Plasmodesmata in integrated cell signalling: insights from development and environmental signals and stresses

Journal of Experimental Botany, Nov 2014

To survive as sedentary organisms built of immobile cells, plants require an effective intercellular communication system, both locally between neighbouring cells within each tissue and systemically across distantly located organs. Such a system enables cells to coordinate their intracellular activities and produce concerted responses to internal and external stimuli. Plasmodesmata, membrane-lined intercellular channels, are essential for direct cell-to-cell communication involving exchange of diffusible factors, including signalling and information molecules. Recent advances corroborate that plasmodesmata are not passive but rather highly dynamic channels, in that their density in the cell walls and gating activities are tightly linked to developmental and physiological processes. Moreover, it is becoming clear that specific hormonal signalling pathways play crucial roles in relaying primary cellular signals to plasmodesmata. In this review, we examine a number of studies in which plasmodesmal structure, occurrence, and/or permeability responses are found to be altered upon given cellular or environmental signals, and discuss common themes illustrating how plasmodesmal regulation is integrated into specific cellular signalling pathways.

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Plasmodesmata in integrated cell signalling: insights from development and environmental signals and stresses

Journal of Experimental Botany Plasmodesmata in integrated cell signalling: insights from development and environmental signals and stresses Ross Sager 0 Jung-Youn Lee 0 0 Department of Plant and Soil Sciences, Delaware Biotechnology Institute, University of Delaware , Newark, DE 19711 , USA To survive as sedentary organisms built of immobile cells, plants require an effective intercellular communication system, both locally between neighbouring cells within each tissue and systemically across distantly located organs. Such a system enables cells to coordinate their intracellular activities and produce concerted responses to internal and external stimuli. Plasmodesmata, membrane-lined intercellular channels, are essential for direct cell-to-cell communication involving exchange of diffusible factors, including signalling and information molecules. Recent advances corroborate that plasmodesmata are not passive but rather highly dynamic channels, in that their density in the cell walls and gating activities are tightly linked to developmental and physiological processes. Moreover, it is becoming clear that specific hormonal signalling pathways play crucial roles in relaying primary cellular signals to plasmodesmata. In this review, we examine a number of studies in which plasmodesmal structure, occurrence, and/or permeability responses are found to be altered upon given cellular or environmental signals, and discuss common themes illustrating how plasmodesmal regulation is integrated into specific cellular signalling pathways. Plasmodesmata; cell-to-cell communication; development; environmental stresses; biotic and abiotic stress; hormones; cell signalling; callose Introduction Throughout their life cycle, plants process an array of internal and external signals and adapt to environmental conditions and challenges. These range from abiotic factors such as daily and seasonal fluctuations in light intensity/duration and temperature, and availability of water, carbon, nitrogen, and minerals, to biotic factors such as microbial infection and insect herbivory. Plant cells are equipped with both common and specific types of receptors necessary for perception of endogenous and environmental signals, as well as other molecular components for downstream signal transduction cascades. Thus, appropriate signal processing and metabolic generation of responses occur primarily at the individual cell level. Within a single cell, signal transduction is often initiated when chemical ligands are bound by cognate receptors, triggering an intracellular signalling cascade that eventually leads to changes in metabolic and/or nuclear activities. Although these cellular capacities meet the very basic need for individual cells to grow and survive, coordination of cellular responses between neighbours is necessary for the complex responses that affect the wellbeing and developmental progression of the plant as a whole. It is now generally accepted that membrane-lined intercellular bridges known as plasmodesmata are key parts of the cellular infrastructure that allows plant cells to communicate with virtually all of adjoining cells as well as distantly located cells, forming a symplastic network within the organism (Lee, 2014; Lucas and Lee, 2004; Roberts and Oparka, 2003) . Plasmodesma-mediated communication allows plant cells to dispatch mobile forms of intracellular information to adjacent cells within a tissue or organ. Moreover, using plasmodesmata in conjunction with a phloem-based transport system, plant cells can deploy supracellular, long-distance signalling. In essence, one might argue that plant cells are seldom fully independent from the physiological and biochemical influences of their neighbouring cells owing to the presence of plasmodesmata. However, although this state might be sufficient for photosynthetic organisms with simple body plans such as green algae, a monotonous cellular synchrony would not afford the complex lifestyles and body plans found in land plants (Lee, 2014). Indeed, plasmodesmata in vascular plants are dynamic channels, as demonstrated by the range in sizes and types of their molecular cargoes, and by their ability to undergo spatiotemporal gating and changes in frequency (Burch-Smith et  al., 2011b; Burch-Smith and Zambryski, 2011; Ehlers and Westerloh, 2013; Heinlein, 2002) . Plants can thus regulate cellular autonomy/non-autonomy by altering plasmodesmal connectivity, allowing them to orchestrate changes in growth and development at the organismal level and produce protective responses to environmental stresses. Plasmodesmata facilitate the movement of non-cell-autonomous signalling molecules, including transcription factors and other regulatory molecules such as long and small RNAs, and viral components. These topics are reviewed in many excellent articles and will not be covered in the current review; interested readers are referred to: Benitez-Alfonso et al., 2010 ; Ding, 2009; Lee et al., 2011a; Lee and Cui, 2009; Lucas, 2006 ; Niehl and Heinlein, 2011; and Wu and Gallagher, 2012 . In the current review, our primary focus will be on a less understood subject: how cellular signalling pathways are integrated into the regulation of plasmodesmata. The nature of the signals and signal transduction pathways that bring about changes in plasmodesmal permeability and the underlying mechanisms linking these signals to control of plasmodesmata have begun to be elucidated only recently. To stimulate new thoughts and discussion on the control of plasmodesmata-mediated intercellular communication, this review will highlight what is known about the intra- and extracellular stimuli, and challenges that lead to changes in plasmodesmal structure and connectivity, including cytosolic calcium, light, cold, and oxidative stresses, hormones, and pathogens. Furthermore, we will discuss various developmental processes that accompany changes in plasmodesmal frequency and permeability. Finally, we will present common and recurring mechanistic themes underlying the integration of cellular signalling pathways into the control of plasmodesmata. Regulation of plasmodesmal connectivity and permeability Plasmodesmata are dynamic channels A fundamental difference between animal and plant cells in terms of the biogenesis of intercellular communication channels is that the formation of plasmodesmata is coupled with cytokinesis in plant cells. The plasmodesmata formed during cell division are termed primary plasmodesmata (Ehlers and Kollmann, 2001) . These have a simple morphology, and each primary plasmodesma is composed of a cytoplasmic sleeve delimited by the plasma membrane (PM) externally, and an appressed endoplasmic reticulum (ER) cylinder internally (Fig. 1A). Formation of the primary plasmodesmata involves insertion or trapping of ER strands in the newly forming cell plate. Strands of cortical ER stretched across the phragmoplast become lined by plasma membrane, which leads to the formation of a cytoplasmic sleeve between daughter cells (Hepler, 1982; Hepler and Newcomb, 1967) . What determines the frequency (i.e. density) of primary plasmodesmata at the newly built cell wall interface is not understood, but it is generally assumed that the trapping of ER strands in the cell plate occurs fairly randomly, and new division walls all contain similar numbers of plasmodesmata (Faulkner et al., 2008). Through this direct coupling between sister cells, all plant cells newly produced by division embark on their cellular life in a non-cell autonomous manner. However, plasmodesmal frequency in vascular plants is not a fixed cellular parameter. As cells expand and differentiate, so-called secondary plasmodesmata can arise de novo in post-cytokinetic walls (Ehlers and Kollmann, 2001) . Both primary and secondary plasmodesmata are subject to various structural modifications as well as disintegration, depending on developmental or physiological cues (Fig. 1). Degeneration and de novo biogenesis of plasmodesmata are frequently associated with developmental progression or cell-type specification (reviewed in Burch-Smith et  al., 2011b) . For example, as guard cells mature and differentiate, plasmodesmata connecting them to adjacent pavement cells are lost (Fig. 1B). Similarly, plasmodesmata connecting pollen to nutritive tissue degenerate as the pollen cells mature. Interestingly, between the cells within each nurturing layer in anther, wide-open PM-lined channels similar to sieve plate pores (Esau and Thorsch, 1984) are formed as the internal structures of the plasmodesmata are removed (Mamun et al., 2005b; Steer, 1977; Willemse and van Went, 1984; Willmer and Sexton, 1979) (Fig.  1B, C). In addition, plasmodesmal frequency in the shoot apex can be transiently