position of SYT1-mGFP was almost indistinguishable from the PM in the vertical view of the z-stack images (Fig. 1Ci; Supplemental Video 2). However, the horizontal view and signal intensity measurement indicated that SYT1-mGFP was located slightly below the PM (Fig. 1, Cii and Ciii). The distance between SYT1-mGFP and the PM was estimated to be less than 0.2 µm. Moreover, deconvolution and 3D reconstruction of the z-stack images more clearly showed that SYT1-mGFP contacted the PM (Fig. 1Civ). Taken together, these results suggest that SYT1-mGFP is specifically localized to the boundary between the ER and PM (i.e. the EPCS) in the normal cytosolic Ca2+ concentration.

EPCS Localize to Immobile ER Tubules

Since the EPCS appeared to localize to ER tubules, we next analyzed the relationship between the distribution of the EPCS and the ER dynamics by time-lapse imaging of SYT1-mGFP and mCherry-HDEL. The ER tubules rapidly changed their anchoring points, whereas the EPCS appeared to be immobile (Supplemental Video 3). The first of the time-lapse images shows two types of ER tubules that SYT1 was either localized to or not (Fig. 2Ai). A persistent region of the EPCS and ER was identified from the time-lapse images by the averaging or minimizing process, respectively (Fig. 2Aii, left and middle). In addition to the minimizing process, we used the averaging process to extract the persistent area because SYT1-mGFP images had lower signal intensity and the resolution of the minimized SYT1-mGFP image was poor. Merging of the first images with the averaged or minimized images revealed that the EPCSs were almost fixed and that mobile and immobile ER tubules existed (Fig. 2Aiii, left and middle). The averaged SYT1-mGFP image was almost perfectly merged with the persistent region of the ER excluding the tubule junctions (Fig. 2Aii, right). Furthermore, when the first SYT1-mGFP image was compared with the minimized mCherry-HDEL image, almost all the EPCS colocalized with the immobile tubules (Fig. 2Aiii, right). The Pearson’s colocalization coefficient (PCC) increased from 0.32 to 0.67 before and after the minimizing process was applied to the mCherry-HDEL images, suggesting that SYT1-mGFP distribution almost coincided with the persistent region of the ER. To investigate whether SYT1-mGFP was localized to growing tubules, SYT1-mGFP and mCherry-HDEL were visualized using variable-angle epifluorescence microscopy (VAEM). The time-lapse images show that SYT1-mGFP was not localized to the growing tubule (Fig. 2B; Supplemental Video 4). Collectively, these results indicate that EPCS localizes to the immobile tubules, but not to mobile tubules, which have changing anchor points or are growing.

Figure 2.

Figure 2.

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EPCS are distributed to the immobile tubules. The cell cortex of transgenic Arabidopsis expressing mCherry-HDEL (pseudocolor magenta) and SYT1-mGFP in the syt1 background. A, Forty frames of images were captured at 2.46-s intervals. (i) The first images of the time lapse. (ii) The persistent areas extracted by averaging (SYT1-mGFP) or minimizing (mCherry-HDEL) the time-lapse images. (iii) The merged images of the first images (i) and persistent areas (ii). The persistent areas of SYT1-mGFP and mCherry-HDEL are pseudocolored magenta and green, respectively. The bottom right is a merged image of the first image of SYT1-mGFP (i) and the persistent area of mCherry-HDEL (ii). The numbers on the images are the average of the Pearson’s colocalization coefficient (n = 3). Arrowheads indicate that SYT1-mGFP was not distributed in mobile tubules. The arrows indicate the tubule junction where SYT1-mGFP was not localized. B, Visualization of a growing tubule by VAEM. Five frames of images are shown at 0.41-s intervals. Arrowheads indicate the growth of tubules where SYT1 was not localized. Epidermal cells of true leaves were observed 13 d after sowing. Bars = 5 µm in A and 1 µm in B.

EPCS Are Mainly Distributed to ER Sheet Edges, Which Are Converted into Immobile ER Tubules over Time

To assess the distribution of the EPCS in the ER sheets, we observed the epidermal cells of petioles, which have well-developed sheets in comparison with the cells in the middle of true leaves. Visualization of mCherry-HDEL by VAEM revealed that large sheets overlapped with tubules and small sheets. SYT1-mGFP was localized predominantly to the edges of sheets and partially found in the tubules overlapped with the sheets (Fig. 3A). Overlapped tubules and sheets were also observed in transgenic Arabidopsis expressing GFP-HDEL, thus excluding the possibility that these overlapped tubules and sheets were artificially generated by the expression of SYT1-mGFP (Supplemental Fig. S1). Time-lapse images obtained by CLSM showed that the EPCS with SYT1-mGFP found in the sheet edges became immobile tubules over time (Fig. 3B; Supplemental Video 5). In contrast, the time-lapse movie obtained by VAEM showed that part of the sheet edges rapidly moved like mobile tubules (Supplemental Video 6). Moreover, mobile tubules and immobile tubules with SYT1-mGFP were occasionally seen in the middle of the ER sheet (Fig. 3C; Supplemental Video 6). Taken together, these data demonstrated that even when overlapped with ER sheets, there are regions with the potential to form mobile tubules or immobile tubules, the latter of which contain EPCS.

Figure 3.

Figure 3.

