Abstract
In eukaryotes, membrane contact sites (MCS) allow direct communication between organelles. Plants have evolved unique MCS, the plasmodesmata intercellular pores, which combine endoplasmic reticulum (ER) - plasma membrane (PM) contacts with regulation of cell-to-cell signalling. The molecular mechanism and function of membrane tethering within plasmodesmata remains unknown.
Here we show that the Multiple C2 domains and Transmembrane region Protein (MCTP) family, key regulators of cell-to-cell signalling in plants, act as ER - PM tethers specifically at plasmodesmata. We report that MCTPs are core plasmodesmata proteins that insert into the ER via their transmembrane region whilst their C2 domains dock to the PM through interaction with anionic phospholipids. A mctp3/4 loss-of-function mutant induces plant developmental defects while MCTP4 expression in a yeast Δtether mutant partially restores ER-PM tethering. Our data suggest that MCTPs are unique membrane tethers controlling both ER-PM contacts and cell-cell signalling.
Introduction
Intercellular communication is essential for the establishment of multicellularity, and evolution gave rise to distinct mechanisms to facilitate this process. Plants have developed singular cell junctions -the plasmodesmata-which span the cell wall and interconnect nearly every single cell, establishing direct membrane and cytoplasmic continuity throughout the plant body (Tilsner et al, 2016). Plasmodesmata are indispensable for plant life. They control the flux of non-cell-autonomous signals such as transcription factors, small RNAs, hormones and metabolites during key growth and developmental events (Gallagher et al, 2014; Tilsner et al, 2016; Vatén et al, 2011; Carlsbecker et al, 2010; Benitez-Alfonso et al, 2013; Wu et al, 2016; Han et al, 2014; Daum et al, 2014; Nakajima et al, 2001; Xu et al, 2011; Ross-elliott et al, 2017). Over the past few years, plasmodesmata have emerged as key components of plant defence signalling (Faulkner et al, 2013; Wang et al, 2013; Lim et al, 2016). Mis-regulation of plasmodesmata function can lead to severe defects in organ growth and tissue patterning but also generate inappropriate responses to biotic and abiotic stresses (Wu et al, 2016; Wong et al, 2016; Han et al, 2014; Sager & Lee, 2014; Caillaud et al, 2014; Faulkner et al, 2013). Plasmodesmata not only serve as conduits, but act as specialised signalling hubs, capable of generating and/or relaying signals from cell-to-cell through plasmodesmata-associated receptor activity (Stahl & Faulkner, 2016; Stahl et al, 2013; Vaddepalli et al, 2014; Lee, 2015).
Plasmodesmata are structurally unique (Nicolas et al, 2017b; Tilsner et al, 2011). They contain a strand of ER, continuous through the pores, tethered extremely tightly (~10 nm) to the PM by spoke-like elements (Ding et al, 1992; Nicolas et al, 2017a) whose function and identity is unknown. Inside plasmodesmata, specialised subdomains of the ER and the PM coexist, each being characterised by a unique set of lipids and proteins, both critical for proper function (Bayer et al, 2014; Grison et al, 2015; Thomas et al, 2008a; Simpson et al, 2009; Zavaliev et al, 2016; Knox et al, 2015; Faulkner et al, 2013; Benitez-Alfonso et al, 2013). Where it enters the pores, the ER becomes constricted to a 15 nm tube (the desmotubule) leaving little room for lumenal trafficking. According to current models, transfer of molecules occurs in the cytoplasmic sleeve between the ER and the PM. Constriction of this gap, by the deposition of callose, is assumed to be the main regulator of the pore size exclusion limit (Vatén et al, 2011; Zavaliev et al, 2011). Recent work however, suggests a more complex picture where plasmodesmal ER-PM gap is not directly related to the pore permeability and may play additional roles (Nicolas et al, 2017a, 2017b). Newly formed plasmodesmata (type I) exhibit such close contact (~2-3nm) between the PM and the ER, that no electron-lucent cytoplasmic sleeve is observed (Nicolas et al, 2017a). During subsequent cell growth and differentiation the pore widens, separating the two membranes, which remain connected by visible electron-dense spokes, leaving a cytosolic gap (type II). This transition has been proposed to be controlled by protein-tethers acting at the ER-PM interface (Bayer et al, 2017; Nicolas et al, 2017b). Counterintuitively, type I plasmodesmata with no apparent cytoplasmic sleeve are open to macromolecular trafficking and recent data indicate that tight ER-PM contacts may in fact favour transfer of molecules from cell-to-cell (Nicolas et al, 2017a).
The close proximity of the PM and ER within the pores, and the presence of tethers qualifies plasmodesmata as a specialised type of ER-to-PM membrane contact site (MCS) (Tilsner et al, 2016; Bayer et al, 2017). MCS are structures found in all eukaryotic cells which function in direct inter-organellar signalling by promoting fast, non-vesicular transfer of molecules and allowing collaborative action between the two membranes (Burgoyne et al, 2015; Prinz, 2014; Phillips & Voeltz, 2016; Gallo et al, 2016; Eden et al, 2010, 2016; Ho et al, 2016; Chang et al, 2013; Kim et al, 2015; Petkovic et al, 2014; Zhang et al, 2005; Omnus et al, 2016). In yeast and mammalian, MCS protein tethers are known to physically bridge the two organelles, to control the intermembrane gap and participate in organelle cross-talk. Their molecular identity/specificity dictate structural and functional singularity to different types of MCS (Eisenberg-Bord et al, 2016; Henne et al, 2015). To date, the plasmodesmal membrane tethers remain unidentified, but by analogy to other types of MCS it seems likely that they play important roles in plasmodesmal structure and function, and given their unique position within a cell-to-cell junction may link intra- and intercellular communication.
Here, we have reduced the complexity of the previously published Arabidopsis plasmodesmal proteome (Fernandez-Calvino et al, 2011) through the combination of a refined purification protocol (Faulkner & Bayer, 2017) and semi-quantitative proteomics, to identify ~120 proteins highly enriched in plasmodesmata, and identify tether candidates. Amongst the most abundant plasmodesmal proteins, members of the Multiple C2 domains and Transmembrane region Proteins (MCTPs) were enriched in post-cytokinetic plasmodesmata with tight ER-PM gap compared to mature plasmodesmata with wider gap and sparse spokes, and exhibit the domain architecture characteristic of membrane tethers, with multiple lipid-binding C2 domains in the N-terminal, and multiple transmembrane domains in the C-terminal region. Using live cell imaging, molecular dynamics, and yeast complementation, we show that MCTP properties are consistent with a role in ER-PM membrane tethering at plasmodesmata. As several MCTP members have been identified as important components of plant intercellular signalling (Liu et al, 2012, 2018; Vaddepalli et al, 2014), our data suggest a link between inter-organelle contacts at plasmodesmata and inter-cellular communication in plants.
