Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2025 Jun 9.
Published in final edited form as: Results Probl Cell Differ. 2024;73:73–86. doi: 10.1007/978-3-031-62036-2_4

Communicating Across Cell Walls: Structure, Evolution, and Regulation of Plasmodesmatal Transport in Plants

Jacob O Brunkard 1
PMCID: PMC12147918  NIHMSID: NIHMS2087441  PMID: 39242375

Abstract

Plasmodesmata are conduits in plant cell walls that allow neighboring cells to communicate and exchange resources. Despite their central importance to plant development and physiology, our understanding of plasmodesmata is relatively limited compared to other subcellular structures. In recent years, technical advances in electron microscopy, mass spectrometry, and phylogenomics have illuminated the structure, composition, and evolution of plasmodesmata in diverse plant lineages. In parallel, forward genetic screens have revealed key signaling pathways that converge to regulate plasmodesmatal transport, including chloroplast-derived retrograde signaling, phytohormone signaling, and metabolic regulation by the conserved eukaryotic Target of Rapamycin kinase. This review summarizes our current knowledge of the structure, evolution, and regulation of plasmodesmatal transport in plants.

4.1. Introduction

Plant cells are connected by plasmodesmata (PD), narrow membrane-lined channels in the cell wall that connect adjacent cytoplasts (Faulkner 2018). Large and small molecules move through PD, including metabolites, signaling molecules, small RNAs, and proteins up to ~80 kDa (Crawford and Zambryski 2001; Brunkard et al. 2015a; Paultre et al. 2016). Even larger molecules can move through PD through specialized mechanisms, such as viruses, which coopt PD to spread beyond infected cells using “movement proteins” (MPs) that modify PD transport to accommodate large viral genomes and associated proteins (Burch-Smith and Zambryski 2016; Brunkard and Zambryski 2017). Given the wide range of molecules that can move through PD, the safest “null hypothesis” in plant cell biology is that any cytosolic molecule under 100 kDa might traffic between cells until shown otherwise.

PD are essential for land plant physiology and development. No mutants, variants, or species of land plants have been identified that lack PD, and any strong perturbation of PD transport has major deleterious effects on plant biology (Kim et al. 2002; Benitez-Alfonso et al. 2009). Only highly specialized cells are not connected by PD, such that almost the entire plant is comprised of continuous cytoplasts, called the “symplast”, with the cell walls and spaces between cells comprising a continuous extracellular space, called the “apoplast” (Faulkner 2018). Although most cells are connected by PD, the rate and properties of molecules that can move between cells are tightly regulated in different cell types and in response to environmental cues. In recent years, more and more plant cell biologists have focused on investigating how plants adjust PD transport and how these adjustments influence developmental patterning, metabolism, disease susceptibility, and agronomic traits.

Here, I will briefly summarize our current understanding of PD biology, focusing on the PD themselves rather than the many biological functions and cargo of PD, which have been reviewed extensively elsewhere (Brunkard et al. 2015a, 2013; Brunkard and Zambryski 2017; Tee and Faulkner 2024; Bayer and Benitez-Alfonso 2024; Reagan et al. 2018a). Specifically, I will present contemporary knowledge of the structure and formation of PD, the molecular composition of PD, the evolutionary history of PD in photosynthetic organisms, and the regulation of PD transport by diverse signaling cues.

4.2. PD Structure and Formation

In land plants, PDs are composed of two membrane layers: an outer membrane continuous with the plasma membrane and an inner “desmotubule” continuous with the endoplasmic reticulum (ER). The desmotubule is tightly compressed to a diameter of ~10–15 nm, with minimal intercellular transport between cells mediated by the desmotubule (Peters et al. 2021; Ostermeyer et al. 2022). Between the desmotubule and the PD outer membrane is the “cytosolic sleeve”, which varies widely in size and morphology. Most intercellular trafficking occurs within the cytosolic sleeve (Crawford and Zambryski 2000). Membrane-associated molecules may also move between cells via PD (Ishiwatari et al. 1998; Meng et al. 2010), although this is less studied than the trafficking of cytosolic molecules.