modified during floral development (Milyaeva, 2007) (Fig.  1D), and new plasmodesmata are formed between non-daughter cells during the establishment of graft unions or heterocystic parasites (Kollmann and Glockmann, 1991; Vaughn, 2003) . Furthermore, the morphology of both primary and secondary plasmodesmata is subject to significant modifications. As cells mature, plasmodesmata often undergo transformation from simple pores to complex multi-channel connections (Ehlers and Kollmann, 2001; Ehlers and Westerloh, 2013; Oparka et  al., 1999) (Fig.  1E). Branched, complex plasmodesmata often have a reduced size exclusion limit (SEL) compared with simple plasmodesmata, restricting the diffusion of larger molecules (Oparka et al., 1999) . The genetic components involved in degeneration, secondary biogenesis, and ultrastructural modification of plasmodesmata remain to be discovered. However, cell wallmodifying enzymes and their regulators probably play crucial roles in these processes (Burch-Smith et al., 2011b; Kollmann and Glockmann, 1991) . Whereas these types of plasmodesmal modifications reflect long-term or permanent cellular decisions, plant cells can also control plasmodesmal connectivity rapidly and transiently (Holdaway-Clarke et al., 2000; Tucker and Boss, 1996) . This level of control allows cells to respond to spatiotemporal changes in physiological and/or environmental conditions. Most well-understood reversible plasmodesmal regulation involves enzymatic biosynthesis and degradation of a polysaccharide called β-1,3-glucan (callose) at the plasmodesmata (Fig.  1F), which is described in the following section. Plasmodesmal gating is modulated by callose levels Plasmodesmata-mediated molecular flux between cells seems to be modulated through reversible deposition of callose (Zavaliev et al., 2011) (Fig. 1A and F). Callose deposits appear at the primary plasmodesmata in newly formed cell walls, and could be either a remnant of cell plate callose or resynthesized upon completion of plasmodesmal formation (Northcote et al., 1989) . In post-cytokinetic walls, callose is often found at the neck regions of plasmodesmata, reflecting a wound response associated with sample preparation for electron microscopy (Radford et al., 1998) . It is currently unclear whether callose deposited around the neck region strictly correlates with plasmodesmal closure as callose accumulating along the length of plasmodesmal channels is not uncommon (Vaten et al., 2011) . However, it is well established that callose levels at plasmodesmata can fluctuate, and the extent of callose accumulation surrounding the channels negatively correlates with plasmodesmal conductivity (Lee et al., 2011b; Rinne et  al., 2001) . The alterations in plasmodesma callose levels involves two counteracting enzymatic activities, i.e. β-1,3-glucan synthases and hydrolases. In fact, a recurring theme in plasmodesmal regulation is that various endogenous and extracellular signals bring about changes in transcriptional or enzymatic activities of β-1,3-glucan synthases or hydrolases (Dong et al., 2008; Rinne et al., 2011) . In Arabidopsis thaliana, twelve genes are predicted to encode the catalytic subunit of β-1,3-glucan synthase and are referred to as CALLOSE SYNTHASE (CALS) or GLUCAN SYNTHASE-LIKE (GSL) genes (Saxena and Brown, 2000; Verma and Hong, 2001) . Functional studies using genetic mutants have shown that various CALS family members are critical for different developmental and physiological processes (Barratt et  al., 2011; Chen et  al., 2009; Dong et al., 2005; Guseman et al., 2010; Hong et al., 2001; Jacobs et  al., 2003; Vaten et  al., 2011; Xie et  al., 2011) . CALS10, CALS3, and CALS7 affect molecular transport across plasmodesmata or sieve elements. CALS10 (also called GSL8) is required for proper cell plate establishment during cytokinesis as well as guard cell and root tissue patterning (Guseman et  al., 2010). Gain-of-function mutations in CALS3 have been tied to increased deposition of callose at plasmodesmata and decreased macromolecular trafficking between root cells, in addition to developmental defects in roots (Vaten et al., 2011) . CALS7 encodes a phloem-specific isoform that is required for normal deposition of callose in developing sieve elements and for phloem transport (Barratt et al., 2011; Xie et  al., 2011) . Highlighting the critical role of CALS in restricting plasmodesmal permeability, two novel CALS family members control basal and induced plasmodesmal closure (J.-Y. Lee, unpublished data). With regard to callose degradation, the Arabidopsis genome encodes approximately fifty β-1,3-GLUCANASE (BGL) genes, each containing a glycosyl hydrolase family 17 domain (Doxey et al., 2007; Zavaliev et al., 2011) . A few characterized BGL genes affect plasmodesmal callose levels and are involved in a range of developmental processes including cotton (Gossypium hirsutum) fibre elongation (Ruan et  al., 2004) , lateral root emergence (Benitez-Alfonso et  al., 2013) , and dormancy breakage (Rinne et al., 2011) . Changes in transcript levels of specific groups of tobacco (Nicotiana tabacum) BGLs (BGLs expressed in tobacco are grouped into five classes according to amino acid sequence identity of the mature proteins) positively correlate with viral spread both locally and systemically. For example, the silencing of genes for class  I  BGLs in tobacco leaves, which led to increased accumulation of callose at plasmodesmata, was enough to significantly delay the systemic movement of several viruses (Beffa et al., 1996; Iglesias and Meins, 2000) . Corroborating this effect, overexpressing genes encoding class  III or class I BGLs in transgenic potato (Solanum tuberosum) and tobacco, respectively, stimulated the spread of potato virus YNTN (Dobnik et al., 2013) and Tobacco mosaic virus (TMV) (Bucher et al., 2001). Given the large number of genes in this family, it would not be surprising to find more members that translate specific developmental and environmental cues into increased cell-to-cell permeability through plasmodesmal callose degradation. Experimental details described in the sections Forming lateral roots, Elongating cotton fibres, and Gibberellic acid plays a role in reestablishment of plasmodesmal connectivity during chilling-induced breakage of dormancy, below, highlight the theme that specific signalling pathways function in spatiotemporal induction of particular BGLs, which in turn reverse restrictions in plasmodesmal flux. Cytoskeletal components affect plasmodesmal targeting and flux: A callose-independent mechanism for plasmodesmal gating A number of cytoskeletal elements have been found to localize to or near plasmodesmata or to have a significant function in cell-to-cell transport. Actin microfilaments have been implicated in the transport of viral movement proteins (MPs) to plasmodesmata, although disruption of actin can have differing effects on plasmodesmata and viral MP, often dependent on the cell type. Protein dual-labelling by fluorescent tagging for co-expression in tobacco leaves showed that a 126-kDa component of the TMV viral replication complex colocalized with, and moved along, actin filaments (Liu et al., 2005) . Similarly, GFP-tagged cauliflower mosaic virus (CaMV) inclusion body protein P6 trafficked intracellularly along microfilaments (Harries et al., 2009a) . However, blocking actin polymerization allowed 10-kDa dextrans to move between tobacco mesophyll cells (Ding et al., 1996), whereas a similar treatment had no influence on basal trichome plasmodesmal function (Christensen et  al., 2009). By contrast, such actin disruption severely reduced plasmodesmal targeting of the TMV MP in tobacco epidermal cells (Kawakami et al., 2004; Wright et al., 2007) , and inhibited CaMV cell-tocell movement (Harries et al., 2009a) . Intriguingly, certain viruses may use targeted destabilization of actin as a means to facilitate their intercellular movement. For example, treatment of tobacco leaves with the drug phalloidin, which stabilizes actin filaments, prevented the cucumber mosaic virus (CMV) MP from increasing the plasmodesmal SEL (Su et  al., 2010) . Further analysis demonstrated that CMV MP binds to actin filaments in vitro and can in fact sever them (Su et al., 2010) . This led the authors to speculate that plasmodesmata-targeted CMV MP increases the SEL by breaking down actin cytoskeletal scaffolding in the plasmodesmal channel. Endogenous factors using such a mechanism to manipulate the plasmodesmal SEL are not yet known. Other viral proteins seem to localize to F-actin and use it as a scaffold for viral complexes. In infected cells, the TGB1 protein from Potato virus X (PVX) forms a structure called the X-body that collects F-actin in a mesh around its outer layer (Tilsner et al., 2012) . Though the function of the actin scaffolding is not yet known, PVX requires TGB1 and intact actin microfilaments to move cell-to-cell (Harries et al., 2009b) . Immunogold labelling has revealed filamentous actin at the plasmodesmal neck region (Baluska et al., 2004; White et al., 1994) , and myosin proteins, such as the unconventional plant myosin VIII, at the plasma membrane and in clusters near plasmodesmata (Baluska et  al., 2004; Radford and White, 1998; Reichelt et al., 1999) . Myosin VIII seems to be involved in targeting viral cargoes to plasmodesmata. Overexpression of a truncated, tail domain-only form of myosin VIII acts as a dominant-negative mutant, blocking cargo recognition by normal myosin VIII; overexpression of this domain in tobacco impairs targeting of the viral protein Hsp70 to plasmodesmata (Avisar et  al., 2008) . Recently, a study showed that cell-to-cell movement of 1- to 3-kDa dextrans was increased in staminal hairs of Tradescantia virginiana upon inhibition of myosin VIII function by treatment with antimyosin antibodies or the drug 2,3-butanedione monoxime, which binds myosin and slows its ATPase activity. On the contrary, permanent binding of myosin to actin induced by the drug N-ethylmalemide decreased cell-to-cell movement (Radford and White, 2011) . As the authors note, the application of these drugs may cause side-effects in plant cells that could alter intercellular transport, so further research into their effects on plasmodesmata is warranted. Furthermore, in