RESULTS
Identification of plasmodesmal ER-PM tethering candidates
To identify putative plasmodesmal MCS tethers, we decided to screen the plasmodesmata proteome for ER-associated proteins (a general trait of ER-PM tethers (Henne et al, 2015; Eisenberg-Bord et al, 2016)) with structural features enabling bridging across two membranes. Published plasmodesmata proteome reported the identification of more than 1400 proteins in Arabidopsis (Fernandez-Calvino et al, 2011), making the discrimination of true plasmodesmata-associated from contaminant proteins a major challenge. To reduce the proteome complexity and identify core plasmodesmata proteins, we used a refined plasmodesmata purification technique (Faulkner & Bayer, 2017) together with label-free comparative quantification (Supplementary Fig. 1a). Plasmodesmata and likely contaminant fractions, namely the PM, microsomal, total cell and cell wall fractions were purified from six-day old Arabidopsis suspension culture cells and simultaneously analysed by liquid-chromatography tandem mass-spectrometry (LC-MS/MS). For each protein identified, its relative enrichment in the plasmodesmata fraction versus “contaminant” fractions was determined (Supplementary Fig. 1b; Supplementary Table 1). Enrichment ratios for selecting plasmodesmal-candidates was set based on previously characterised plasmodesmal proteins (see M&M for details). This refined proteome dataset was reduced to 115 unique proteins, cross-referenced with two published ER-proteomes (Nikolovski et al, 2012; Dunkley et al, 2006) and used as a basis for selecting MCS-relevant candidates.
Alongside, we also analysed changes in protein abundance during the ER-PM tethering transition from very tight contacts in post-cytokinetic plasmodesmata (type I) to larger ER-PM gap and sparse tethers in mature plasmodesmata (type II) (Nicolas et al, 2017a). For this we obtained a similar semi-quantitative proteome from four and seven-day old culture cells, enabling a comparison of plasmodesmata composition during the tethering transition (Nicolas et al, 2017a) (Supplementary Fig. 2).
A survey of our refined proteome identified several members of the Multiple C2 domains and Transmembrane region Proteins (MCTPs) family, namely AtMCTP3-7, 9, 10, 14-16, as both abundant and highly enriched at plasmodesmata (Supplementary Fig. 1b, Supplementary Table 1). In addition to being “core” plasmodesmata-associated proteins, our data also suggests that MCTPs are differentially regulated during the ER-PM tethering transition from post-cytokinetic to mature plasmodesmata (Nicolas et al, 2017a) (Supplementary Fig. 2). Amongst the 47 plasmodesmal proteins differentially enriched, all MCTPs were more abundant (1.4 to 3.6 times) in type I (tight ER-PM contacts) compared with type II (open cytoplasmic sleeves) plasmodesmata (Supplementary Fig. 2).
MCTPs are ER-associated proteins that stably cluster at plasmodesmata and present structural features of membrane tethers
MCTPs are structurally reminiscent of the ER-PM tether families of mammalian extended-Synaptotagmins (HsE-Syts) and plant Arabidopsis Synaptotagmins (AtSYTs) (Pérez-Sancho et al, 2015b; Giordano et al, 2013), possessing lipid-binding C2 domains at one end and multiple transmembrane domains (TMDs) at the other, a domain organization consistent with the function of membrane tethers (Supplementary Fig. 3). Unlike HsE-Syts and AtSYTs, the transmembrane region of MCTPs is located at the C-terminus and three to four C2 domains at the N-terminus (Fig. 1a; Supplementary Fig. 3). Two members of the Arabidopsis MCTP family, AtMCTP1/Flower locus T Interacting Protein (FTIP) and AtMCTP15/QUIRKY (QKY) have previously been localised to plasmodesmata in Arabidopsis and implicated in cell-to-cell trafficking of developmental signals (Vaddepalli et al, 2014; Liu et al, 2012). However, two recent studies indicate that other MCTP members, including AtMCTP3, 4, 9, which show high plasmodesmata-enrichment in our proteome, do not associate with the pores in vivo (Liu et al, 2017, 2018).
We investigated the in vivo localisation of MCTPs identified in our proteomic screen by transiently expressing N-terminal fusions fluorescent proteins in Nicotiana benthamiana leaves. As the MCTP family is conserved in N. benthamiana (Supplementary Fig. 4) and to avoid working in a heterologous system we also examined the localisation of NbMCTP7, whose closest homolog in Arabidopsis was also identified as highly-enriched in plasmodesmata fractions (AtMCTP7; Supplementary Fig. 1). Confocal imaging showed that all selected MCTPs, namely AtMCTP3, 4, 6, 9 and NbMCTP7, displayed a similar subcellular localisation, with a faint ER-like network at the cell surface and a punctate distribution along the cell periphery at sites of epidermal cell-to-cell junctions (Fig. 1b, c). Time-lapse imaging showed that peripheral fluorescent punctae were immobile, which contrasted with the high mobility of the ER-like network (Supplementary Mov. 1). Colocalisation with RFP-HDEL confirmed MCTPs association with the cortical ER, while the immobile spots at the cell periphery perfectly co-localised with plasmodesmal markers (mCherry-PDCB1; (Simpson et al, 2009; Grison et al, 2015); Fig. 1c). Co-labelling with general ER-PM tethers such as VAP27.1-RFP and SYT1-RFP (Pérez-Sancho et al, 2015a; Wang et al, 2014), showed partial overlap with GFP-NbMCTP7, while co-localisation with mCherry-PDCBl was significantly higher (Supplementary Fig. 5). To further quantify and ascertain MCTP association with plasmodesmata, we measured a plasmodesmal enrichment ratio, hereafter named “plasmodesmata-index”. For this we calculated fluorescence intensity at plasmodesmata pit-fields (indicated by mCherry-PDCBl or aniline blue) versus cell periphery. All MCTPs tested displayed a high plasmodesmata-index, ranging from 1.85 to 4.15, similar to PDLP1 (1.36) and PDCB1 (1.45) two-well established plasmodesmata markers (Thomas et al, 2008b; Simpson et al, 2009) (Fig. 1d), confirming enrichment of MCTPs at pit-fields. When stably expressed in Arabidopsis thaliana under the moderate promoter UBIQUITIN 10 or 35S promoters AtMCTP4, AtMCTP6 and AtMCTP9 were found mainly restricted to plasmodesmata (Supplementary Fig. 6a, white arrows), as indicated by an increase of their plasmodesmata-index compared with transient expression in N. benthamiana (Supplementary Fig. 6b). A weak but consistent ER localisation was also visible in stably transformed Arabidopsis (Supplementary Fig. 6a red stars).