PDs are divided into two classes by the mechanism of their formation: primary PDs and secondary PDs (Meeuse 1941). Primary PDs form during cell division when strands of ER are trapped in the developing phragmoplast (Knox et al. 2015; Kriechbaumer et al. 2015; Hepler 1982), which is an assembly of cytoskeletal proteins, membranes, and vesicles filled with carbohydrates that establish the plane of a nascent plant cell wall during cytokinesis. Secondary PDs form after cell division, whether de novo or through fission of existing PD (Burch-Smith et al. 2011a). Both primary and secondary PD can adopt a range of morphologies, including simple channels, highly branched structures, and funnel shapes, among others, that presumably influence the size and rate of particles moving through each PD (Ostermeyer et al. 2022).

Studying PD morphology has proven challenging due to various technical limitations. PDs are substantially smaller than the diffraction limit of light, and even cutting-edge superresolution light microscopy (Fitzgibbon et al. 2010) only achieves resolutions of ~20nm (Schermelleh et al. 2019) that are insufficient to visualize PD structure. Transmission electron microscopy (TEM) can achieve the necessary resolution to observe PD, but TEM cannot be used to visualize living tissues; moreover, classical fixation and staining techniques for TEM introduced various artifacts (Gilkey and Staehelin 1986) that misled early efforts to define PD ultrastructure. Recent advances in electron microscopy (EM), such as TEM tomography, focused ion beam milling coupled with scanning electron microscopy (FIB-SEM) (Reagan et al. 2018b, 2022), and serial block face SEM (SBF-SEM) (Paterlini and Belevich 2022), are finally allowing us to accurately visualize PD ultrastructure at high resolution and in large numbers so that we can begin to deeply investigate the role of PD morphology in plant cell biology (Sankoh and Burch-Smith 2021). Nonetheless, given that PD ultrastructure can currently only be accurately visualized in fixed tissues, we can only use these techniques to begin drawing correlations between PD structure and function.

4.3. Molecular Composition of PD in Land Plants

PDs are comprised of three primary classes of molecules: polysaccharides in the cell wall, lipids in the outer membrane and desmotubule, and proteins anchored in the surrounding cell wall, outer membrane, cytosolic sleeve, or desmotubule. Early investigations of PD using microscopy already revealed several key components of the PD structure, which have now been further elucidated using biochemical approaches. With rare exceptions, PD do not form wholly distinct structures composed of unique features in the sense that, for instance, many chloroplast proteins are only found in chloroplasts. Instead, PD are “enriched” for some molecular components over others, without clear-cut defining features, and there is likely considerable diversity in the degree of this enrichment even when comparing PD in the same tissue.

4.3.1. Polysaccharides

The cell wall surrounding PD is distinguished by the frequent presence of callose, a class of polysaccharides composed primarily of β−1,3,-glucan linkages, that surrounds the PD membranes (Esau 1939, 1950; Currier 1957). Large callose deposits in the cell wall around PD prevent intercellular transport, leading to the popular model that callose “constricts” the PD channel. Callose levels surrounding PD vary depending on cell type and physiological status, dynamically regulating intercellular transport between plant cells (Zavaliev et al. 2011). Beyond callose, immunohistological and biochemical studies suggest that PD cell walls are enriched for a handful of other polysaccharides, such as relatively flexible forms of the pectins homogalacturonan and rhamnogalacturon-I, as well as fucosylated xyloglucans (Knox and Benitez-Alfonso 2014; Paterlini et al. 2022). Unlike callose, the functional relevance of these enrichments has not been established and is likely less dynamic, but they are proposed to contribute to PD biogenesis and/or structural plasticity; for example, mutants with reduced cellulose and increased pectin content in cell walls form more PD, suggesting that these polysaccharides could broadly impact PD biogenesis (Okawa et al. 2023).