this particular study, inhibition of actin polymerization with latrunculin B did not disrupt intercellular trafficking, in contrast to previous reports (Ding et al., 1996; Wright et al., 2007) , indicating there may be tissue- or species-specific differences in how actin affects plasmodesmal function. Other results support a possible role for myosins in plasmodesmal transport. In one study using grapevine fanleaf virus, which forms MP-derived tubules that stretch through plasmodesmata to allow the virus to move (Amari et  al., 2011) , blocking the function of myosin XI by overexpressing its tail domain alone suppressed the formation of viral tubules, and thereby also inhibited the cell-to-cell movement of the virus. It has also been shown that certain viruses can selectively use particular myosin proteins; silencing myosin XI-2 in tobacco repressed the intercellular spread of turnip mosaic virus by a factor of ten, whereas silencing myosin VIII-1, VIII-2, or XI-F did not affect its infection (Agbeci et al., 2013) . Similar results were obtained for TMV (Harries et  al., 2009b) . Finally, certain treatments, like calcium, may result in modulation of plasmodesmal conductivity by affecting the function of myosin proteins (see Calcium fluxes rapidly close plasmodesmata, below). The role of the microtubule-based cytoskeleton in plasmodesmal targeting remains more contentious (Niehl et al., 2013) . Certain viruses seem to use microtubules for targeting to plasmodesmata, whereas others do not. For instance, TGB1 of potato mop top virus requires association with microtubules with its N-terminal domain to localize to plasmodesmata (Shemyakina et  al., 2011; Wright et  al., 2010) . Treatment of tobacco with the microtubule-disassembling drug colchicine inhibited targeting of TGB1–GFP to plasmodesmata, whereas blocking microtubule polymerization with colchicine or oryzalin had no effect on the targeting of TMV MP to plasmodesmata in tobacco epidermal cells (Wright et al., 2007). However, other studies have suggested that whereas plasmodesmal targeting of TMV MP–GFP is unaffected, interaction of the MP with microtubules is required for effective cell-to-cell spread of viral RNA (Boyko et al., 2000; Boyko et al., 2007) . When tobacco plants infected with TMV were shifted to a high temperature, a stronger association of TMV MP–GFP with microtubules correlated highly with greater spread of the viral RNA. Intact microtubules may also be required for proper movement of endogenous non-cell autonomous proteins, as shown by trafficking of GFP-tagged SHORTROOT along microtubules into endodermal cells of Arabidopsis roots (Wu and Gallagher, 2013). Plasmodesmata undergo degeneration and structural remodelling during organogenesis, cell growth, and development Isolating mature guard cells from their neighbouring epidermal cells Guard cells require cellular autonomy to regulate stomatal aperture by modulation of their turgor pressure. Several lines of evidence indicate that this autonomy is achieved by severing their plasmodesmal connections to surrounding epidermal cells. Ultrastructural studies using transmission electron microscopy on Vicia faba, Allium cepa, Beta vulgaris, and tobacco guard cells revealed that the intact plasmodesmata between developing guard cells and their neighbouring cells disintegrate by the time of guard cell maturity (Wille and Lucas, 1984; Willmer and Sexton, 1979) . The deposition of a thick layer of cell wall material on the guard cell side of the shared wall results in severed (or halved) plasmodesmata in the adjacent epidermal cell wall (Wille and Lucas, 1984; Willmer and Sexton, 1979) . Consistent with these observed ultrastructural changes to guard cell plasmodesmata, numerous studies utilizing microinjection have shown that mature guard cells are symplasmically isolated from surrounding epidermal cells. Before maturation, fluorescent dyes can diffuse freely between immature guard cell pairs and subsidiary cells. For instance, when the membrane-impermeable fluorescent dye lucifer yellow carbohydrazide (LYCH) was injected into immature guard cells and subsidiary cells of A. cepa and Commelina communis, the probe was mobile between cells (Palevitz and Hepler, 1985) . However, when the dye was injected into mature subsidiary cells, the fluorescent signal spread into neighbouring epidermal cells but was excluded from mature guard cells. Furthermore, when one of a mature guard cell pair of A. cepa and Commelina communis was injected, the fluorescent signal was retained within that single cell, indicating that in these species each mature guard cell is symplasmically isolated not only from surrounding epidermal cells but also from its sister (Palevitz and Hepler, 1985) . By contrast, injecting one of a guard cell pair in maize (Zea mays) with the dye 2’,7’-bis-(2carboxyethyl)-5(and 6)carboxyflurescein revealed cytoplasmic sharing only between mature guard cells (Mumm et al., 2011) . This indicates that plasmodesmal connectivity within the stomatal complex itself may differ somewhat between species. Further supporting the idea that immature guard cell pairs remain symplasmically connected with surrounding epidermal cells, gene silencing signals penetrate into guard cells before their maturation (Voinnet et al., 1998) . Likewise, free GFP and fluorescently tagged viral MP were confined when transiently expressed in the mature guard cells of various dicots (tobacco, potato, Arabidopsis, and V.  faba) through biolistic bombardment (Hofius et al., 2004; Isono et al., 1997; Lee et  al., 2005; Myouga et  al., 2008) , which indicates that there is neither diffusion nor active transport out of mature half-stoma. Generating pollen, megagametophytes, and zygotes A developing anther contains multiple cell layers, including the tapetum which surrounds sporogenous tissue and provides nutrients to the developing pollen mother cells (Fig.  2A). Electron microscopy together with microinjection experiments show that the development of reproductive tissues accompanies major alterations in plasmodesmal connectivity. In early stages of pollen development, plasmodesmal connections exist at the cellular junctions between tapetum–tapetum and tapetum–pollen mother cell layers (Radice et al., 2008; Steer, 1977) . The presence of plasmodesmata in this tissue is thought to be crucial for the feeding of the developing pollen mother cells with nutrients and metabolites by the tapetal cells. Plasmodesmata were also observed between pre-meiotic pollen mother cells; however, as pollen mother cells progress into meiotic cell division, tapetal cells secrete callose that accumulates within the newly forming cell walls of pollen tetrads (Mamun et  al., 2005a) . The thickening of callose walls coincides with the disappearance of plasmodesmata at the end of this stage, which leads to symplasmic isolation of pollen grains (Mamun et  al., 2005b; Steer, 1977) (Fig.  2A). During meiosis, the tapetum layer itself also becomes symplasmically isolated from the surrounding tissue. This means that soluble nutrients produced in the inner middle layers must be transported apoplastically when they enter into and exit from the tapetal layers before reaching the developing pollens (Clement and Audran, 1995; Roschzttardtz et  al., 2011) . It is speculated that the symplasmic isolation of the tapetum during pollen development is necessary to protect neighbouring cell layers from potentially detrimental chemicals produced during programmed cell death of the tapetum. The cell death of the tapetum is thought to provide a mechanism to salvage the energy that should go into supporting the developing pollen cells (Wu and Cheun, 2000). Consistent with this notion, it was recently shown that plants contain underdeveloped or aborted pollens when tapetal cells are prevented from undergoing programmed cell death in rice (Niu et al., 2013) . Notably, the plasmodesmal connections remaining among tapetum cells seem to go through structural modification during pollen development. Sometimes much larger connections called cytomictic channels form between tapetum cells following symplasmic disconnection from the middle layer (Clement and Audran, 1995; Mursalimov et al., 2010; Perdue et  al., 1992; Vinckier and Smets, 2005) (see Fig.  1C & 2A). These large channels (or holes) are thought to facilitate the movement of organelles and chromatin, producing syncytia. Furthermore, whereas fully developed pollen grains are enveloped completely in callose, and thus lack plasmodesmal connections with each other (Mamun et  al., 2005b; Steer, 1977) , cytomictic channels can form between pollen mother cells (Mamun et  al., 2005a; Wang et  al., 2004) . The function of these channels is unknown, but it is conceivable that they facilitate exchange of essential nutrients or help coordinate the timing of meiotic division (Wang et  al., 2004). It remains to be determined whether cytomictic channels form de novo, or through the modification/fusion of existing plasmodesmata through the help of pit field-localized cellulase (Henslop-Harrison, 1966; Wang et al., 1998) . Megagametophytes, the female reproductive cells, also seem to require a symplastic system to fine-tune the timing of nutrient distribution during development (Willemse and van Went, 1984) . Ultrastructural analysis of the chalaza, the tissue connecting the ovule to the placenta, has shown that plasmodesmal frequency is higher in the chalazal cells than in neighbouring cell types (Russell, 1979; Thijssen, 2003) . This feature may have to do with a potential role of the chalazal tissue in regulating the symplastic flow of nutrients from the phloem to the egg sacs. In fact, during the tetrad stage of megagametophyte development, the ovule primordium excluding the tetrad is symplastically connected to the vasculature (Werner et al., 2011) (Fig. 2B.1). However, by stage 12 of anthesis during ovule maturation, the plasmodesmata in chalazal cells apparently become occluded (Thijssen, 2003) , coinciding with a lack of symplastic dye movement from the vasculature (Werner et  al., 2011) (Fig.  2B.2). Furthermore, no plasmodesmal connections between the cells of the nucellus (egg sac tissue) and integuments (outer protective tissue) are observed, other than within the chalazal zone, in late stages of