To get a better understanding of MCTP distribution within the plasmodesmal pores, we further analysed transiently-expressed GFP-NbMCTP7 by 3D structured illumination superresolution microscopy (3D-SIM) (Fitzgibbon et al, 2010) (Fig. 1e). We found that NbMCTP7 is associated with all parts of plasmodesmata including the neck regions and central cavity, as well as showing continuous fluorescence throughout the pores. In some cases, lateral branching of plasmodesmata within the central cavity was resolved. The very faint continuous fluorescent threads connecting neck regions and central cavity correspond to the narrowest regions of the pores and may indicate association with the central desmotubule (Fig. 1e, white arrows).
Using Fluorescence Recovery After Photobleaching (FRAP) we then assessed the mobility of NbMCTP7. We found that, when associated with the cortical ER the fluorescence recovery rate of GFP-NbMCTP7 was extremely fast and similar to RFP-HDEL with half-times of 1.16 sec and 0.99 sec, respectively (Fig. 2). By contrast, when GFP-NbMCTP7 was associated with plasmodesmata, the recovery rate slowed down to a half-time of 4.09 sec, indicating restricted mobility, though still slightly faster than for the cell wall-localised plasmodesmal marker mCherry-PDCB1 (5.98 sec). Overall, these results show that NbMCTP7 mobility is high at the cortical ER but becomes restricted inside the pores.
From our data we concluded that MCTPs are ER-associated proteins, whose members specifically and stably cluster at plasmodesmata. They display the structural features required for ER-PM tethering and are differentially associated with the pores during the transition in ER-PM contacts.
AtMCTP4 is a core plasmodesmata-associated protein and loss-of-function mctp3/mctp4 double mutants show pleiotropic developmental defects
We next focused on AtMCTP4, which according to our proteomic screen qualifies as a core plasmodesmal constituent considering that it is one of the most abundant proteins in our refined proteome (Supplementary Table 1). The implication of AtMCTP4 association with plasmodesmata is that the protein contributes functionally to cell-to-cell signalling. Given the importance of plasmodesmata in tissue patterning and organ growth, a loss-of-function mutant is expected to show defects in plant development. We first obtained T-DNA insertion lines for AtMCTP4 and its closest homolog AtMCTP3, which share 92.8% identity and 98.7% similarity in amino acids with AtMCTP4, but both single knockouts showed no apparent phenotypic defects (data not shown). We therefore generated an Atmctp3/Atmctp4 double mutant, which presented pleiotropic developmental defects with a severely dwarfed and bushy phenotype, twisted leaves with increased serration (Fig. 3 a-d), and multiple inflorescences (not shown). Whilst preparing this manuscript another paper describing the Atmctp3/Atmctp4 mutant was published (Liu et al, 2018), reporting similar developmental defects. We noted additional phenotypic defects in particular aberrant pattern in the root apical meristem organisation specifically within the quiescent center (QC) (Fig. 3e). Instead of presenting the typical four-cell layer organisation, we observed aberrant cell division pattern in Atmctp3/Atmctp4 mutant with asymmetrical division in the QC, suggesting that both proteins may play a general role in cell stem niche maintenance (Liu et al, 2018).
AtMCTP4 has recently been reported as an endosomal-localised protein (Liu et al, 2018), which is in conflict with our data indicating plasmodesmata association. To further check AtMCTP4 localisation, we expressed the protein as a GFP N-terminal fusion protein under its own promoter and analysed its localisation in Arabidopsis stable lines. Similar to transient expression experiments (Fig. 1), we found that pMCTP4:GFP-AtMCTP4 was located at stable punctate spots at the cell periphery (Fig. 3f white arrows; Suppl movie 2), in all tissues examined, i.e. leaf epidermal and spongy mesophyll cells, hypocotyl epidermis, lateral root primordia, root tip, and inflorescence shoot apical meristem. These immobile dots colocalised perfectly with aniline blue indicating plasmodesmata association (Fig.3f top row), which was also evident in leaf spongy mesophyll cells where the dotty pattern of pMCTP4:GFP-AtMCTP4 was present on adjoining walls (containing plasmodesmata), but absent from non-adjoining walls (without plasmodesmata) (Fig.3f white arrowheads). We also observed a weak but consistent ER-association of AtMCTP4 (Fig.3f, red stars).
In summary we concluded that whatever the tissue and organ considered, AtMCTP4 is strongly and consistently associated with plasmodesmata but also presents a steady association with the ER and that Atmctp3/Atmctp4 loss-of-function is detrimental for normal plant development, including previously uncovered defect in the root apical meristem.
The C-terminal transmembrane regions of MCTPs serve as ER-anchors
A requirement for tethers is that they physically bridge two membranes. Often this is achieved through lipid-binding module(s) at one terminus of the protein, and transmembrane domain(s) at the other (Eisenberg-Bord et al, 2016; Henne et al, 2015). All sixteen Arabidopsis MCTPs contain two to three predicted TMDs near their C-terminus (collectively referred to as the transmembrane region, TMR). To test whether the MCTP TMRs are determinants of ER-insertion, we generated truncation mutants lacking the C2 domains for NbMCTP7, AtMCTP3, AtMCTP4, AtMCTP6, AtMCTP9 as well as AtMCTP1/FTIP and AtMCTP15/QKY (Fig. 4a). When fused to YFP at their N-terminus, all truncated mutants retained ER-association, as demonstrated by co-localisation with RFP-HDEL (Fig. 4b left panels). Meanwhile plasmodesmata association was completely lost and the plasmodesmata-index of all truncated MCTP_TMRs dropped below one, comparable to RFP-HDEL (Fig. 4b right panels and c), quantitatively confirming the loss of plasmodesmata association when the C2 modules were deleted. We therefore concluded that, similar to the HsE-Syt and AtSYT ER-PM tether families (Giordano et al, 2013; Levy et al, 2015; Pérez-Sancho et al, 2015b), MCTPs insert into the ER through their TMRs, but the TMR is not sufficient for MCTP plasmodesmal localisation.
MCTP C2 domains can bind membranes in an anionic lipid-dependent manner
Members of the HsE-Syt and AtSYT tether families bridge across the intermembrane gap and dock to the PM via their C2 domains (Pérez-Sancho et al, 2015b; Pérez-sancho et al, 2016; Giordano et al, 2013; Saheki et al, 2016). Arabidopsis MCTPs contain three to four C2 domains, which may also drive PM-association through interactions with membrane lipids. C2 domains are independently folded structural and functional modules with diverse modes of action, including membrane docking, protein-protein interactions and calcium sensing (Corbalan-Garcia & Gómez-Fernández, 2014).