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EPCS distribution in ER sheets. A to C, The cell cortex of transgenic Arabidopsis expressing mCherry-HDEL (pseudocolor magenta) and SYT1-mGFP in the syt1 background. A, VAEM images. Tubules (arrow) and cisternae (arrowhead) were overlapping in the ER sheets. Color-coded images based on signal intensity (bottom row). B, CLSM images. Forty frames of images were captured at 4.41-s intervals. (i) The first images of the time lapse. (ii) The persistent areas extracted by minimizing (mCherry-HDEL) or averaging (SYT1-mGFP) the time-lapse images. (iii) The merged images of the first images (i) and persistent areas (ii). The persistent areas of mCherry-HDEL and SYT1-mGFP are pseudocolored green and magenta, respectively. The bottom right is a merged image of the first image of mCherry-HDEL (i) and the persistent area of SYT1-mGFP (ii). EPCS with SYT1-mGFP found in the edges of the sheet became an immobile tubule (arrowheads). C, Merged images of the persistent area of SYT1-mGFP and the first image or the persistent area of mCherry-HDEL. Using VAEM, 100 frames were captured at 0.050-s intervals. A mobile tubule (arrowheads) and an immobile tubule (arrows) were observed in the middle of the sheet. Epidermal cells of petioles (A–C) were observed 13 d after sowing. Bars = 5 µm in A, 2 µm in B, and 1 µm in C.

To characterize the 3D structure of the sheets and tubules, we stained the PM of transgenic Arabidopsis plants expressing the ER marker, GFP-HDEL, with FM4-64 and visualized the PM and ER using SCLIM. The horizontal view of the ER showed that the sheets were raised more toward the cytoplasmic side than the tubules (Fig. 4, A and B). 3D reconstruction with deconvolution of z-stack images also supported the idea that sheets are raised more than tubules (Fig. 4C). We estimated the distance of a tubule and a sheet from the PM by calculating the median distance of the signal intensity measurements (Fig. 4D) and found that ER sheets are more distant (approximately 150 nm) from the PM than tubules. The distance between the PM and the tubule was less than 0.4 to 0.5 µm, and this value was in close agreement with the sum of the distances of PM-EPCS and EPCS-tubule mentioned above (Fig. 1, Biii and Ciii). This verified the accuracy of these estimations. Given that sheet edges are converted from immobile tubules (Fig. 3), which are in contact with the PM (Figs. 1, 2, and 4), the ER sheet has a flat dome-like structure that is in contact with the PM at part of its edge.

Figure 4.

Figure 4.

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ER sheets are more distant from the PM than tubules. The cell cortex of transgenic Arabidopsis expressing GFP-HDEL, as visualized by SCLIM. A, Vertical view. FM4-64 staining of the PM is shown by magenta. The dotted line indicates the y axis of B. B, xz view. Arrows and arrowheads indicate sheets and tubules, respectively. C, Z-stack images were deconvolved and reconstructed into 3D images. FM4-64 staining of the PM is shown by blue. D, The signal intensities along the dotted lines in B. The signal intensity of the FM4-64-stained PM was calculated by averaging signal intensities along the two dotted lines. The coordinate of the peak of FM4-64 signal intensity was set to 0. The solid lines in the graph indicate the median distances of the sheet and tubule from the PM. Epidermal cells of true leaves were observed 13 d after sowing. Bar, 1 µm.

ER Tubules Associated with EPCS Are LatB Insensitive

To gather further information on the relationship between the distribution of EPCS and the dynamics of the ER, latrunculin B (LatB), an inhibitor of actin polymerization, was applied. Large ER aggregations containing SYT1-mGFP were occasionally observed in LatB-treated cells (Supplemental Fig. S2), in accordance with a previous study (Pérez-Sancho et al., 2015). Excluding the aggregation area, almost all tubules became labeled with SYT1-mGFP, and enlargement of the sheet area was observed (Fig. 5, A and B, left). This effect is probably because mobile tubules, which are maintained by the function of actin filaments, formed into sheets after the LatB treatment. In agreement with this hypothesis, the PCC values did not significantly change after the LatB treatment (Fig. 5B, right). Alternation of the distribution of SYT1-mGFP was not found, indicating that immobile tubules and EPCS are independent of actin filaments. This experiment suggests that there are structural and functional differences between mobile and immobile tubules.

Figure 5.

Figure 5.

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LatB tolerance in immobile tubules with SYT1 localization. Transgenic Arabidopsis expressing mCherry-HDEL (pseudocolor magenta) and SYT1-mGFP in the syt1 background were treated with 0.5% DMSO or 10 µm LatB for 1 h. A, Images visualized by CLSM. B, Quantification of the LatB effect. Left, the proportion of the number of tubules labeled with SYT1-mGFP. n = 3 per treatment. Right, the average of the Pearson’s colocalization coefficient. n = 6 per treatment. Error bars indicate the sd. *P < 0.01 using Student’s t test. Epidermal cells of true leaves were observed 13 d after sowing. Bars = 5 µm.