ovule development (Thijssen, 2003) . Interestingly, symplastic dye movement into ovules from the vasculature resumes by late stage 13, but the tracer can move only into the outer integument (Werner et al., 2011) (Fig. 2B.3). Studies utilizing electron microscopy, symplastic dye tracers, and outer/inner integument-specific expression of GFP have all revealed that plasmodesmal connections are retained within the nucellus and integument tissues themselves (Han et al., 2000; Thijssen, 2003; Wang and Fisher, 1994; Werner et  al., 2011) . For example, examination of symplasmic domains within late-stage ovules using transcriptional GFP reporters driven by the promoters of AtGL2 and AtTT12 showed that the outer and inner integuments are isolated from each other (Fig. 2B.4). However, although AtGL2:GFP drives expression only within the outermost cell layer of the outer integument, GFP could move into the inner cell layer of the outer integument. Similarly, AtTT12:GFP is active only within the innermost layer of the inner integument, and GFP could diffuse outward through all inner integument layers. Collectively, these data suggest that ovules develop tissue-specific symplasmic and symplastic barriers, and isolate megagametophytes following a period of initial development during which photosynthates are directed symplastically through the chalaza to the growing megagametophytes. Similar to the pollen development, the symplasmic isolation of megagametophytes may be required to prevent cytoplasmic leakage of signalling molecules produced by neighbouring cells that could interfere with the gameto- promoter, which is active in the hypocotyl epidermis, can phytic differentiation. diffuse throughout the whole embryo (Stadler et  al., 2005) A similar increase in symplasmic isolation also occurs dur- (Fig.  2C.2). Furthermore, the small transcription factor ing embryo development. Before fertilization, the egg cell TARGET OF MONOPTEROS 7 (TMO7) is trafficked interof Torenia fournieri is cytoplasmically connected to the sur- cellularly from embryonic cells to an adjacent suspensor cell rounding tissue in the embryo sac, as evidenced by movement called the hypophysis, or primary root meristem founder cell of a fluorescent reporter conjugated to a 10-kDa dextran of the developing seedling (Schlereth et al., 2010) . Expression from microinjected central cells into the egg cell and syner- of TMO7–GFP was observed in globular and heart stage gids (Han et al., 2000) (see Fig. 2B.2, inset). Starting around embryos in both the hypophysis and adjacent embryonic cells 24 hours post-fertilization, however, movement of even free (Fig. 2C.2), whereas a much larger form of TMO7 fused to dye reporter into the newly formed zygote is prevented. Early triple GFP could not traffic from the embryo into the hypoultrastructural imaging of soybean and Arabidopsis showed physis. Another study showed that early in embryogenesis that embryonic plasmodesmata are disconnected from the and up to the heart stage, fluorescent tracers, e.g. 0.5-kDa surrounding endosperm except within the suspensor, a spe- 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS) and 10-kDa cialized tissue that draws nutrients from maternal tissues into F-dextrans, spread cell-to-cell throughout the whole embryo the embryo (Dute et al., 1989; Mansfield and Briarty, 1991) . (Kim et al., 2002). By the torpedo stage, however, movement In Arabidopsis, suspensor-specific expression of diffus- of 10-kDa dextrans was restricted to only a few cells. Use of ible GFP under the SUC3 promoter revealed movement of multimeric GFPs as fluorescent reporters expressed from the the protein into the globular stage embryo (Fig.  2C.1), yet embryonic SAM produced similar results (Kim et al., 2005a; such cell-to-cell movement was prevented during the heart Kim et al., 2005b) . In early heart stage embryos, both the sinstage (Stadler et al., 2005) (Fig. 2C.2). This result is consist- gle and a tandem double GFP could move throughout the ent with ultrastructural study of Crassulaceae ovules at dif- hypocotyl into embryonic cotyledons and root. However, by ferent embryonic stages, which revealed that the complexity the late heart stage, double GFP was blocked from moving of suspensor cell plasmodesmata increases as the embryo into embryonic cotyledons, but not root. A  triple GFP was matures; transitioning from simple, unblocked plasmodes- always restricted to the hypocotyl. GFP driven by the SUC3 mata to complex, branched plasmodesmata that are plugged promoter, which is active in the embryonic root during the with an electron-dense substance (Kozieradzka-Kiszkurno torpedo stage, spread into a wider symplasmic zone of the and Plachno, 2012). The authors noted the similarity of embryo through the mid-torpedo stage (Stadler et al., 2005) this material to the callose that was accumulated in the plas- (Fig. 2C.3). However, in late torpedo stage, GFP signal was modesmata of birch (Betula pubescens) shoot apical meris- confined to where the promoter was active, demonstrating tem (SAM) during dormancy (Rinne and van der Schoot, the overall decrease in intercellular permeability as embryos 1998) . However, intriguingly, shadows within the plugs have mature (Fig. 2C.4). These studies underscore that spatiotemcontinuity with the ER, suggesting that the appressed ER of poral regulation of plasmodesmata is probably critical for suspensor plasmodesmata may have been structurally modi- coordinating the development of distinct tissues and organs fied to increase the filtration of nutrients or disable transport from the very early stage of embryo formation. altogether (Kozieradzka-Kiszkurno and Bohdanowicz, 2010; Kozieradzka-Kiszkurno and Plachno, 2012). Preparing organs for abscission As zygotes develop, internal signalling factors need to be contained and directed to the developing areas, which Organ separation during abscission results from dissolution is apparently aided by changes in plasmodesmal connec- of the cell walls between the organ and the main plant body tivity and symplasmic zoning. During the heart stage of at specialized areas called abscission zones (Estornell et  al., embryo development, GFP expressed under the AtGL2 2013; Sexton and Roberts, 1982) . Within abscission zones, layers, though still blocked from the outer integument. (C) Changes in plasmodesmal connectivity during embryo development. 1. Globular stage: The suspensor (Sus) and embryo (Em) are cytoplasmically connected, as indicated by the diffusion of GFP from the suspensor into the embryo (green gradient) reported by AtSUC3:GFP. Dashed bracket, expression domain of AtSUC3:GFP; broken brown lines, open plasmodesmata within suspensor. 2. Heart stage: The suspensor becomes cytoplasmically isolated as its plasmodesmata increase in structural complexity and are occluded by a calloselike substance (filled red boxes), preventing GFP diffusion from suspensor to embryo (green). However, the heart stage embryo itself remains a symplastic domain, as indicated by the diffusion of GFP throughout the embryo from the hypocotyl-located expression of AtGL2:GFP (yellow). The plasmodesmata between the hypophysis and the embryo also remain open, as indicated by the movement of GFP signal in AtTMO7:TMO7–GFP from the embryo into the hypophysis (orange). Dashed bracket, expression domain of AtSUC3:GFP. 3. Mid-Torpedo stage: Plasmodesmata remain open in this embryo stage, as indicated by the diffusion of GFP from the root tip (dark green) throughout the embryo (green), driven by the AtSUC3 promoter. 4. Late Torpedo stage: The symplastic connectivity of the torpedo stage embryo becomes restricted as the embryo matures, and GFP does not diffuse out of its expression domain in the root tip (dark green). At this stage, AtSUC3 is active in other tissues of the embryo, but GFP remains restricted to these cells as well (green line and dashes). (D) Plasmodesmata undergo structural remodelling and form highly complex channels during abscission. In the abscission zone, multivesicular (MVB) and paramural bodies (PB) are frequently found nearby or associated with complex plasmodesmata. This occurs almost exclusively in the proximal tissue of the zone. It is hypothesized that these bodies contain cell wall (CW)-remodelling enzymes, such as cell wall hydrolases (CWH) and peroxidases (Per), to loosen and degrade the middle lamella (ML) between the main plant body and the abscised organ, allowing it to separate. Post-separation, these vesicles may also play a role in bringing cell wall material to repair the outer layer and prevent cytoplasmic leakage through plasmodesmata. PM, plasma membrane; AER, appressed endoplasmic reticulum. plasmodesmata undergo extensive structural remodelling, with a very high occurrence of branched plasmodesmata (Bar-Dror et al., 2011; Jensen and Valdovinos, 1967; Osborne and Sargent, 1976; Webster, 1973) . Complex plasmodesmata, large cavities, and membrane vesicularization are observed within the middle lamella region of the abscission zone (BarDror et al., 2011; Osborne and Sargent, 1976) , indicating that cell wall separation may preferentially initiate there (Fig. 2D). Diaminobenzidine staining also revealed high levels of hydrogen peroxide, which is known to weaken cell wall connections, within plasmodesmata, cell walls, and Golgi vesicles in the abscission zones of tobacco flower pedicels (Henry, 1979) . Furthermore, paramural bodies—membrane vesicles that form between the cell wall and PM—accumulate at complex plasmodesmata during abscission, with the majority forming in the proximal side of the zone (Bar-Dror et  al., 2011) (Fig. 2D). These vesicular bodies have been hypothesized to contain cell wall-modifying enzymes, such as hydrolases and peroxidases, brought there to enhance the breakdown of the cell wall between the strands of complex plasmodesmata, enabling separation of the distal tissue (Osborne and Sargent, 1976) . Alternatively, they may contain cell wall material, deposited to plug the plasmodesmata of the proximal tissue once the distal tissue separates (An et  al., 2006) (Fig.  2D). It is noteworthy that, in addition to complex branching of plasmodesmata, sieve plate pores in the vasculature of P. vulgaris have significantly reduced levels of callose plugs during abscission (Scott et  al., 