To investigate the function of MCTP C2 modules, we first searched for homologs of AtMCTP individual C2 domains (A, B, C, and D) amongst all human and A. thaliana proteins using the HHpred webserver (Zimmermann et al, 2018) for remote homology detection. The searches yielded a total of 1790 sequence matches, which contained almost all human and A. thaliana C2 domains. We next clustered the obtained sequences based on their all-against-all pairwise similarities in CLANS (Frickey & Lupas, 2018). In the resulting map (Supplementary Fig. 7a), the C2 domains of Arabidopsis MCTPs (AtMCTPs, coloured cyan) most closely match the C2 domains of membrane-trafficking and -tethering proteins, including human MCTPs (HsMCTPs, green), human Synaptotagmins (HsSyts, orange), human Ferlins (HsFerlins, blue), human HsE-Syts (HsE-Syts, magenta) and Arabidopsis SYTs (AtSYTs, red), most of which dock to membranes through direct interaction with anionic lipids (Giordano et al, 2013; Saheki et al, 2016; Pérez-Lara et al, 2016; Abdullah et al, 2014; Marty et al, 2014). By comparison to the C2 domains of these membrane-trafficking and -tethering proteins, the C2 domains of most other proteins do not make any connections to the C2 domains of AtMCTPs at the P-value cut-off chosen for clustering (1e-10). Thus, based on sequence similarity, the plant AtMCTP C2 domains are expected to bind membranes.
We next asked whether the C2 modules of MCTPs are sufficient for PM association in vivo. Fluorescent protein fusions of the C2A-D or C2B-D modules without the TMR were generated for NbMCTP7, AtMCTP3, AtMCTP4, AtMCTP6, AtMCTP9 as well as AtMCTP1/FTIP and AtMCTP15/QKY and expressed in N. benthamiana. We observed a wide range of sub-cellular localisations from cytosolic to PM-associated and in all cases plasmodesmata association was lost (Supplementary Fig. 7b-d).
To further investigate the potential for MCTP C2 domains to interact with membranes, we employed molecular dynamics modelling. We focussed on AtMCTP4, as a major plasmodesmal constituent and whose loss-of-function in conjunction with AtMCTP3, induces severe plant development defects (Liu et al, 2018) (Fig. 3). We first generated the 3D structures of all three C2 domains of AtMCTP4 using 3D homology modelling, and then tested the capacity of individual C2 to dock to membrane bilayers using coarse-grained dynamic simulations (Fig. 5a; Suppl movie 3). Molecular dynamics modelling was performed on three different membranes; 1) a neutral membrane composed of phosphatidylcholine (PC), 2) a membrane with higher negative charge composed of PC and phosphatidylserine (PS; 3:1) and 3) a PM-mimicking lipid bilayer, containing PC, PS, sitosterol and the anionic phosphoinositide phosphatidyl inositol-4-phosphate (PI4P; 57:19:20:4). The simulations showed that all individual C2 domains of AtMCTP4 can interact with lipids and dock on the membrane surface when a “PM-like” lipid composition was used (Fig. 5a). The PC-only membrane showed only weak interactions, whilst the PC:PS membrane allowed only partial docking (Fig. 5a). Docking of AtMCTP4 C2 domains arose mainly through electrostatic interactions between lipid polar heads and basic amino acid residues at the protein surface. We further tested two other MCTP members, namely AtMCTP15/QKY and NbMCTP7, which possess four rather than three C2 domains. We found that similar to AtMCTP4, the individual C2 domains of AtMCTP15/QKY and NbMCTP7 exhibited membrane interaction in the presence of the negatively charged lipids (Supplementary Fig. 8).
Our molecular dynamics data thus suggests that membrane docking of the AtMCTP4 C2 domains depends on the electrostatic charge of the membrane and more specifically on the presence of PI4P, a negatively-charged lipid which has been reported as controlling the electrostatic field of the PM in plants (Simon et al, 2016).
To confirm the importance of PI4P for MCTP membrane interactions and thus, potentially subcellular localisation, we used a short-term treatment with phenylarsine oxide (PAO), an inhibitor of PI4-kinases (Simon et al, 2016). We focused on Arabidopsis root tips where effects of PAO have been thoroughly characterised (Simon et al, 2016). In control-treated roots of Arabidopsis plants stably expressing UB10:YFP-AtMCTP4, the fluorescent signal was most prominent at the apical-basal division plane of epidermal root cells, where numerous plasmodesmata are established during cytokinesis (Grison et al, 2015) (Fig. 5b white arrowheads). The YFP-AtMCTP4 fluorescence pattern was punctate at the cell periphery, each spot of fluorescence corresponding to a single or group of plasmodesmata (Fig. 5c, white arrows). We found that 40 min treatment with PAO (60 μM) induced a loss in the typical spotty plasmodesmata-associated pattern and instead AtMCTP4 became more homogenously distributed along the cell periphery (Fig.5b-c). To confirm the effect of PAO on the cellular PI4P pool, we used a PI4P-biosensor (1XPH FAPP1) which showed a clear shift from PM-association to cytosolic localisation upon treatment with PAO (Simon et al, 2016) (Fig. 5b). This control not only demonstrates that the PAO treatment was successful, but also highlights that the majority of PI4P was normally found at the PM, rather than the ER, of Arabidopsis root cells. Therefore, the effect of PAO on YFP-AtMCTP4 localisation is likely related to a perturbance of PM docking by the MCTP4 C2 domains.
Altogether, our data suggest that the C2 domains of plant MCTPs can dock to membranes in the presence of negatively charged phospholipids, and that PI4P depletion reduced AtMCTP4 stable association with plasmodesmata.
AtMCTP4 expression is sufficient to partially restore ER-PM contacts in yeast
To further test the ability of MCTPs to physically bridge across membranes and tether the ER to the PM, we used a yeast Δtether mutant line deleted in six ER-PM tethering proteins resulting in the separation of the cortical ER (cER) from the PM (Manford et al, 2012) and expressed untagged AtMCTP4. To monitor recovery in cortical ER, and hence, ER-PM contacts, upon AtMCTP4 expression, we used Sec63-RFP (Metzger et al, 2008) as an ER marker combined with confocal microscopy. In wild-type cells, the ER was organised into nuclear (nER) and cER. The cER was visible as a thread of fluorescence along the cell periphery, covering a large proportion of the cell circumference (Fig. 6a white arrows). By contrast and as previously reported (Manford et al, 2012), we observed a substantial reduction of cER in the Δtether mutant, with large areas of the cell periphery showing virtually no associated Sec63-RFP (Fig. 6a). When AtMCTP4 was expressed into the Δtether mutant line, we observed partial recovery of cER, visible as small regions of Sec63-RFP closely apposed to the cell cortex. We further quantified the extent of cER in the different lines by measuring the ratio of the length of cER (Sec63-RFP) against the cell perimeter (through calcofluor wall staining) and confirmed that ATMCTP4 expression induced an increase of cER from 7.3 % to 23.1% when compared to the Δtether mutant (Fig. 6b). This partial complementation is similar to results obtained with yeast deletion mutants containing only a single endogenous ER-PM tether, IST2, or all three isoforms of the tricalbin (yeast homologs of HsE-Syts) (Manford et al, 2012), supporting a role of AtMCTP4 as ER-PM tether.