A Decrease in the Intracellular Ca2+ Concentration Causes Disruption of the ER Network and an Increase in the Area of EPCS

Since SYT1 has Ca2+-dependent lipid-binding domains, we manipulated the intracellular Ca2+ concentration by treatment with lanthanum ion (La3+) or ethylene glycol tetra-acetic acid acetoxymethyl ester (EGTA-AM). Treatment with a high concentration of La3+ causes a decrease in the intracellular Ca2+ concentration by blocking the binding of Ca2+ to the PM in a competitive manner (Segal, 1986). EGTA-AM also decreases the intracellular Ca2+ concentration by chelating Ca2+ (Idevall-Hagren et al., 2015). The controls, 0.2% DMSO and 2 mm NaCl, showed no effect on the ER and EPCS (Fig. 6A). Treatment with 1 mm LaCl3 caused severe effects on the distribution of EPCSs and the ER structure. A number of dot-shaped EPCSs appeared, and the total area of the EPCS increased (Fig. 6, A and B). The ER showed fragmentation of the network and segregation from the EPCS (Fig. 6C), and fine pores were formed in ER sheets. To characterize the morphological change of the ER, we quantified the ER shape using distance transformation (Fig. 6D). Distance transformation changes each point of a binary image to the distance from the closest boundary (Supplemental Fig. S3). A greater number of pixels with low intensity indicates that the ER network was more tubular or fragmented. Using this approach, we successfully quantified the fragmented ER when LaCl3 was applied. Similar effects were seen when 200 µm EGTA-AM was applied, but the EPCS became tubule-like structures unlike the numerous dot-like EPCS observed when LaCl3 was applied. The ER was more severely affected and formed ring-shaped aggregations at the tubule junctions. These observations indicate that the intracellular Ca2+ concentration is intimately involved in the distribution of EPCS and the ER structure.

Figure 6.

Figure 6.

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A decrease of the intracellular Ca2+ concentration affects the ER network and EPCS distribution. Transgenic Arabidopsis expressing mCherry-HDEL (pseudocolor magenta) and SYT1-mGFP in the syt1 background were treated with 0.2% DMSO, 2 mm NaCl, 1 mm LaCl3, or 200 µm EGTA-AM. A, Images visualized by CLSM. Images of mCherry-HDEL were binarized and processed with distance transformation (bottom left; see Supplemental Fig. S3). Epidermal cells of true leaves were observed at 13 d after sowing. Bars = 5 µm. B to D, Quantification of the EPCS (SYT1-mGFP) area (B), colocalization of the ER (mCherry-HDEL) with the EPCS (SYT1-mGFP) using the Pearson’s correlation coefficient (C) and morphological change of the ER (mCherry-HDEL) using distance transformation (D). The inset shows an enlargement of the region with higher signal intensities. Error bars indicate sd. **P < 0.05; *P < 0.01, n = 6 per treatment; using Student’s t test (B and C). The DMSO and EGTA-AM or NaCl and LaCl3 data sets were significantly different according to Kolmogorov-Smirnov tests (P < 0.01; D).

The Number of Immobile ER Tubules Is Reduced in the syt1 Mutant

We next investigated the ER morphology of the syt1 mutant. Although the ER morphology of the syt1 mutant has been investigated in previous studies (Levy et al., 2015; Pérez-Sancho et al., 2015; Siao et al., 2016), no consensus has been reached because quantitative analysis using an ER marker was not carried out. In this study, to minimize differences associated with the developmental stage and cell type, we used the hypocotyls of etiolated seedlings expressing the ER marker GFP-HDEL at 6 d after sowing. In the syt1 mutant, the number of ER tubules was reduced and the sheet area increased in comparison with the wild type. Moreover, the meshes of the ER network were larger than those of the wild type (Fig. 7A). Quantification of ER morphology, the number of meshes, the mesh size, and the area occupied by each size of mesh supported these observations (Fig. 7, Ci and Cii). The number of meshes in the syt1 mutant was approximately half that of the wild type, whereas the average mesh size was approximately double. Smaller meshes (1–10 µm2) were significantly reduced in abundance in the syt1 mutant, and larger meshes (>20 µm2) were significantly increased in abundance. Time-lapse analysis showed that the persistent area of the ER was reduced by half in the syt1 mutant (Fig. 7, B and Ciii). Combined with the localization of immobile tubules in SYT1, these results indicate that SYT1 plays central roles in forming and maintaining immobile tubules.

Figure 7.

Figure 7.

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Immobile tubules are reduced in the syt1 mutant. Transgenic Arabidopsis expressing GFP-HDEL in the wild-type (wt) or syt1 background. A, Images visualized by CLSM (top row). These images were binarized and processed with distance transformation (bottom row). B, Forty frames of images were captured at 2.46-s intervals. The first images of the time lapse are presented in the top row. The persistent area was extracted by minimizing the time-lapse images (bottom row). C, Quantification of the ER profile. (i) Morphological changes in the ER were quantified using distance transformation. The inset shows an enlargement of the region with higher signal intensities. The two data sets are significantly different according to a Kolmogorov-Smirnov test (P < 0.01). (ii) The number of meshes, the average mesh size, and the area occupied by each size of mesh. (iii) The proportion of persistent area in the ER. Error bars indicate the sd. *P < 0.01, n = 10 per group; using Student’s t test. D, 3D Images visualized by SCLIM. (i) GFP-HDEL in the wild-type background. The same z-stack images as Fig. 4A were used. The PM was stained with FM4-64 (pseudocolor magenta). (ii) The syt1 mutant expressing GFP-HDEL. The dotted line indicates the y axis of the xz views. The signal intensity along the arrows is shown to the right. The coordinate of the peak of FM4-64 signal intensity was set to 0. The z-stack images of (ii) were deconvolved and reconstructed into 3D images (bottom). E, Complementation assay. The wild type expressing mCherry-HDEL (wt), the syt1 mutant expressing mCherry-HDEL (syt1), and the syt1 mutant expressing mCherry-HDEL and SYT1-mGFP (rescued) were used. Quantification of the average mesh size (left) and the area occupied by each size of mesh (right). Error bars indicate the sd. **P < 0.05; *P < 0.01, n = 10 per group; using Student’s t test. The epidermal cells of the hypocotyls of etiolated seedlings at 6 d after sowing were used in A to C. Epidermal cells of the true leaves at 13 d after sowing were used in D and E. Bars = 5 µm in A and B and 1 µm in D.