4.3.2. Membrane Lipids

Due to their distinct structures and functions, PD membranes are exemplary models for investigating lateral segregation of lipids and proteins (i.e., “lipid rafts”) within membranes. Biochemical investigations of PD membrane fractions from Arabidopsis roots and cell suspension cultures have revealed that PD outer membranes are enriched for sterols and complex sphingolipids containing very-long-chain fatty acids (Grison et al. 2015). Overall, there are fewer glycerolipids and glycerophospholipids in PD outer membranes than in the rest of the plasma membrane, and the phospholipids that are found in PD tend to be more highly saturated. Genetically or chemically inhibiting sterol and/or sphingolipid biosynthesis deregulates PD trafficking and prevents proper sorting of PD-enriched proteins to PD membranes, suggesting that the lipid composition of PD outer membranes could play regulatory roles in PD transport (Yan et al. 2019; Dettmer et al. 2014; Kraner et al. 2017). In contrast to the PD outer membranes, relatively little is known about the lipid composition of desmotubules that are continuous with ER. Remarkably, the desmotubule membranes are often appressed to form a rod only ~10 nm in diameter; since lipid bilayers are ~5 nm in width, this is effectively the minimum possible diameter for a cylinder of lipid bilayer membranes and therefore must involve substantial reorganization of membrane proteins and lipids to maintain its morphology.

4.3.3. Proteins

All plant viruses that spread beyond the vascular system use specialized movement proteins that facilitate the transport of their genomes between cells through PD. Many movement proteins accumulate at PD, often showing a selective preference for some PD over others; for example, Tobacco mosaic virus (TMV) P30 accumulates at diverse PD, whereas Potato leaf roll virus (PLRV) P17 accumulates only at PD with branched morphologies (Burch-Smith et al. 2011a). TMV P30 is the model viral movement protein that drives the movement of the single-stranded (ss) RNA genome from infected cells into uninfected cells (Citovsky et al. 1990), where the TMV ssRNA is translated by the host, replicates, and spreads. Despite nearly four decades of intensive research, we still have a minimal understanding of how P30 alters PD to drive TMV ssRNA trafficking. Genetic and molecular approaches have, however, identified short amino acid sequences in P30 that are necessary and sufficient for P30 to accumulate at PD, in a process that is proposed to depend on synaptotagmins that establish ER-plasma membrane junctions (Yuan et al. 2016, 2018).

Many plant proteins also accumulate primarily at PD, which have been identified primarily from two complementary approaches. In the first approach, putative membrane-associated proteins were cloned and internally fused with fluorescent proteins to avoid interfering with signal peptides or post-translational modifications at the N- and C-termini of the proteins (Thomas et al. 2008; Simpson et al. 2009). These fluorescent proteins were then screened for enrichment at puncta in the cell walls and further validated by colocalization with established PD markers, such as high callose deposits or TMV P30 fused to fluorescent proteins. In the second approach, proteins were extracted from PD-enriched membranes isolated from cell wall fractions and then analyzed using mass spectrometry (Fernandez-Calvino et al. 2011; Lee et al. 2011; Johnston et al. 2023; Gombos et al. 2023). Both approaches revealed similar sets of highly-enriched PD proteins, including PD CALLOSE-BINDING PROTEINs (PDCBs), PD-LOCALIZED PROTEINS (PDLPs), and PD β-GLUCANASES (PDBGs), among others. These proteins all associate with the PD outer membrane, participate in the synthesis or stability of callose at PD, and can dynamically regulate PD transport. PDLP5, for example, is induced upon recognition of a pathogen infection and promotes callose synthesis at PD to effectively isolate infected cells from the rest of the symplasm (Lee et al. 2011; Wang et al. 2013).