Discussion
In plants, communication between cells is facilitated and regulated by plasmodesmata, ~50 nm diameter pores that span the cell wall and provide cell-to-cell continuity of three different organelles: the PM, cytoplasm, and ER. The intercellular continuity of the ER and the resulting architecture of the pores make them unique amongst eukaryotic cellular junctions, and qualify plasmodesmata as a specialised type of ER-PM MCS (Bayer et al, 2017; Tilsner et al, 2016). Like other types of MCS, the membranes within plasmodesmata are physically connected but so far the molecular components and function of the ER-PM tethering machinery remain an enigma.
Here, we provide evidence that members of the MCTP family, some of which have been described as key regulators of intercellular trafficking and cell-to-cell signalling (Vaddepalli et al, 2014; Liu et al, 2018, 2012), also act as ER-PM tethers inside the plasmodesmata pores.
MCTPs as core plasmodesmal components
Whi1st several MCTPs have previously been characterised as regulators of cell-to-cell trafficking or signalling (Liu et al, 2012, 2018; Vaddepalli et al, 2014; Liu et al, 2017), only some have also been localised to plasmodesmata, Whilst other studies reported alternative localisations which include PM, ER, Golgi, endosomes and cytoplasm (Trehin et al, 2013; Liu et al, 2017, 2018, 2012; Kraner et al, 2017; Vaddepalli et al, 2014), perhaps depending on the isoform, orientation of fluorescent protein fusions and expression context. Here, we have identified several MCTPs (6-10 out of 16) as belonging to the most abundant proteins at plasmodesmata through both in vivo and proteomic data. This includes AtMCTP3 and AtMCTP4 recently identified as modulators of SHOOTMERISTEMLESS trafficking (Liu et al, 2018), for which we find that a Atmctp3/Atmctp4 loss-of-function mutant displays a severe developmental phenotype, including defects in the root QC, that agrees with the findings of Liu et al, 2018. Whereas Liu et al. (2018) observed endosomal-localisation of AtMCTP3 and AtMCTP4, our data suggest they are primarily plasmodesmata-associated. We therefore propose that MCTPs are core plasmodesmata-constituents and that AtMCTP3 and AtMCTP4 may possibly regulate the transport of SHOOTMERISTEMLESS, at the pores.
MCTPs as plasmodesmata-specific ER-PM tethers
While ER-PM contacts within plasmodesmata have been observed for decades (Ding et al, 1992; Tilsner et al, 2011; Tilney et al, 1991; Nicolas et al, 2017b), the molecular identity of the tethers has remained elusive. Here we propose that MCTPs are prime plasmodesmal membrane tethering candidates as they possess all required features: 1) strong association with plasmodesmata; 2) structural similarity to known ER-PM tethers such as HsE-Syts and AtSYTs (Levy et al, 2015; Pérez-Sancho et al, 2015b; Giordano et al, 2013) possessing an ER-inserted TMR at one end and multiple lipid-binding C2 domains at the other for PM docking; 3) the ability to partially restore ER-PM tethering in a yeast Δtether mutant.
Similarly to other ER-PM tethers (Eisenberg-Bord et al, 2016; Wong et al, 2016; Henne et al, 2015; Giordano et al, 2013), MCTP C2 domains dock to the PM through electrostatic interaction with anionic lipids, especially PI4P and to a lesser extent PS. In contrast with animal cells, PI4P is found predominantly at the PM in plant cells and defines its electrostatic signature (Simon et al, 2016). Although plasmodesmata are MCS, they are also structurally unique: both the ER and the PM display extreme, and opposing membrane curvature inside the pores; the ER tubule is linked to the PM on all sides; and the membrane apposition is unusually close (2-3 nm in type I post-cytokinetic pores (Nicolas et al, 2017a)). So while structurally related to known tethers, MCTPs are also expected to present singular properties. For instance, similar to the human MCTP2, MCTPs could favour ER membrane curvature through their TMR (Joshi et al, 2017). Plasmodesmata also constitute a very confined environment, which together with the strong negative curvature of the PM, may require the properties of MCTP C2 domains to differ from that of HsE-Syts or AtSYTs. All of these aspects will need to be investigated in the future.
Inter-organellar signalling at the plasmodesmal MCS?
In yeast and animals, MCS have been shown to be privileged sites for inter-organelle signalling by promoting fast, non-vesicular transfer of molecules such as lipids (Gallo et al, 2016; Saheki et al, 2016; Wong et al, 2016). Unlike the structurally analogous tethering proteins AtSYTs and HsE-Syts, MCTPs do not harbour known lipid-binding domains that would suggest that they participate directly in lipid transfer between membranes. However, MCTPs are likely to act in complex with other proteins (Fulton et al, 2009; Trehin et al, 2013) which may include lipid shuttling proteins. For instance, AtSYTl, which contains a lipid-shuttling SMP (synaptotagmin-like mitochondrial-lipid binding protein) domain (Reinisch & De Camilli, 2016), is recruited to plasmodesmata during virus infection and promotes virus cell-to-cell movement (Levy et al, 2015). MCS tethers typically interact with other MCS components and locally regulate their activity, act as Ca2+ sensors, or modulate membrane spacing to turn lipid shuttling on or off (Eden et al, 2010, 2016; Ho et al, 2016; Chang et al, 2013; Kim et al, 2015; Giordano et al, 2013; Idevall-Hagren et al, 2015a; Petkovic et al, 2014; Zhang et al, 2005; Fernàndez-Busnadiego et al, 2015; Omnus et al, 2016; Saheki et al, 2016). Similar activities could be performed by MCTPs at plasmodesmata. To date however, ER-PM cross-talk at plasmodesmata remains hypothetical.
Combining organelle tethering and cell-to-cell signalling functions
Several members of the MCTP family have previously been implicated in regulating either macromolecular trafficking or intercellular signalling through plasmodesmata. AtMCTP1/FTIP interacts with, and is required for phloem entry of the Flowering Locus T (FT) protein, triggering transition to flowering at the shoot apical meristem (Liu et al, 2012). Similarly, AtMCTP3/AtMCTP4 regulate trafficking of SHOOTMERISTEMLESS in the shoot apical meristem, however in this case they prevent cell-to-cell trafficking (Liu et al, 2018). AtMCTP15/QKY promotes the transmission of an unidentified non-cell-autonomous signal through interaction with the plasmodesmata/PM-located receptor-like kinase STRUBBELIG (Vaddepalli et al, 2014). Thus, previously characterised MCTP proteins regulate intercellular trafficking/signalling either positively or negatively.