Since the loss of SYT1 was expected to cause detachment of the ER from the PM, we investigated whether the ER was associated with the PM in the syt1 mutant using SCLIM. The signal intensity of the horizontal view showed that the distance between the ER and the PM was slightly greater in the syt1 mutant (Fig. 7, Di and Dii). However, this was hardly distinguishable even in the deconvoluted images, and we were unable to quantify the mean distance between the ER and the PM. Considering that the ER sheets were more distant from the PM than the tubules (Fig. 4), the reduction in immobile tubules and the increase in sheets in the syt1 mutant probably indicates detachment of the ER from the PM.

To determine whether the transgenic line expressing SYT1-mGFP in the syt1 background rescued the syt1 phenotype, we compared the ER morphology between the wild type, the syt1 mutant, and SYT1-mGFP-rescued lines using mCherry-HDEL. Although there was no significant difference between the wild type and SYT1-mGFP-rescued lines in terms of the average mesh size, an increase in the average mesh size was observed in the syt1 mutant compared to the wild type and the rescued lines (Fig. 7E). The reduction in the abundance of smaller meshes (1–3 µm2) and the increase in the abundance of larger meshes (>6 µm2) were abolished in the rescued line. Therefore, SYT1-mGFP complements the intrinsic SYT1 function.

LatB and La3+ Differentially Affect the syt1 and Wild-Type Plants

To further investigate the function of SYT1, the syt1 mutant was treated with LatB or LaCl3. When treated with LatB, the ER network was severely disrupted. The mesh number decreased and the mean mesh size increased in both the syt1 and wild-type plants (Fig. 8, A and B). LatB had a stronger effect in the syt1 mutant than in the wild type, and there was a greater increase in the abundances of large meshes in the syt1 mutant. This effect may be caused by the reduction of immobile tubules, which are LatB insensitive (as shown in Fig. 5). Conversely, with LaCl3 treatment, the mesh number increased and the average mesh size decreased in both the syt1 mutant and the wild type. Fine pores in the sheets were frequently observed in the wild type but were rarely observed in the syt1 mutant. This indicates that the loss of SYT1 caused the different responses to La3+ and raises the possibility that the decrease in the intracellular Ca2+ concentration directly affects SYT1.

Figure 8.

Figure 8.

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Different effects of LatB and La3+ on the ER morphology between the syt1 mutant and the wild type. Transgenic Arabidopsis expressing GFP-HDEL in the wild-type or syt1 background. A, The effect of 0.5% (v/v) DMSO, 10 µm LatB, 2 mm NaCl, or 1 mm LaCl3 on the ER in the wild type or in the syt1 mutant. Epidermal cells of the hypocotyls of etiolated seedlings at 6 d after sowing. Bars = 5 µm. B, The average number of meshes and the area occupied by each size of mesh in each treatment (±sd). Asterisks indicate significant differences from the corresponding controls using Student’s t test (P < 0.01, n = 10 per treatment).

DISCUSSION

Previous studies in plants, yeast, and mammals analyzed the localization of E-Syt family proteins in 2D (Schapire et al., 2008; Manford et al., 2012; Giordano et al., 2013; Levy et al., 2015; Siao et al., 2016); however, 3D information on the localization of E-Syt family proteins was lacking. Since the closest distance between the ER and PM in EPCS is expected to be less than 20 nm (Fernández-Busnadiego et al., 2015) and the ER rapidly moves throughout the cytoplasm in plant cells, it was difficult to distinguish EPCS from the ER and PM using CLSM. In this study, we have successfully visualized SYT1 that was specifically localized to EPCS using SCLIM, which enables high-speed scanning at a better resolution than CLSM (Figs. 1 and 9). This means that SYT1 can be used as an EPCS marker at least at the normal cytosolic Ca2+ concentration.

Figure 9.

Figure 9.

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Schematic illustration of EPCS distribution in the ER on the PM. 3D illustration (top) and 2D illustration (bottom). EPCS (deep green) are localized to immobile tubules and the edges of sheets in the cortical ER (pale green). Mobile tubules are mainly driven by actin filaments (purple). Ribosomes (beige balls) are frequently found on the ER sheets and eliminated from EPCS.

We quantitatively demonstrated that SYT1 was localized to immobile tubules (Fig. 2) and that SYT1 deficiency caused a reduction of the immobile ER tubules (Fig. 7). Our finding indicated that there are two types of ER tubules, mobile tubules without SYT1 and immobile tubules with SYT1, which probably function differently in the ER network, and that the formation of EPCS and immobile tubules by SYT1 are essential for proper organization of the ER network (Fig. 9). Although several previous studies investigated SYT1 localization in the ER and the ER phenotype of the syt1 mutant (Schapire et al., 2008; Levy et al., 2015; Pérez-Sancho et al., 2015), SYT1 distribution and function in the context of ER morphology and ER dynamics is largely unclear. One major reason for this is that no consensus result was obtained from phenotyping of the syt1 mutant. Pérez-Sancho et al. (2015) reported no phenotypic alterations in the ER, whereas Levy et al. (2015) reported the collapse of the ER network. In this study, we quantified the various aspects of the ER dynamics and morphology using etiolated seedlings and revealed that the syt1 mutant shows an intermediate phenotype between those reported previously (Levy et al., 2015; Pérez-Sancho et al., 2015). In these previous studies, the reduction in the ER tubules may have been overlooked or overestimated because developing leaves, which have cells of various sizes at various developmental stages, were used without quantification. Consistent with this hypothesis, a previous study showed that the densities of SYT1-labeled EPCS and ER change depending on the developmental stage and cell type (McFarlane et al., 2017).