1967) . This parallel modification in sieve plate pores may facilitate nutrient mobilization from the distal to the proximal tissue prior to abscission of the former. All the abscission studies discussed above involved ethylene treatment. Considering this experimental condition, it is tempting to speculate that there might be a direct link between the changes in plasmodesmal structure and the ethylene signalling pathways. This possibility may be tested by assessing the effect of ethylene treatment on plasmodesmal permeability as has been shown for the salicylic acid-induced plasmodesmal closure (Wang et al., 2013) . Producing larger pores in sieve tubes for phloem mass flow The formation of sieve plate pores is a fascinating example of a developmental process in which plasmodesmata at specific cellular junctions are structurally modified to provide the cells with enhanced intercellular transport capacity (see Fig. 1C). Sieve elements are elongated, specialized conducting cells of the phloem that are alive at maturity. Conjoined end to end via the sieve plate, these cells form functional sieve tubes, and mass flow of phloem contents occurs through enlarged pores within the sieve plate (Knoblauch and van Bel, 1998; van Bel et al., 2002) . During sieve element maturation, the sieve pores form through major structural modifications of the primary plasmodesmata that once connected daughter sieve elements. Initially, hyperaccumulation of callose creates a thick cell wall layer surrounding sieve plate plasmodesmata. This is followed by disappearance of the appressed ER and degradation of the callose, eventually leaving much wider pores that are lined only with PM (Esau and Thorsch, 1984) . The functional importance of the transient callose accumulation in sieve pore formation was recently supported by mutant analysis of the phloem-specific callose synthase Arabidopsis CALS7 (Xie et  al., 2011) . Loss-of-function mutation in CALS7 resulted in poorly developed sieve plates that had greatly decreased callose accumulation and fewer sieve pores with narrower openings. Moreover, using 14CO2 labelling, the rate of phloem transport of photosynthate from source to sink tissues was found to be decreased in cals7 (Barratt et al., 2011) . Shoot apical meristems undergoing developmental phase change Symplasmically regulated cellular zoning of SAMs ensures that signals involved in organ differentiation remain in the intended groups of cells. In the vegetative SAM of birch, iontophoretic microinjection of LYCH dye into a single cell of the central zone tunica led to spread of the dye throughout only central zone tunica cells (Fig. 3A.2). Similar results were obtained when a tunica cell within the peripheral zone was microinjected (Rinne and van der Schoot, 1998) . The SAM/phloem interface is also symplastically regulated (Gisel et al., 1999) . When HPTS dye was loaded into cut petioles of Arabidopsis leaves, cell-to-cell movement through the vasculature into younger SAM tissue was restricted until the plants were about 4 weeks old (Fig. 3A.1). At this point, the dye diffused into the peripheral zone and tunica of the central zone except for a region of cells called the inner central zone, which may be the organization centre (Wang and Fiers, 2010) , a site of high levels of signalling and growth regulation (Fig. 3A.2). The control of cell-to-cell connectivity is especially dynamic in the SAM, as seen during developmental processes like floral transition (Fig.  3A) and dormancy cycling (Fig.  3B, detailed in Gibberellic acid plays a role in reestablishment of plasmodesmal connectivity during chilling-induced breakage of dormancy below). SAMs undergo several massive changes during reproductive transition, with not only gene expression profiles, but also zoning patterns being altered (Chandler, 2012). Arabidopsis requires 1–4 consecutive long days to commit to flowering, during which time HPTS dye could unload into the SAM from the vasculature. However, from the time immediately before floral commitment, no dye moved into the SAM (Gisel et  al., 1999; Gisel et  al., 2002) (Fig.  3A.3). During and immediately after the floral transition, there was a period of symplastic isolation, but dye movement in the SAM eventually returned to normal (Gisel et  al., 1999) (Fig.  3A.4). Surprisingly, however, the time it took Arabidopsis plants to recover symplastic connectivity between the SAM and stem vasculature depended upon their initial growth conditions. When grown in long days from the start, dye could move from leaves to the SAM almost immediately after floral transition. By contrast, when plants were initially grown in short days, this symplastic recovery took 2–3 weeks (Gisel et  al., 2002) . This prompts the intriguing question of how symplastic transport can be altered by initial photoperiod and light levels. As described below, the addition of secondary plasmodesmata to cell-cell junctions could be one aspect of this phenomenon. The Sinapis alba SAM central zone changes shape and size upon long day-induced floral transition. By the second long day, the central zone symplasmic field became more circular, and the area of dye spread more than tripled, though the sizes of SAM cells remained unchanged (Ormenese et al., 2002) (Fig. 3A.3). These changes in zoning pattern may be caused by alterations to plasmodesmal frequency (see Fig. 1D), modulating the connectivity between cell layers in the SAM (Fig. 3A.3, inset). A single long-day treatment sufficient to induce floral transition in Sinapis also induced transient changes in plasmodesmal formation in the SAM (Ormenese et al., 2000) . Serial thin sections of each Sinapis meristematic layer revealed that starting about 28 hours after floral induction, the plasmodesmal frequency had increased in all meristematic layers compared with uninduced plants, peaking at about 4-fold higher. However, at 48 hours post-long day induction, the frequencies of plasmodesmata in all SAM layers had returned to normal. The plasmodesmata in all observed Sinapis SAM tissues were almost always simple, and their ultrastructure unmodified, in both control and long day-induced plants (Ormenese et al., 2000) . Consistent with these observations, electron microscopic analyses of Rudbeckia and Perilla SAMs that were induced to the reproductive phase by changes in photoperiod also revealed that plasmodesmal frequency between SAM cells in all layers increased significantly at first (Milyaeva, 2007) . Intriguingly however, after this initial increase, only the cells of the SAM central zone retained their higher plasmodesmal frequency, whereas the number of plasmodesmata in the medullar zone dropped to below that of non-induced control plants. In Iris inflorescence meristem, the L2 layer has the highest plasmodesmal frequencies in comparison to the L1 and corpus layers (Bergmans et al., 1997). However, during the development of the inflorescence meristem into a floral meristem plasmodesmal densities of the L2 cells are significantly reduced in all interfaces, whereas the number of plasmodesmata between clonal cells within the L1 and those within the corpus remain essentially the same. As the L2 layer is positioned between the inner corpus and the outer L1 in the SAM, Bergmans et al. (1997) posit that the L2 layer may play a central role in dictating the symplasmic integration and thus the developmental fate of meristem in Iris. Collectively, the studies described above suggest that transient changes in plasmodesmal frequency or patterning in the shoot meristem probably reflect dynamic intercellular signalling, necessary for determining the shift in the developmental phase and to specify the zones of floral organ differentiation. Upon floral induction, symplastic transport from the vasculature is blocked, and plasmodesmal frequencies within the SAM initially increase, possibly to prevent movement of unwanted signals from the vasculature into the SAM, while allowing necessary floral differentiation signals to move from one SAM zone into another. However, plasmodesmal frequencies in these tissues eventually either return to pre-transition levels or decrease permanently, underscoring that cellular autonomy and non-autonomy are highly plastic parameters which well reflect developmental status of the plant. Forming lateral roots Lateral root primordia are initiated in the xylem pole pericycle, growing via a series of controlled cell divisions from a few cells into the new lateral organ, which pushes through outer cell layers of the primary root (Malamy and Benfey, 1997) . During development, the lateral root becomes symplastically isolated from the primary root phloem and, phloem-loaded symplastic tracer cannot move into the newly developing lateral root primordium (Fig. 4A). However, this symplastic isolation is temporary, lasting only until the new phloem elements within growing lateral root have differentiated, at which point phloem unloading resumes (Oparka et  al., 1995) (Fig.  4A). Transient callose deposition at plasmodesmata within the lateral root primordium plays a critical role in the formation and maintenance of this symplastic domain (Benitez-Alfonso et al., 2013) (Fig. 4A). Using GFP as a reporter, Benitez-Alfonso et al. (2013) showed that cells of lateral root primordia are symplastically connected to the pericycle at initial stages of development, but gradually become isolated. As detected by immunofluorescence, the timing of symplastic discontinuity correlates with an increase in callose deposition not only within lateral root primordia, but also in the underlying vasculature and overlying endodermal cells. Using a combination of transcriptome data mining with localization studies, the authors also identified plasmodesmal-localized β-1,3-glucanase 1 (PdBG1) as an important player in this process. A lack of PdBG1, and its closely related isoform PdBG2, resulted in higher callose accumulation in growing lateral root primordia, reduced GFP unloading from the phloem, and increased lateral root initiation and density. Elongating cotton fibres Cotton fibres, unique trichomes formed on the surface of seeds, are single cells that elongate from microns to centimetres in a matter of weeks (Lee et al., 2007; Schubert et al., 1973) . Transient inhibition of plasmodesmal connectivity during