Whilst the mechanisms by which these MCTP proteins regulate intercellular transport/signalling have not been elucidated, MCTP physical interaction with mobile factors or receptor is critical for proper function (Vaddepalli et al, 2014; Liu et al, 2017, 2018, 2012). In AtMCTP1/FTIP, the interaction is mediated by the C2 domain closest to the TMR (Liu et al, 2017). For the C2 domains of HsE-Syts, conditional membrane docking is critical for their function and depends on intramolecular interactions, cytosolic Ca2+ and the presence of anionic lipids (Idevall-Hagren et al, 2015b; Fernàndez-Busnadiego et al, 2015; Saheki et al, 2016; Bian et al, 2018; Giordano et al, 2013). With three to four C2 domains, it is conceivable that MCTPs assume different conformations within the cytoplasmic sleeve in response to changes in the plasmodesmal PM composition, Ca2+, and the presence of interacting mobile signals (Fig.7), which could link membrane tethering to cell-cell signalling. Understanding in detail how MCTPs function in the formation and regulation of the plasmodesmal MCS will be an area of intense research in the coming years.
Material & Methods
Biological material and growth conditions
Arabidopsis (Columbia) and transgenic lines were grown vertically on solid medium composed of Murashige and Skoog (MS) medium including vitamins (2.15g/L), MES (0.5g/L) and plant-Agar (7g/L), pH 5.7, then transferred to soil under long-day conditions at 22 °C and 70% humidity.
Arabidopsis (Landsberg erecta) culture cells were cultivated as described in(Nicolas et al, 2017a) under constant light (20μE/m/s) at 22°C. Cells were used for experimentation at various ages ranging from four to seven-day-old (mentioned in individual experiment).
MCTP sequence alignment and phylogenetic tree
The 16 members of Arabidopsis thaliana MCTP family, gathering a total of 59 C2 domains, were dissected using a combination of several bioinformatic tools. The alignment of A. thaliana MCTP members from(Liu et al, 2017) combined with Pfam predictions was used as a first step to segregate the MCTP members into “sub-families”: the short MCTPs, which contain three C2 domains (C2B to C2D) and the long MCTPs, which contain four C2 domains (C2A to C2D). The short MCTPs lack the C2A domain, whereas the C2B-C-D are conserved in all members.
The prediction and delimitation of C2 domains in proteins, including MCTPs, from databases such as Pfam are rather imprecise. In order to provide stronger and more accurate predictions for the delimitation of each C2 domain, we used both the PSIRED(Buchan et al, 2013; Jones, 1999) protein sequence analysis (http://bioinf.cs.ucl.ac.uk/psipred/) and Hydrophobic Cluster Analysis(Callebaut et al, 1997) (HCA; http://www-ext.impmc.upmc.fr/~callebau/HCA.html). Multiple sequence alignment was performed using Clustal Omega (http://www.ebi.ac.uk/Tools/msa/clustalo/).
Cluster map of Human and A. thaliana C2 domains
To generate a C2 cluster map, we first collected all A. thaliana and human C2 domains, using the HHpred webserver(Alva et al, 2016; Söding et al, 2005). The obtained set was filtered to a maximum of 100% pairwise sequence identity at a length coverage of 70% using MMseqs2(Steinegger & Söding, 2017) to eliminate all redundant sequences. The sequences in the filtered set, comprising almost all human and A. thaliana C2 domains (~1800 in total), was next clustered in CLANS(Frickey & Lupas, 2018) based on their all-against-all pairwise sequence similarities as evaluated by BLAST P-values.
Cloning of MCTPs and transformation into Arabidopsis
The different constructs used in this study were either PCR amplified from cDNA or genomic DNA (Col-0) using gene specific primers (Supplementary Table S2), or were synthesised and cloned into donor vectors by GenScript® (Supplementary Table S2). For N-terminal tag fusion, the PCR/DNA products were cloned into the Multisite Gateway® donor vectors pDONR-P2RP3 (Invitrogen, Carlsbad, CA), and then subcloned into pB7m34GW or pK7m34GW using the multisite LR recombination system(Karimi et al, 2002), the moderate promoter UBIQUITIN10 (UBQ10/pD0NR-P4P1R previously described in(Marquès-Bueno et al, 2016)) and eYFP/pD0NR221. For C-terminal tag fusion, the PCR/DNA products were first cloned into pD0NR221, then multisite recombined using a mVenus/pD0NR-P2RP3 and UBQ10/pD0NR-P4P1R.
For the expression of GFP-MCTP4 driven by its native promotor we used the binary vector pRBbar-OCS harboring a BASTA resistance, a multiple cloning side (MCS) and an octopine synthase (OCS) terminator within the left and right borders. The vector derived from the pB2GW7 (Karimi et al, 2002) by cutting out the expression cassette with the restriction enzymes SacI and HindIII and replaced it with a synthesized MCS and an OCS terminator fragment. To combine promoter region and GFP-MCTP4 coding sequence we used In-Fusion cloning (Takara Bio Europe). To PCR amplify the coding sequence for GFP-MCTP4 with its respective primers (Supplementary Table2) we used the plasmid coding for GFP-MCTP4 as template (previously described as GFP-C2-89 by (Kraner et al, 2017)). The resulting pRBbar-pMCTP4: plasmid was linearized with BamH1/Pst1 the amplified GFP-MCTP4 was fused in to generate the MCTP4 promoter driven GFP-MCTP4 construct (pMCTP4:GFP-MCTP4). Expression vectors were transformed in Arabidopsis Col-0 by floral dip(Clough & Bent, 1998), and transformed seeds were selected based on plasmid resistance.
N. benthamiana homologs of Arabidopsis MCTP isoforms were identified by protein BLAST searches against the SolGenomics N. benthamiana genome (https://solgenomics.net). An ortholog of AtMCTP7, NbMCTP7 (Niben101Scf03374g08020.1) was amplified from N. benthamiana leaf cDNA. The recovered cDNA of NbMCTP7 differed from the SolGenomics reference by the point mutation G287D and three additional silent nucleotide exchanges, as well as missing base pairs 1678-1716 which correspond to thirteen in-frame codons (encoding the amino acid sequence LKKEKFSSRLHLR). We note that this nucleotide and amino acid sequence is exactly repeated directly upstream (bp 1639-1677) in the SolGenomics reference and may thus represent an error in the N. benthamiana genome assembly. The recovered NbMCTP7 sequence has been submitted to database.
Generation of Atmctp3/Atmctp4 loss-of-function Arabidopsis mutant
Atmctp3 (Sail_755_G08) and Atmctp4 (Salk_089046) T-DNA insertional Arabidopsis mutants (background Col-0) were obtained from the Arabidopsis Biological Resource Center (http://www.arabidopsis.org/). Single T-DNA insertion lines were genotyped and homozygous lines were crossed to obtain double homozygous Atmctp3/Atmctp4.