We showed that some ER tubules are overlapped with ER sheets by using VAEM (Fig. 3) and that the ER sheets are more distant from the PM than tubules by using SCLIM (Fig. 4). SYT1 are frequently localized to sheet edges that were transformed into immobile tubules over time, indicating that ER sheets contact the PM mainly at the edges, which are functionally and morphologically related to immobile tubules (Fig. 3). These findings suggested that the ER sheet has a flat dome-like structure and is in contact with the PM at its edge (Fig. 9). Consistent with this finding, the previous report on the structure of ER tubules and sheets in animal cells suggested structural and functional similarities between sheet edges and tubules (Shibata et al., 2010). Immobile tubules occasionally seen in the middle of sheets can be interpreted in two ways: immobile tubules associated with the edges of small sheets below the sheets or independent immobile tubules below the ER sheets. Either or both of these immobile tubules probably correspond with the persistent region found in the middle of ER sheets in previous studies (Sparkes et al., 2009a, 2010).

Although EPCS distribution almost coincided with the persistent region of the ER, the EPCSs were not distributed to the tubule junctions despite their immobility (Fig. 2A). Tubule junctions are anchor/growth sites of mobile tubules and are mainly regulated by actin filaments and microtubules (Sparkes et al., 2009b; Hamada et al., 2014). Another type of EPCS, which consists of VESICLE-ASSOCIATED MEMBRANE PROTEIN-ASSOCIATED PROTEIN27 (VAP27) and NETWORKED SUPERFAMILY PROTEIN3C proteins, may be distributed to these junctions (Saravanan et al., 2009; Wang et al., 2014, 2016; Siao et al., 2016). This idea is supported by the previous finding that EPCS consisting of VAP27/NETWORKED SUPERFAMILY PROTEIN3C interact with actin filaments and microtubules (Wang et al., 2014). Moreover, in our study, the SYT1-associated EPCS were LatB insensitive (Fig. 5), indicating that the EPCS associated with SYT1 are independent of actin filaments. This suggested functional and structural differences between VAP-associated EPCS and SYT-associated EPCS.

Our study provides insights into the functional difference between tubules and sheets. Previous studies using electron microscopy showed that ER sheets have higher ribosome density than ER tubules and that ribosomes are excluded from the region where the ER is in contact with the PM (Puhka et al., 2007; Friedman and Voeltz, 2011; McFarlane et al., 2017). Furthermore, transformation of the sheeted ER into a branched tubular network correlates with a decrease in ribosomes on the ER membrane (Puhka et al., 2007). These data are highly compatible with our model; immobile ER tubules, which are associated with EPCS, are specialized for lipid exchange and regulation of Ca2+ homeostasis and consequently may not require ribosomes. Moreover, EPCS probably play pivotal roles such as scaffolding for the ER and determine the tubule:cisterna ratio, because a lack of SYT1 resulted in transformation into the sheeted ER (Fig. 7, A and C). Although several studies have presented confocal or electron microscopy images of the ER ultrastructure, information connecting the ER morphology and the subcellular distribution of certain proteins has been limited (Puhka et al., 2007; Shibata et al., 2010; Friedman and Voeltz, 2011; West et al., 2011; Nixon-Abell et al., 2016; McFarlane et al., 2017). Our findings suggested SYT1 as a key determinant of morphology and functional differentiation of the ER. Unfortunately, our data explain neither the function of the mobile tubules nor the mechanism by which immobile tubules are arranged to organize the ER network. Further studies are required to reveal the biological function of mobile tubules and the process by which immobile tubules are formed in the ER network.

Treatment with La3+ or EGTA-AM suggested that reduction of the intracellular Ca2+ concentration caused the expansion of the EPCS area and segregation between the ER and the EPCS (Fig. 6). This seemingly contradicts with the previous study, in which the C-terminal domain of SYT1 was shown to bind to phospholipids in a Ca2+-dependent manner in vitro (Schapire et al., 2008). However, previous studies in E-Syts similarly reported contradictory results between in vivo and in vitro experiments. E-Syt2 accumulated at EPCS in lower Ca2+ concentrations in vivo (Giordano et al., 2013; Idevall-Hagren et al., 2015), although the C-terminal domain of E-Syt2 bound to phospholipids in a Ca2+-dependent manner in vitro (Min et al., 2007). This indicates that the Ca2+-dependent PM binding of E-Syt family proteins in vivo are intricately regulated by various factors. It is likely that Ca2+ deprivation caused a stronger binding of SYT1 to the PM in plant cells, as was the case of E-Syt2.