cotton fibre elongation correlates with the maintenance of the high turgor pressure necessary for driving the elongation process (Ruan et  al., 2001) . During the elongation period of 10–16  days after anthesis, movement of fluorescent reporter from the phloem into the fibre cells was blocked (Fig.  4B). Ultrastructural imaging revealed that the plasmodesmata connecting the fibre to the seed coat gradually change from simple to complex as elongation takes place. Moreover, using immunogold labelling, callose accumulation in plasmodesmata was shown to transiently increase at the fibre base during the elongation phase (Fig. 4B). The reversal in callose accumulation was correlated with the induction of fibre-specific β-1,3glucanase named GhGlu1 and plasmodesmal re-opening (Ruan et al., 2004) (Fig. 4B), similar to the role of PdBG1 during lateral root initiation (Benitez-Alfonso et al., 2013) . Interestingly, in the cotton mutant line fls, which shows a severe short-fibre phenotype, fluorescent symplastic dyes consistently moved into the fibres throughout fibre elongation, concomitant with an inability to close plasmodesmata during any stage of elongation (Ruan et al., 2005) . These studies point to a well-coordinated signalling pathway integrating spatiotemporal regulation of plasmodesmal conductivity at the seed coat/cotton fibre cell interface into developmental programme. Intra- and extracellular signals alter plasmodesmal permeability Salicylic acid is critical for plasmodesmal closure during defence Salicylic acid (SA) is the central hormone in innate immune responses to biotrophic and hemibiotrophic pathogens. SA is produced in chloroplasts and triggers nuclear localization of the SA-response coordinator NPR1 to up-regulate genes essential for protecting cells from infection (Mou et al., 2003) . Our recent studies have shown that SA, and SA signalling components including NPR1, play a critical role in plasmodesmal closure through PLASMODESMATALOCATED PROTEIN 5 (PDLP5) (Lee et al., 2011b; Wang et al., 2013) (Fig. 5, top right). PDLP5 is a type I transmembrane protein containing a cysteine-rich extracellular domain and a transmembrane domain with short C-terminal tail, and belongs to a family of eight members in Arabidopsis (Thomas et al., 2008) . PDLP5 was initially isolated from a biochemical extraction of plasmodesma-enriched cell walls prepared from young Arabidopsis seedlings (Lee et al., 2011b) . In this study, immunogold labelling showed that PDLP5 was localized at the central region of plasmodesmal channels. It was also shown that PDLP5 was expressed at a very low level in young plants under normal growth conditions, but was induced in senescing tissues and upon SA treatment or pathogenic bacterial infection. Loss of PDLP5 function in the pdlp5-1 mutant results in enhanced basal plasmodesmal permeability, whereas overexpression of PDLP5 severely restricts plasmodesmal permeability, underscoring its role in constraining plasmodesmata (Lee et al., 2011b) . Interestingly, although no enzymatic function has been predicted for PDLP5, plasmodesmal callose deposition is positively correlated with the level of PDLP5. Moreover, SA- or pathogen-induced plasmodesmal callose hyperaccumulation and subsequent plasmodesmal closure require PDLP5, suggesting its key role in controlling plasmodesmal callose deposition (Wang et  al., 2013) . Further highlighting the role of PDLP5 in mediating immune signalling through plasmodesmal regulation, the enhancement of plasmodesmal permeability in pdlp5-1 is directly correlated with susceptibility to pathogenic bacteria, whereas restriction of plasmodesmal permeability in PDLP5-overexpressing plants correlates with resistance against bacterial pathogens. Excitingly, plants overexpressing PDLP5 hyperaccumulate SA, indicating a positive feedback loop between SA-directed PDLP5 induction and PDLP5-regulated SA accumulation (Lee et  al., 2011b) . Notably, the SA downstream regulator NPR1 is critical not only for PDLP5 induction but also for PDLP5-based activity in plasmodesmal inhibition (Wang et al., 2013) , underscoring a tight link between SA signalling and plasmodesmal closure. Interestingly, SA has also been implicated in enhancing plasmodesmal complexity. Seedlings treated with SA for five days had approximately three times more plasmodesmal branching than did wild-type plants (Fitzgibbon et al., 2013). SA biosynthesis is up-regulated during the basal defence Given that SA functions also in promoting senescence, this response, and its accumulation is often inextricably linked to effect on plasmodesmal modification probably reflects over- reactive oxygen species (ROS) production and intracellular all changes in developmental programming. It is tempting redox changes. Both ROS production and SA biosynthesis to speculate, based on the known relationship between SA occur in chloroplasts (Serrano et al., 2013) , and as described level and changes in cellular redox states, that such SA effect in the following section, changes in the redox status of the on plasmodesmal complexity could possibly be explained plastids can affect plasmodesmal permeability and complexby altered redox status of the cell (Sager and Lee, 2012) . ity. Thus, perhaps SA- and ROS-based redox changes in the plastids and cytoplasm during defence and senescence may together contribute to the fine-tuning of the structure and permeability of plasmodesmata. Gibberellic acid plays a role in reestablishment of plasmodesmal connectivity during chilling-induced breakage of dormancy Dormancy is a period when plant growth slows to a halt; to survive cold winters, shoots may develop protective buds over meristematic zones, and plant cells induce a set of freezetolerance proteins. A  crucial step for enabling dormancy is the arrest of growth within shoot apical meristems (SAMs), accomplished in part by sealing the plasmodesmata within the entire SAM zone and the vasculature of the stem below it. Microinjection of LYCH into a single cell of the proliferating SAM of birch plants showed cell-to-cell movement of the dye (Rinne and van der Schoot, 1998) (see Fig. 3B). However, no dye diffusion was observed in the dormant SAM, concomitant with the high level of callose deposited within the plasmodesmata of all cells within the SAM at that stage (Rinne et  al., 2001) (Fig.  3B.1). However, a chilling treatment sufficient to break dormancy restored the dye-coupling patterns found in active SAMs, indicating that dormancy breakage involves re-establishing the symplastic connection of the SAM (Fig. 3B.2). Consistently the strong labelling of plasmodesmata by anti-callose antibodies within dormant birch SAM tissue was reversed after dormancy was released by chilling treatment. As dormancy breakage eliminated most of the accumulated callose at plasmodesmata, it was hypothesized that subsets of the family GH17 enzymes may be expressed and localize to plasmodesmata (Rinne et  al., 2001) . Immunoblotting using antibodies produced against a conserved sequence of GH17 family BGL proteins revealed that some members are present during all stages of dormancy, whereas others accumulate only during the release period. Immunogold labelling showed changes in BGL cellular localization depending on the cycle of dormancy (Rinne et  al., 2001) . Normally present in cell plates and young division walls of active SAM, BGLs in dormant SAM were found predominantly within the lumen of cortical ER-associated lipid bodies that became abundant during short day induction. Importantly, upon chilling to break dormancy, these lipid bodies labelled with BGL antibody were often associated with plasmodesmata, while plasmodesmal orifices were also occasionally labelled. These data suggest that ER-derived lipid bodies may retain BGLs during dormancy and act as vehicles for delivery to plasmodesmata as soon as dormancy is lifted. Using this mechanism, degradation of plasmodesmal callose, and thereby unsealing of plasmodesmata, may occur in time to promote cellto-cell movement of signalling molecules that support the growth and differentiation of SAM when it is released from dormancy. The phytohormone gibberellic acid (GA) has been tied to dormancy establishment and release (Saure, 1985; Zanewich and Rood, 1995) , while chilling of dormant hybrid aspen (Populus tremula × Populus tremuloides) buds promotes up-regulation of GA biosynthesis genes (Rinne et al., 2011) . The GA3 and GA4 analogues induced specific subsets of GH17 family BGL genes. Furthermore, transport of the symplastic tracer dye calcein into the shoot apex was restored after a chilling period sufficient to release dormancy (8 weeks) or after the addition of GA4, but not GA3, to dormant buds (Rinne et  al., 2011) . FLOWERING LOCUS T (FT) and CENTRORADIALIS-LIKE 1 (CENL1) are signalling peptides targeting regions in the SAM, and their expression is regulated by dormancy cycling (Rinne et  al., 2011) . FT transcripts were up-regulated in dormant aspen buds by 800-fold during chilling, but after bud burst were markedly reduced. By contrast, CENL1 transcripts were low in buds during chilling, but a shift to long day and higher temperature conditions greatly induced CENL1 preceding bud burst (Rinne et al., 2011) . Both GA analogues had little effect on FT, but strongly induced CENL1 in buds. Collectively, these data have led to a model explaining the role of symplasmic and symplastic reestablishment during dormancy breakage (Rinne et al., 2011) (Fig. 5, bottom right). According to this model, chilling up-regulates GA3 biosynthesis and FT expression together in the embryonic leaves of dormant buds. At this stage, FT cannot move from the leaves into the SAM because phloem sieve pores are blocked by callose. GA3 also induces the production of a specific subset of BGLs that localize to lipid bodies produced during dormancy. After a chilling period sufficient to break dormancy, the BGL-containing lipid bodies are mobilized to plasmodesmata and sieve plate pores, where they release the BGLs to degrade callose deposits. FT can now move through the vasculature into the SAM, where it acts to prime SAM cells for full dormancy breakage and bud burst. Increasing temperature then triggers GA4 production, which activates another subset of BGLs that fully degrade dormancy callose accumulated at