For genotyping, genomic DNA was extracted from Col-0, Atmctp3 (GABI-285E05) and Atmctp4 (SALK-089046) plants using chloroform:isoamyl alcohol (ratio24:1), genomic DNA isolation buffer (200mM Tris HCL PH7.5, 250mM NaCl, 25mM EDTA and 0.5% SDS) and isopropanol. PCR were performed with primers indicated in Supplementary Table2. For transcript expression, total mRNA was extracted from Col-0 and Atmctp3/Atmctp4 using RNeasy® Plant Mini Kit (QIAGEN) and cDNA was produced using random and oligodT primers. The expression level of AtMCTP3, AtMCTP4 and ubiquitous Actin2 (ACT2) transcript was tested by PCR amplification using primers listed in Supplementary Table2.
Confocal Laser Scanning Microscopy
For transient expression in N. benthamiana, leaves of 3 week-old plants were pressure-infiltrated with GV3101 agrobacterium strains, previously electroporated with the relevant binary plasmids. Prior to infiltration, agrobacteria cultures were grown in Luria and Bertani medium with appropriate antibiotics at 28°C for two days then diluted to 1/10 and grown until the culture reached an OD600 of about 0.8. Bacteria were then pelleted and resuspended in water at a final OD600 of 0.3 for individual constructs, 0.2 each for the combination of two. The ectopic silencing suppressor 19k was co-infiltrated at an OD600 of 0.15. Agroinfiltrated N. benthamiana leaves were imaged 3-4 days post infiltration at room temperature. ~ 2 by 2 cm leaf pieces were removed from plants and mounted with the lower epidermis facing up onto glass microscope slides.
Transgenic Arabidopsis plants were grown as described above. For primary roots, lateral roots and hypocotyl imaging, six to seven days old seedlings or leaves of 5-8 leaf stage rosette plants were mounted onto microscope slides. For shoot apical meristem imaging, the plants were first dissected under a binocular then transferred to solid MS media and immediately observed using a water-immersion long-distance working 40X water immersion objective. Confocal imaging was performed on a Zeiss LSM 880 confocal laser scanning microscope equipped with fast AiryScan using Zeiss C PL APO x63 oil-immersion objective (numerical aperture 1.4). For GFP, YFP and mVenus imaging, excitation was performed with 2-8% of 488 nm laser power and fluorescence emission collected at 505-550 nm and 520-580 nm, respectively. For RFP and mCherry imaging, excitation was achieved with 2-5% of 561 nm laser power and fluorescence emission collected at 580-630 nm. For aniline blue (infiltrated at the concentration of 25 μg/mL) and Calcofluor White (1 μg /mL), excitation was achieved with 5% of 405 nm laser and fluorescence emission collected at 440-480 nm. For colocalisation sequential scanning was systematically used.
For quantification of NbMCTP7 co-localisation with VAP27.1, SYT1 and PDCB1, coexpression of the different constructs was done in N. benthamiana. An object based method was used for colocalization quantification(Bolte & Cordelières, 2006). Images from different conditions are all acquired with same parameters (zoom, gain, laser intensity etc.) and channels are acquired sequentially. These images are processed and filtered using ImageJ software (https://imagei.nih.gov/ij/) in order to bring out the foci of the pictures. These foci were then automatically segmented by thresholding and the segmented points from the two channels were assessed for colocalization using the ImageJ plugin Just Another Colocalization Plugin (JACoP)(Bolte & Cordelières, 2006). This whole process was automatized using a macro (available upon demand).
Pseudo-Schiff-Propidium iodide stained Arabidopsis root tips was performed according to(Truernit et al, 2008). Aniline blue staining was performed according to(Grison et al, 2015). Brightness and contrast were adjusted on ImageJ software (https://imagej.nih.gov/ij/).
Plasmodesmata (PD) index
Plasmodesmata depletion or enrichment was assessed by calculating the fluorescence intensity of GFP/YFP-tagged full-length MCTP, truncated MCTPs and the proton pump ATPase GFP-PMA2(Gronnier et al, 2017), at 1) plasmodesmata (indicated by mCHERRY-PDCB1, PDLPl-mRFP or aniline blue) and 2) at the cell periphery (i.e. outside plasmodesmata pitfields). For that, confocal images of leaf epidermal cells (N. benthamiana or Arabidopsis) were acquired by sequential scanning of mCHERRY-PDCB, PDLP1-mRFP or aniline blue (plasmodesmata markers) in channel 1 and GFP/YFP-tagged MCTPs in channel 2 (for confocal setting see above). About thirty images of leaf epidermis cells were acquired with a minimum of three biological replicates. Individual images were then processed using ImageJ by defining five regions of interest (ROI) at plasmodesmata (using plasmodesmata marker to define the ROI in channel1) and five ROIs outside plasmodesmata. The ROI size and imaging condition were kept the same. The GFP/YFP-tagged MCTP mean intensity (channel 2) was measured for each ROI then averaged for single image. The plasmodesmata index corresponds to intensity ratio between fluorescence intensity of MCTPs at plasmodesmata versus outside the pores. For the plasmodesmata-index of RFP-HDEL, PDLP1-RFP and mCHERRY-PDCB1 we used aniline to indicate pitfields. R software was used for making the box plots and statistics.
FRAP analysis
For FRAP analysis, GFP-NbMCTP7, RFP-HDEL and mCHERRY-PDCB1-expressing N. benthamiana leaves were used. The experiments were performed on a Zeiss LSM 880 confocal microscope equipped with a Zeiss C PL APO x63 oil-immersion objective (numerical aperture 1.4). GFP and mCherry were respectively excited at 488nm and 561nm with 2% of Argon or DPSS 561-10 laser power, and fluorescence was collected with the GaAsp detector at 492-569nm and 579-651nm, respectively. To reduce as much as possible scanning time during FRAP monitoring, the acquisition window was cropped to a large rectangle of 350 by 50 pixels, with a zoom of 2.7 and pixel size of 0.14μm. By this mean, pixel dwell time was of 0.99μs and total frame scan time could be reduced down to 20 ms approximately. Photobleaching was performed on rectangle ROIs for the ER-network and on circle ROIs for the pitfields with the exciting laser wavelengths set to 100%. The FRAP procedure was the following: 30 pre-bleach images, 10 iterations of bleaching with a pixel dwell time set at 1.51 μs and then 300 images post-bleach with the “safe bleach mode for GaAsp”, bringing up the scan time up to approximately 200ms. The recovery profiles were background substracted and then double normalized (according to the last prebleach image and to the reference signal, in order to account for observational photobleaching) and set to full scale (last pre-bleach set to 1 and first post-bleach image set to 0), as described by Kote Miura in his online FRAP-teaching module (EAMNET-FRAP module, https://embl.de). Plotting and curve fitting was performed on GraphPad Prism (GraphPad Software, Inc.).