In mammals, three E-Syts form homo- and heterodimers and work in a coordinated manner to bind to the PM (Giordano et al., 2013; Idevall-Hagren et al., 2015). Ca2+ dependence is varied among E-Syt1 to 3 (Idevall-Hagren et al., 2015). Elevation of the intracellular Ca2+ concentration led to a proportional increase of E-Syt1 binding to the PM. In contrast, E-Syt2 more strongly bound to the PM at lower Ca2+ concentrations as described above. Thus, SYT1 is more similar to E-Syt2 than to E-Syt1 in its Ca2+ dependence. In plants, synaptotagmin proteins other than SYT1 have not been found in EPCS; however, other synaptotagmin proteins that have different Ca2+ sensitivity are probably involved in EPCS functions. This prediction is supported by the finding that immobile tubules, although lower in abundance, were still observed in the syt1 mutant (Fig. 7B). This suggests that other SYTs may function redundantly in ER-PM tethering in plant cells.

MATERIALS AND METHODS

Plant Materials and Growth Conditions

Arabidopsis (Arabidopsis thaliana) ecotype Columbia-0 (Col-0; CS60000) was used as the wild-type plant. The Arabidopsis transgenic line, GFP-HDEL, was previously reported (Matsushima et al., 2002). The SAIL_775A08 T-DNA insertion line was used as the syt1 mutant. Arabidopsis seeds were surface-sterilized and then sown onto 0.5% gellan gum containing 1% (w/v) Suc and Murashige and Skoog medium. Plants were grown at 22°C under continuous light. For etiolated seedlings, plants were grown at 22°C under continuous dark after germination.

Confocal Microscopy

Plant tissues were mounted on microscope slides (CLSM and VAEM) or cover slips (SCLIM). All samples were observed at room temperature. For CLSM, a Zeiss LSM780 confocal laser scanning microscope with 488-nm and 543-nm line lasers, a 493- to 550-nm band-pass emission filter, and a 63 × 1.20 numerical aperture water immersion objective lens were used. SCLIM consists of the Olympus model IX-71 fluorescence microscope with a custom-made high-speed confocal scanner (Yokogawa Electric), image intensifiers (Hamamatsu Photonics) with a custom-made cooling system, and EM-CCD cameras (Hamamatsu Photonics) (Kurokawa et al., 2013). VAEM and SCLIM images were acquired as described previously (Yamaoka et al., 2013; Ito et al., 2018). Image analysis was performed using ImageJ software (NIH). Signal intensity was measured by the Plot Profile tool. Persistent areas were extracted from sequential images consisting of 40 frames at 2.46-s intervals by applying the minimizing or the averaging process of the Z-Projection tool. To create a heat map showing signal intensity, images were color coded by 16 colors in Lookup Tables. To quantify tubules labeled with SYT1-mGFP, the tubules associated with a SYT1-mGFP signal above the threshold were manually counted. The area and number of ER meshes were measured by the Analyze Particles tool. We defined ER meshes as regions in which all of the following conditions were met: (1) the fluorescence intensity was below the threshold; (2) they were completely surrounded by a region whose fluorescence intensity was above the threshold; and (3) they were larger than 0.156 µm2 (corresponding to 3 × 3 pixels). For the distance transformation, binarized images prepared using the Otsu method were transformed by the Distance Map tool. The distance transformation is schematically illustrated in Supplemental Fig. S3. PCCs were measured using a Fiji Colocalization plug-in (Schindelin et al., 2012).

Inhibitor Treatment

True leaves or etiolated seedlings were incubated with 10 µm LatB in 0.5% DMSO, 200 µm EGTA-AM in 0.2% DMSO, or 1 mm LaCl3 in distilled water for 1 h.

Plasmid Construction and Transformation

Transgenic Arabidopsis expressing SYT1-mGFP or mCherry-HDEL were generated using the floral-dip method (Clough and Bent, 1998) with Agrobacterium tumefaciens harboring the SYT1-mGFP or mCherry-HDEL expression vector. To construct the SYT1-mGFP expression vector, a region spanning 1.5 kb upstream of the SYT1 transcriptional start site to just before the stop codon (At2g20990) was amplified by PCR from Col-0 genomic DNA using the following primers: 5′-AACCAATTCAGTCGACATGGGCTTTTTCAGTACG-3′ and 5′-AAGCTGGGTCTAGATATCCAGAGGCAGTTCGCCACTC-3′. The amplified ProSYT1::SYT1 fragment was cloned into a pENTR1a plasmid vector (Invitrogen) using the In-Fusion HD Cloning Kit (Clontech) and the SalI and EcoRV restriction sites and subsequently transferred to the pGWB404m vector (Nakagawa et al., 2007; Segami et al., 2014) using an LR reaction (Invitrogen). For the construction of the mCherry-HDEL expression vector, an ER-targeted mCherry sequence was PCR amplified from er-rk/er-rb (Nelson et al., 2007) using the following primers: 5′-GCAGGCTCCGCGGCCGCATGAAGGTACAGGAGGGTTTG-3′ and 5′-AGCTGGGTCGGCGCGCCTTACAGCTCGTCATGAGATC-3′. Part of pENTR/D-TOPO was amplified by inverse PCR using the following primers: 5′-GGCGCGCCGACCCAGCT-3′ and 5′-GCGGCCGCGGAGCCTGC-3′. The amplified fragments were then cloned using the In-Fusion HD Cloning Kit and subsequently transferred to the pB0GW7 vector, which harbors a UBIQUITIN10 promoter (Ichino et al., 2014), using an LR reaction. Col-0 and the syt1 mutant expressing SYT1-mGFP and mCherry-HDEL or GFP-HDEL were generated by crossing.

Statistical Analyses

The data sets used for each analysis were obtained from the plants grown at the same time. The numbers of cells used for statistical analyses are given in the figure legends. Statistical significance was determined by Student’s t test or Kolmogorov-Smirnov tests.