the plasmodesmata between SAM cells. This initiates bud burst and stem elongation in the dormancy-released SAM. CENL1, induced by GA4, is known to be expressed in the rib meristem of Populus, but it is speculated that the degradation of callose in the SAM may allow CENL1 to move into nearby cells. It would be interesting to determine if CENL1 has a non-cell autonomous function outside the rib meristem tissue, similarly to its Arabidopsis orthologue TFL1 (Conti and Bradley, 2007), which affects dormancy release or bud burst. Also, it would be highly informative to test whether GA-mediated BGL induction is conserved across plant species, and if so, how prevalent this singling pathway is for spatiotemporal control of plasmodesmal permeability. Changes in redox status modulate plasmodesmal permeability Reactive oxygen species (ROS), such as hydrogen peroxide and superoxide, can be utilized as second messengers when produced in controlled bursts during various physiological, developmental, and stress responses in plants (Nanda et al., 2010; Tian et  al., 2013) . Several studies have shown close relationships between cellular redox status and regulation of plasmodesmal trafficking (Fig.  5, top left). ISE1 and ISE2, encoding a mitochondrion-localized DEAD-box RNA helicase and a chloroplast-localized DEVH-box RNA helicase, respectively (Burch-Smith et  al., 2011a; Stonebloom et  al., 2009) , were identified from a mutant screen of Arabidopsis embryos exhibiting increased SEL (Kim et  al., 2002). Mutations in these genes rendered the embryonic plasmodesmata more open to the movement of large dextrans. Moreover, whereas wild-type embryos mostly had simple plasmodesmata at the torpedo stage, ise1 and ise2 contained more branched and twinned plasmodesmata (Burch-Smith and Zambryski, 2010). Similarly, silencing ISE1 or ISE2 in adult tobacco leaves coincided with an increase in frequency or proportion of branched plasmodesmata, respectively. Further, silencing ISE1 heightened ROS production in mitochondria and increased intercellular permeability to TMV MP–GFP (Stonebloom et  al., 2009) . Strongly oxidizing the mitochondria or reducing the chloroplasts also increased cell-to-cell movement (Stonebloom et  al., 2012) . Enzymes producing ROS are known to function in cell wall remodelling processes (Liu et  al., 2014) , and specific targeting of such enzymes to plasmodesmata may lead to the enhanced plasmodesmal structural complexity seen in ise1 and ise2 (Burch-Smith and Zambryski, 2010). Detection of a class of peroxidases that produce hydroxyl radicals at or near plasmodesmata in the cambial zone of tomato (Solanum lycopersicum) stems (Ehlers and van Bel, 2010) is consistent with the notion that these enzymes may contribute to the changes in local redox state, which in turn affects plasmodesmal permeability and remodelling. The importance of redox status for control of intercellular trafficking was further supported by a recent study showing that mutating the chloroplast-localized antioxidant thioredoxin m3 (TRX-m3) severely hindered phloem unloading of GFP in the roots of pSUC2:GFP compared with wildtype seedlings (Benitez-Alfonso et  al., 2009) . The mutant, called gat1, died very early in development, similar to ise1 and ise2, but unlike in those mutants embryonic diffusion of carboxyfluorescein was unaffected in gat1, suggesting that defects in symplastic connectivity manifest later in development. However, diaminobenzidine and aniline blue staining detected high levels of ROS and callose, respectively, in the root apices of gat1 and similar gating mutants (BenitezAlfonso and Jackson, 2009) . When ectopically overexpressed in leaf tissues, GAT1/TRX-m3 could increase the movement of GFP that was transiently expressed through biolistic bombardment (Benitez-Alfonso et  al., 2009) . Overexpression of GAT1/TRX-m3 in chloroplasts led to a more reduced plastid environment, whereas the knockout gat1 mutant contained oxidized plastids, along with an increase and decrease in plasmodesmal trafficking. These results suggest the existence of a system by which plant cells detect intracellular redox states and transmit a signal to modify plasmodesmal structure and permeability (Fig. 5, top left). Interestingly, another thioredoxin family member, TRXh9, was shown to move cell-to-cell through plasmodesmata (Meng et al., 2010) . TRX-h9–GFP localized to the plasma membrane, unlike many of its family members, though it has no known transmembrane domains. However, analysis of the TRX-h9 protein sequence revealed N-terminal domain Gly and Cys residues in a pattern frequently associated with palmitoylation, the post-translational addition of fatty acids that can enhance the membrane association of an otherwise soluble protein (Meng et al., 2010) . This putative palmitoylation site was critical for the intercellular trafficking of TRX-h9, as when TRX-h9–GFP was expressed under the root endodermis-specific SCARECROW promoter, its movement out of the endodermal layer was halted by mutations in the palmitoylation motif. It is not yet known whether the intercellular movement of TRX-h9 has any effect on redox-based intercellular signalling and/ or plasmodesmal modulation. It is tantalizing, though, to speculate that plant cells may utilize intercellular antioxidant proteins, like Trx-h9, to help prevent oxidative stress from spreading into neighbouring cells through plasmodesmata (Fig. 5, top left). Calcium fluxes rapidly close plasmodesmata Cytoplasmic calcium is involved in numerous signalling pathways as a secondary messenger, and has been shown to affect plasmodesmal permeability (Fig.  5, bottom left). When the cytosolic free calcium concentration was raised in staminal hairs of Setcreasea purpurea using microinjected calcium-loaded calcium chelator, fluorescent dye movement out of these cells was blocked for several hours, indicating that the increase in cytosolic free calcium stimulated closure of plasmodesmata (Tucker, 1990) . A  transient closure of plasmodesmata was also observed when calcium influx inducers mastoparan and inositol-1,4,5-triphosphate were microinjected into the staminal hairs (Tucker and Boss, 1996) . Notably, this plasmodesmal closure response was suppressed when the hair cells were pre-treated with the calcium channel blocker La3+, supporting the idea that calcium influx itself was the signal causing the plasmodesmal change. Further evidence linking calcium to plasmodesmal responses, comes from experiments using direct calcium injection or chilling treatments that induce an increase in cytosolic calcium (Holdaway-Clarke et  al., 2000) . The electrical resistance between sister maize suspension culture cells was monitored through the use of microelectrodes placed within the cytoplasm of each cell, assuming an increase in resistance indicative of plasmodesmal closure (Holdaway-Clarke et  al., 1996). When chilled the culture cells at 0–5  °C, plasmodesmata were closed and remained restricted until the temperature rose again, while accompanying a measured 2-fold increase in cytosolic Ca2+ concentration. However, when cells were injected, Ca2+ caused only a transient plasmodesmal closure, and the pores reopened within ten seconds. Assuming that the cytosolic calcium release induced in these experiments was physiologically relevant, callose deposition and degradation, which takes hours, is unlikely to be responsible for the plasmodesmal closure and reopening in response to calcium influx (Holdaway-Clarke et al., 2000; Tucker and Boss, 1996) . Instead, the authors suggested that calcium-dependent structural protein rearrangement might take place on this short time scale to alter plasmodesmal permeability (Fig. 5, bottom left). Intriguingly, several calcium-sensitive proteins have been found to localize to plasmodesmata. For example, immunogold labelling detected a 17-kDa centrin-like protein in the plasmodesmal neck regions of onion (Allium cepa) root tips and cauliflower (Brassica oleracea) florets (Blackman et al., 1999) . Centrins are a major part of filamentous scaffolds that contract upon binding to calcium in algal centrosomes and the basal regions of flagella (Geimer and Melkonian, 2005; Hu et al., 2004; Sanders and Salisbury, 1989) . Analogously, perhaps a centrin-containing scaffold configured across the plasmodesmal entranceway could quickly contract the pore neck region in response to intracellular calcium flux. Immunodetection also found calreticulin and myosin VIII at plasmodesmata in maize root cells (Baluska et al., 1999; Reichelt et al., 1999; Baluska et al., 2001) . Calreticulin is a calcium-binding protein with diverse roles in plants, including calcium signalling and sequestration (Jia et  al., 2009). Myosin VIII contains within its neck region potential binding domains for calmodulins (Knight and Kendrick-Jones, 1993), a calcium-binding family of intermediate messengers mediating transduction of calcium signalling (Ranty et al., 2006) . Finding that these proteins, along with actin (White et al., 1994) , localize at plasmodesmata attracts the hypothesis that an acto-myosin and centrin-filamentous system within the neck of plasmodesma translates the fluctuation of calcium flux exerted by calcium channels and calreticulin into rapid but transient contractions of the pore (Fig. 5, bottom left). Osmotic stress transiently increases plasmodesmal flux Plant root tips were shown to increase plasmodesmal flux temporarily and enhance sugar transport to osmotically stressed tissues (Schulz, 1994, 1995) . Electron microscopy of pea (Pisum sativum) root tip plasmodesmal ultrastructure showed that the diameter of the cytoplasmic sleeves in the cortical cells increased markedly, along with a loss in plasmodesmal neck constriction, after one hour of osmotic stress imposed using 350 mM mannitol (Schulz, 1995). However, this plasmodesmal response to mannitol was also transient, thereby the morphological alterations in root tip cortical plasmodesmata were reversed within a few hours. This result is reminiscent of previous observations that pea root tips subjected to osmotic stress displayed a transient up-regulation in the phloem unloading of 14C-sucrose into the root tip cortical cells after two hours (Schulz, 1994) . The author suggests that root tip plasmodesmal dilation during the