3D-SIM imaging
For 3D structured illumination microscopy (3D-SIM), an epidermal peal was removed from a GFP-NbMCTP7-expressing leaf and mounted in Perfluorocarbon PP11(Littlejohn et al, 2014) under a high precision (170mm+/-5mm) coverslip (Marie Enfield). The sample chamber was sealed with non-toxic Exaktosil N 21 (Bredent, Germany). 3D-SIM images were obtained using a GE Healthcare / Applied Precision OMX v4 BLAZE with a 1.42NA Olympus PlanApo N 60X oil immersion objective. GFP was excited with a 488nm laser and imaged with emission filter 504-552nm (528/48nm). SR images were captured using Deltavison OMX software 3.70.9220.0. SR reconstruction, channel alignment and volume rendering were done using softWoRx V. 7.0.0.
Yeast
Wild-type (SEY6210) and delta-tether yeast strain(Manford et al, 2012) were transformed with Sec63.mRFP (pSM1959). Sec63.mRFP(Metzger et al, 2008) was used as an ER marker and was a gift from Susan Mickaelis (Addgene plasmid #41837). Delta-tether/Sec63.mRFP strain was transformed with AtMCTP4 (pCU416: pCU between SacI and SpeI sites, Cyc1 terminator between XhoI and KpnI sites and AtMCTP4 CDS between BamHI and SmaI sites, Supplementary table S2). Calcofluor White was used to stain the cell wall of yeast. All fluorescent microscopy was performed on midlog cells, grown on selective yeast media (-URA -LEU for AtMCTP4 and Sec63 expression, and -LEU for Sec63). Images were acquired with Airyscan module, using a 63X oil immersion lens and sequential acquisition. Brightness and contrast were adjusted on ImageJ software (https://imagei.nih.gov/ii/).
Supplementary methods
Methods for plasmodesmata label-free proteomic analysis and dynamic modelling are described in details in Supplementary methodl.
Sequence data for genes in this article can be found in the GenBank/EMBL databases using the following accession numbers: AtMCTP1, At5g06850; AtMCTP2, At5g48060; AtMCTP3, At3g57880; AtMCTP4, At1g51570; AtMCTP5, At5g12970; AtMCTP6, At1g22610; AtMCTP7, At4g11610; AtMCTP8, At3g61300; AtMCTP9, At4g00700; AtMCTP10, At1g04150; AtMCTP11, At4g20080; AtMCTP12, At3g61720; AtMCTP13, At5g03435; AtMCTP14, At3g03680; AtMCTP15, At1g74720; AtMCTP16, At5g17980 and NbMCTP7, Niben101Scf03374g08020.1.
Contributions
F. I., M.S.G., M.F. and S.C. carried out the proteomic analysis. M.L.B. cloned the MCTPs, produced and phenotyped the Arabidopsis transgenic lines, with the exception of AtMCTP4:GFP-MCTP4 and 35S:GFP-MCTP6 which were generated by M.K.. M.L.B. and J.D.P. imaged the MCTP reporter lines. W.N. carried out the FRAP analysis and image quantification for co-localisation with the help of L.B.. A.G. performed the phylogenic analysis. J.D.P. carried out the PAO experiments. M.L.B. performed the yeast experiments. T.J.H. and J.T. performed the 3D-SIM. V.A. carried out the C2 cluster map analysis. J.D.P. carried out the molecular dynamic analysis with the help of J-M.C. and L.L..
E.M.B. conceived the study and designed experiments with the help of J.T and L.L. E.M.B, J.D.P., J.T. and M.L.B. wrote the manuscript. All the authors discussed the results and commented on the manuscript.
Competing interests
The authors declare no competing financial interests.
Movie S1. Confocal time lapse imaging of 35S:GFP-NbMCTP7 in N. benthamiana epidermal leaves. One image every 0.2 seconds.
Movie S2. Confocal time lapse imaging of AtMCTP4:GFP-AtMCTP4 in transgenic Arabidopsis epidermal leaves. One image every 0.2 seconds.
Movie S3. Docking of the C2B, C2C and C2D domains of AtMCTP4 on a “PM-like” membrane (see Fig. 5), containing phosphatidylcholine (PC), phosphatidylserine (PS), sitosterol (Sito) and phosphoinositol-4-phosphate (PI4P) in the following ratio: PC/PS/Sito/PI4P 57:19:20:4. Please note that 0.5μs out of total (2.5μs) simulation is shown (moment of docking). The amino acid colour code is as follow: red, negatively charged (acidic) residues; blue, positively charged (basic) residues; green, polar uncharged residues; and white, hydrophobic residues. The lipid colour code is as follow: PC is depicted as light-pink polar heads and grey acyl chains, PS is depicted as dark-pink polar heads and light-purple acyl chains, PI4P is depicted as orange (inositol ring) and yellow (phosphate 4) polar heads and light-blue acyl chains and sitosterol is light-green.
Acknowledgements
This work was supported by the National Agency for Research (Grant ANR-14-CE19-0006-01 to E.M.B), the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation programme (grant agreement No 772103-BRIDGING to E. M.B), Fonds National de la Recherche Scientifique (NEAMEMB PDR T.1003.14, BRIDGING CDR J.0114.18 and RHAMEMB CDR J.0086.18 to L. L. and M D.). J.D.P. is funded by a PhD fellowship from the Belgian “Formation à la Recherche dans l’Industrie et l’Agriculture” (FRIA). Work in J.T, lab is supported by grant BB/M007200/1 from the U.K. Biotechnology and Biomedical Sciences Research Council (BBSRC).
Fluorescence microscopy analyses were performed at the plant pole of the Bordeaux Imaging Centre (http://www.bic.u-bordeaux.fr). The proteomic analyses were performed at the Functional Genomic Center of Bordeaux, (https://proteome.cgfb.u-bordeaux.fr). We thank Steffen Vanneste and Abel Rosado for providing the VAP27.1.RFP and SYT1.GFP binary vectors and Yvon Jaillais for providing the 1xPH(FAPP1) Arabidopsis transgenic lines. The plasmid pRBbar-OCS was kindly provided by Prof. Frederik Börnke (IGZ—Leibniz Institute of Vegetable and Ornamental Crops, Großbeeren, Germany). We thank Christophe Trehin and Patrice Morel for providing the AtMCT15_C2s construct and Alenka Copic for providing the yeast WT and A-tether strains. We thank Fabrice Cordelières for his help for the fluorescence image quantification and Paul Gouget, Yvon Jaillais, Andrea Paterlini, and Yrjo Helariutta for critical review of the article prior to submission.