Accession Numbers

Accession numbers are as follows: SYT1 (gene, NM127668; protein, NP565495).

Supplemental Data

The following supplemental materials are available.

Acknowledgments

We thank Shelley Robison from Edanz Group (www.edanzediting.com/ac) for editing a draft of this manuscript.

References

  1. Clough SJ, Bent AF (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16: 735–743 [DOI] [PubMed] [Google Scholar]
  2. Fernández-Busnadiego R, Saheki Y, De Camilli P (2015) Three-dimensional architecture of extended synaptotagmin-mediated endoplasmic reticulum-plasma membrane contact sites. Proc Natl Acad Sci USA 112: E2004–E2013 [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Friedman JR, Voeltz GK (2011) The ER in 3D: a multifunctional dynamic membrane network. Trends Cell Biol 21: 709–717 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Gallo A, Vannier C, Galli T (2016) Endoplasmic reticulum-plasma membrane associations: structures and functions. Annu Rev Cell Dev Biol 32: 279–301 [DOI] [PubMed] [Google Scholar]
  5. Giordano F, Saheki Y, Idevall-Hagren O, Colombo SF, Pirruccello M, Milosevic I, Gracheva EO, Bagriantsev SN, Borgese N, De Camilli P (2013) PI(4,5)P(2)-dependent and Ca(2+)-regulated ER-PM interactions mediated by the extended synaptotagmins. Cell 153: 1494–1509 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Hamada T, Ueda H, Kawase T, Hara-Nishimura I (2014) Microtubules contribute to tubule elongation and anchoring of endoplasmic reticulum, resulting in high network complexity in Arabidopsis. Plant Physiol 166: 1869–1876 [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Ichino T, Fuji K, Ueda H, Takahashi H, Koumoto Y, Takagi J, Tamura K, Sasaki R, Aoki K, Shimada T, et al. (2014) GFS9/TT9 contributes to intracellular membrane trafficking and flavonoid accumulation in Arabidopsis thaliana. Plant J 80: 410–423 [DOI] [PubMed] [Google Scholar]
  8. Idevall-Hagren O, Lü A, Xie B, De Camilli P (2015) Triggered Ca2+ influx is required for extended synaptotagmin 1-induced ER-plasma membrane tethering. EMBO J 34: 2291–2305 [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Ito Y, Uemura T, Nakano A (2018) The Golgi entry core compartment functions as a COPII-independent scaffold for ER-to-Golgi transport in plant cells. J Cell Sci [DOI] [PubMed]
  10. Kurokawa K, Ishii M, Suda Y, Ichihara A, Nakano A (2013) Live cell visualization of Golgi membrane dynamics by super-resolution confocal live imaging microscopy. Methods Cell Biol 118: 235–242 [DOI] [PubMed] [Google Scholar]
  11. Levy A, Zheng JY, Lazarowitz SG (2015) Synaptotagmin SYTA forms ER-plasma membrane junctions that are recruited to plasmodesmata for plant virus movement. Curr Biol 25: 2018–2025 [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Manford AG, Stefan CJ, Yuan HL, Macgurn JA, Emr SD (2012) ER-to-plasma membrane tethering proteins regulate cell signaling and ER morphology. Dev Cell 23: 1129–1140 [DOI] [PubMed] [Google Scholar]
  13. Matsushima R, Hayashi Y, Kondo M, Shimada T, Nishimura M, Hara-Nishimura I (2002) An endoplasmic reticulum-derived structure that is induced under stress conditions in Arabidopsis. Plant Physiol 130: 1807–1814 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Matsuura-Tokita K, Takeuchi M, Ichihara A, Mikuriya K, Nakano A (2006) Live imaging of yeast Golgi cisternal maturation. Nature 441: 1007–1010 [DOI] [PubMed] [Google Scholar]
  15. McFarlane HE, Lee EK, van Bezouwen LS, Ross B, Rosado A, Samuels AL (2017) Multiscale structural analysis of plant ER-PM contact sites. Plant Cell Physiol 58: 478–484 [DOI] [PubMed] [Google Scholar]
  16. Min S-W, Chang W-P, Südhof TC (2007) E-Syts, a family of membranous Ca2+-sensor proteins with multiple C2 domains. Proc Natl Acad Sci USA 104: 3823–3828 [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Nakagawa T, Suzuki T, Murata S, Nakamura S, Hino T, Maeo K, Tabata R, Kawai T, Tanaka K, Niwa Y, et al. (2007) Improved Gateway binary vectors: high-performance vectors for creation of fusion constructs in transgenic analysis of plants. Biosci Biotechnol Biochem 71: 2095–2100 [DOI] [PubMed] [Google Scholar]
  18. Nelson BK, Cai X, Nebenführ A (2007) A multicolored set of in vivo organelle markers for co-localization studies in Arabidopsis and other plants. Plant J 51: 1126–1136 [DOI] [PubMed] [Google Scholar]
  19. Nixon-Abell J, Obara CJ, Weigel AV, Li D, Legant WR, Xu CS, Pasolli HA, Harvey K, Hess HF, Betzig E, et al. (2016) Increased spatiotemporal resolution reveals highly dynamic dense tubular matrices in the peripheral ER. Science 354: aaf3928. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Peremyslov VV, Morgun EA, Kurth EG, Makarova KS, Koonin EV, Dolja VV (2013) Identification of myosin XI receptors in Arabidopsis defines a distinct class of transport vesicles. Plant Cell 25: 3022–3038 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Pérez-Sancho J, Vanneste S, Lee E, McFarlane HE, Esteban Del Valle A, Valpuesta V, Friml J, Botella MA, Rosado A (2015) The Arabidopsis synaptotagmin1 is enriched in endoplasmic reticulum-plasma membrane contact sites and confers cellular resistance to mechanical stresses. Plant Physiol 168: 132–143 [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Pérez-Sancho J, Tilsner J, Samuels AL, Botella MA, Bayer EM, Rosado A (2016) Stitching organelles: organization and function of specialized membrane contact sites in plants. Trends Cell Biol 26: 705–717 [DOI] [PubMed] [Google Scholar]