initial osmotic stress response may increase the symplastic unloading of sugars into the stressed cells, to lower their water potential and help prevent plasmolysis. It is possible that the underlying mechanism involves callose degradation and re-deposition at plasmodesmata between root cells. Alternatively, calcium influx and/or ROS burst, both of which can be quickly and highly induced in response to mechanical stress of the plasma membrane during osmotic shock (Kurusu et al., 2013; Xiong and Zhu, 2002) , may play a role in regulating plasmodesmal aperture in a callose-independent manner. Cold and high light induce plasmodesmal remodelling The onset of cold temperatures not only slows overall plant growth but also can reduce photoassimilate transport rates. A  change in plasmodesmal ultrastructure or frequency is thought to relieve this stress by increasing sugar movement from source to sink tissues (Bilska and Sowinski, 2010) . Analyses of the plasmodesmal ultrastructure at the interface between Kranz mesophyll (KMS), bundle sheath (BS), and vascular parenchyma (VP) cells in cold-tolerant (CT) and cold-sensitive (CS) maize subjected to chilling temperatures revealed that CT maize could better maintain a state of open plasmodesmata compared with the CS lines. Intriguingly, a previous study comparing CS and CT maize found that plasmodesmal frequencies between KMS, BS, and VP cells increased by the greatest amount in the CT line grown at suboptimal temperatures, compared with CS plants. Under such conditions, the rate of photoassimilate movement into the transport path also increased in the CT line (Sowinski et  al., 2003) . These data suggest that enhancing plasmodesmal permeability allows the plants to better survive the cold environment. Plasmodesmata of the KMS/BS junction typically contain an internal ring of an unknown globular element called the internal sphincter (Bilska and Sowinski, 2010) . Compared with control CS maize grown in normal temperatures, CS lines subjected to chilling responded with a swelling of the internal sphincter. Following four hours of treatment, KMS/BS plasmodesmata of the CS lines had about 25% higher incidence of sphincter enlargement, compared with CT maize. The cross-area of the cytoplasmic sleeves in BS/BS and VP/VP also shrank significantly in the coldtreated CS maize, but remained unaffected in the CT lines. Immunogold labelling showed that callose and calreticulin in the plasmodesmal neck regions of KMS/BS and BS/VP interfaces greatly increased in cold-treated CS lines, but not in the CT lines. Finally, 14C transport capacity post-chilling was much reduced in the CS compared with CT maize, even after only 1 hour of chilling (Bilska and Sowinski, 2010) . These results indicate that CS maize respond very rapidly to chilling by blocking plasmodesmata. Although callose deposition probably accounts for the long-term sealing of plasmodesmata in the CS lines, cold-induced calcium flux has previously been shown to rapidly close plasmodesmata (Holdaway-Clarke et al., 2000) . The enhanced level of calreticulin found at the plasmodesmal neck regions in cold conditions in CS lines may represent a negative feedback response to sequester free calcium. Elucidation of the mechanisms used by the CT maize to maintain plasmodesmal opening and increase their frequency in response to chilling could be useful for improving cold-stress tolerance in many crop plants. It is notable that a putative member of the dehydrin protein family localizes to the plasmodesmal neck region in a highly cold-tolerant dogwood species (Karlson et al., 2003). Dehydrins are hypothesized to protect and stabilize other proteins and membranes against abiotic stresses caused by high salinity, drought, and cold. It will be interesting to determine whether these proteins indeed have a role in maintaining plasmodesmal connectivity in cold-tolerant species. High light intensity, a condition that can enhance the rate of photosynthesis, was also shown to elevate plasmodesmal density at KMS/BS/VP junctions (Sowinski et  al., 2007) , similar to cold-tolerant maize adaptation to chilling temperatures. C4 grasses were grown in low, medium, and high intensity light, and plasmodesmal density between KMS/BS and BS/VP junctions in each condition was compared, as well as the rate of photosynthate export. Higher light enhanced the export of photosynthate through the mesophyll into the vasculature, as measured using radiolabelled carbon dioxide (14CO2). Plasmodesmal density increased significantly between KMS/BS, and to a lesser extent between BS/VP junctions, as the light levels intensified (Sowinski et al., 2007) . Collectively, these results indicate that de novo biosynthesis of plasmodesmata may help alleviate the stresses caused by certain abiotic stresses by enhancing intercellular nutrient flux. Toxic metals and wounding trigger plasmodesmal closure Toxic metals have been shown to increase callose deposition at plasmodesmata. Treatments with 5–20  µM aluminium (Al3+) induced closure of plasmodesmata within three hours, as measured by microinjected LYCH diffusion in wheat (Triticum aestivum) root epidermal and tobacco mesophyll tissue (Sivaguru et al., 2000) . This response is callose-dependent, as it was preventable by pre-treatment with a callose synthesis inhibitor, 2-deoxy-d -glucose. Similarly, treatment with subtoxic levels of cadmium ions (Cd2+) can severely inhibit the cell-to-cell spread of turnip vein clearing virus, with 10 µM cadmium completely halting viral systemic movement (Citovsky et  al., 1998). Toxic metals other than aluminium and cadmium have also implicated in plasmodesmal closure. For example, six hours after treatment with lead cations (Pb2+), Lemna minor radial root cell walls and plasmodesmata had a high level of callose accumulation (Samardakiewicz et al., 2012) . Similar experiments were performed with arsenic (As3+; (Pirselova et  al., 2012) , but although aniline blue staining resulted in punctate signals within root cell walls in response to arsenic, whether these signals indicated plasmodesmata was not determined. Wound stresses have also been shown to induce accumulation of plasmodesmal callose (Radford et al., 1998) (Fig. 1F). Reversal of wound-induced plasmodesmal callose seems to involve Arabidopsis BETA-1,3-GLUCANASE PUTATIVE PLASMODESMATA-ASSOCIATED PROTEIN (AtBG_ ppap; Levy et  al., 2007). Aniline blue staining revealed that after wounding, there was higher accumulation of plasmodesmal callose in mutant leaves lacking AtBG_ppap compared with wild type. An AtBG_ppap–GFP fusion protein infiltrated into tobacco leaves confirmed that this glucanase could localize to both plasma and ER membranes, with a noticeably brighter punctate fluorescent pattern across cell perimeters that overlapped with plasmodesmal callose at cell–cell junctions. Bombardment of GFP into AtBG_ppap mutant and wild-type Arabidopsis leaves revealed that the overaccumulation of plasmodesmal callose in the mutant also blocked the cell-to-cell spread of GFP, supporting its effect on plasmodesmal permeability (Levy et al., 2007) . Perspectives Since plasmodesmata were first described some hundred years ago, plant biologists have been striving to elucidate their role in plant biology. Recent discoveries have demonstrated that plasmodesmata are not just simple “holes in the walls” but rather are highly dynamic intercellular communication channels, enabling plant cells to organize into a synchronized entity. However, it has also become evident that gaining cellular autonomy by severing or closing plasmodesmata serves as a crucial mechanism, not only for cell differentiation and specialization, but also for plant survival under drastic environmental conditions. Challenges ahead include identifying various intracellular signalling pathways and factors that convert primary intrinsic and extrinsic signals into alterations in plasmodesmal structure, formation, and permeability. Given the essential role of plasmodesmata in cell-to-cell transport, understanding how plasmodesmal regulation is integrated into specific cellular signalling pathways that enhance intercellular flux could provide insights into improving health and fitness of agriculturally important crops. Furthermore, advancing knowledge about various signalling factors that regulate plasmodesmata during environmental stress conditions could help improve traits such as pathogen resistance, drought and cold tolerance, and fruit/seed yield, in preparation for imminent climate changes. In this regard, it is inevitable that we will have to pay special attention to how plasmodesmal alterations bring about changes in distribution of the most critical plant signalling molecules, hormones, and it will not be surprising to find more crosstalk between plasmodesmata and hormonal signalling beyond SA and GA. Excitingly, a very recent finding showed that auxin may regulate callose synthase GSL8 to prevent its own diffusion out of cells during the phototropic response (Han et al., 2014) . As our understanding advances, complex feed-forward and feed-back regulatory circuits may prove to be a common theme linking hormonal and other signalling pathways to plasmodesmal control. Plasmodesmal research is one of the most vital, but extremely challenging, fields of biology. Plasmodesmata are recalcitrant to compositional and anatomical studies at the molecular level, owing to the difficulties associated with isolating them not only as biochemically intact entities but also as genetic mutants. However, the refinement of certain techniques, such as mass spectroscopy-based plasmodesmal protein identification from plasmodesmata-enriched cell wall fractions, improved fluorescent-tagging-based localization studies, and genetic approaches, has provided much-needed insights into the role of plasmodesmata in developmental and intercellular signalling processes. Thus, although the field is still far from grasping the complete picture of the molecular anatomy and genetic networks of plasmodesmata, the future is bright. 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Ross Sager, Jung-Youn Lee. Plasmodesmata in integrated cell signalling: insights from development and environmental signals and stresses, Journal of Experimental Botany, 2014, 6337-6358, DOI: 10.1093/jxb/eru365