  23. Prinz WA. (2014) Bridging the gap: membrane contact sites in signaling, metabolism, and organelle dynamics. J Cell Biol 205: 759–769 [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Puhka M, Vihinen H, Joensuu M, Jokitalo E (2007) Endoplasmic reticulum remains continuous and undergoes sheet-to-tubule transformation during cell division in mammalian cells. J Cell Biol 179: 895–909 [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Saravanan RS, Slabaugh E, Singh VR, Lapidus LJ, Haas T, Brandizzi F (2009) The targeting of the oxysterol-binding protein ORP3a to the endoplasmic reticulum relies on the plant VAP33 homolog PVA12. Plant J 58: 817–830 [DOI] [PubMed] [Google Scholar]
  26. Schapire AL, Voigt B, Jasik J, Rosado A, Lopez-Cobollo R, Menzel D, Salinas J, Mancuso S, Valpuesta V, Baluška F, et al. (2008) Arabidopsis synaptotagmin 1 is required for the maintenance of plasma membrane integrity and cell viability. Plant Cell 20: 3374–3388 [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, et al. (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9: 676–682 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Segal J. (1986) Lanthanum increases the rat thymocyte cytoplasmic free calcium concentration by enhancing calcium influx. Biochim Biophys Acta 886: 267–271 [DOI] [PubMed] [Google Scholar]
  29. Segami S, Makino S, Miyake A, Asaoka M, Maeshima M (2014) Dynamics of vacuoles and H+-pyrophosphatase visualized by monomeric green fluorescent protein in Arabidopsis: artifactual bulbs and native intravacuolar spherical structures. Plant Cell 26: 3416–3434 [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Shibata Y, Shemesh T, Prinz WA, Palazzo AF, Kozlov MM, Rapoport TA (2010) Mechanisms determining the morphology of the peripheral ER. Cell 143: 774–788 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Siao W, Wang P, Voigt B, Hussey PJ, Baluška F (2016) Arabidopsis SYT1 maintains stability of cortical endoplasmic reticulum networks and VAP27-1-enriched endoplasmic reticulum-plasma membrane contact sites. J Exp Bot 67: 6161–6171 [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Sparkes IA, Frigerio L, Tolley N, Hawes C (2009a) The plant endoplasmic reticulum: a cell-wide web. Biochem J 423: 145–155 [DOI] [PubMed] [Google Scholar]
  33. Sparkes I, Runions J, Hawes C, Griffing L (2009b) Movement and remodeling of the endoplasmic reticulum in nondividing cells of tobacco leaves. Plant Cell 21: 3937–3949 [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Sparkes I, Tolley N, Aller I, Svozil J, Osterrieder A, Botchway S, Mueller C, Frigerio L, Hawes C (2010) Five Arabidopsis reticulon isoforms share endoplasmic reticulum location, topology, and membrane-shaping properties. Plant Cell 22: 1333–1343 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Ueda H, Yokota E, Kutsuna N, Shimada T, Tamura K, Shimmen T, Hasezawa S, Dolja VV, Hara-Nishimura I (2010) Myosin-dependent endoplasmic reticulum motility and F-actin organization in plant cells. Proc Natl Acad Sci USA 107: 6894–6899 [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Wang P, Hawkins TJ, Richardson C, Cummins I, Deeks MJ, Sparkes I, Hawes C, Hussey PJ (2014) The plant cytoskeleton, NET3C, and VAP27 mediate the link between the plasma membrane and endoplasmic reticulum. Curr Biol 24: 1397–1405 [DOI] [PubMed] [Google Scholar]
  37. Wang P, Richardson C, Hawkins TJ, Sparkes I, Hawes C, Hussey PJ (2016) Plant VAP27 proteins: domain characterization, intracellular localization and role in plant development. New Phytol 210: 1311–1326 [DOI] [PubMed] [Google Scholar]
  38. Wang P, Hawes C, Hussey PJ (2017) Plant Endoplasmic Reticulum-Plasma Membrane Contact Sites. Trends Plant Sci 22: 289–297 [DOI] [PubMed] [Google Scholar]
  39. West M, Zurek N, Hoenger A, Voeltz GK (2011) A 3D analysis of yeast ER structure reveals how ER domains are organized by membrane curvature. J Cell Biol 193: 333–346 [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Yamaoka S, Shimono Y, Shirakawa M, Fukao Y, Kawase T, Hatsugai N, Tamura K, Shimada T, Hara-Nishimura I (2013) Identification and dynamics of Arabidopsis adaptor protein-2 complex and its involvement in floral organ development. Plant Cell 25: 2958–2969 [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Zacharias DA, Violin JD, Newton AC, Tsien RY (2002) Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 296: 913–916 [DOI] [PubMed] [Google Scholar]

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