On the desmotubule side, a class of proteins called MCTPs (Multiple C2 domain and Transmembrane region Proteins) accumulate at PD. These MCTPs are anchored in the desmotubule ER by C-terminal transmembrane domains and then extend via N-terminal C2 domains to associate with phospholipids at the surface of the PD outer membrane. MCTPs are one of three classes of PD-enriched proteins, along with β-glucanases and tetraspanins, that are found in both the model flowering plant Arabidopsis thaliana and the model moss Physcomitrium patens, hinting that they evolved functions at PD when PD first arose in the land plant lineage (Johnston et al. 2023).

Using less stringent definitions, many proteins that are otherwise found in the plasma membrane, endoplasmic reticulum, cell wall, or cytoplasm may sometimes localize to PD. Proteins with C-terminal glycosylphosphatidylinositol (GPI) anchors are often enriched at PD, for instance, likely because GPI anchors tend to sort with sphingolipids that are also enriched at PD (Zavaliev et al. 2016). The GPI anchors of PDCBs are necessary and sufficient for PDCBs to localize to PD outer membranes, for example. Given the small size of PD, however, it is unlikely that very many proteins are consistently resident within PD, and even fewer are likely to directly regulate PD transport. New approaches for defining subcellular proteomes, such as biotin ligase-mediated proximity labeling, may expand our understanding of the dynamic PD proteome in diverse tissues.

4.4. Repeated Evolution of PD in Multicellular Algae

Land plants evolved from a common green algal ancestor approximately 500 million years ago and subsequently diverged into several groups, including bryophytes (hornworts, mosses, liverworts) and polysporangiophytes (ferns, lycophytes, gymnosperms, and flowering plants); the precise sequence of events during the early diversification of land plants remains under debate (Donoghue et al. 2021; Brunkard et al. 2015b; Harris et al. 2022; Bowles et al. 2023). Phylogenomic studies over the last decade agree, however, that the sister group of land plants is most likely Zygnematophycheae, a class of green algae. These algae are unicellular or from simple filamentous strands that are not connected by any PD-like structures in the cell walls. Land plants, in contrast, all have complex, three-dimensional (“parenchymatous”) multicellular life stages, and they all form PD in the cell wall. The frequency, morphology, and functions of PD may vary considerably across land plant lineages, but they share the same basic features: outer membranes, a central desmotubule, and a cytosolic sleeve between these membranes that allow diverse molecules to move between cells (Brunkard and Zambryski 2017).

Several other lineages of multicellular algae have evolved PD-like structures, including the aquatic green algal classes Charophyceae and Coleochaetophyceae, multiple lineages of more distantly related chlorophyte green algae, and lineages of red and brown algae (Niklas and Newman 2013; Wegner et al. 2023). These PD do not have a desmotubule continuous with the ER, although they are similarly thought to form by incomplete cytokinesis, and some algal PD have central structures of unknown composition. Strikingly, PD-like structures are present in every lineage with cellulosic cell walls that have evolved complex multicellularity. This suggests that PD-like structures are critical for central physiological processes in multicellular organisms separated by cellulosic cell walls. A deeper understanding of the composition, formation, and functions of PD-like structures in green and brown algae will be needed to determine whether these structures evolved through convergence or in parallel, i.e., whether the genetic “toolbox” for PD already existed in their last common ancestor (parallelism) or if PD have evolved independently as analogous structures (convergence).

4.5. Dynamic Regulation of PD Transport

Bulk rates of PD transport are dynamically regulated by diverse cues, including a wide range of physiological and developmental conditions. Here, I will focus on how genetic approaches have revealed unexpected regulatory pathways that are major regulators of PD trafficking between plant cells: organelle-nucleus-PD signaling (ONPS) (Burch-Smith et al. 2011b; Azim and Burch-Smith 2020), TARGET OF RAPAMYCIN (TOR) metabolic signaling (Brunkard et al. 2020; Brunkard 2020), and phytohormone signaling. The connections between these pathways and PD were revealed by a forward genetic screen for increased rates of PD transport in Arabidopsis thaliana embryos (Kim et al. 2002), among other approaches. Mutants with increased PD transport are called “ise” mutants, including four mutants identified through forward genetic approaches and several more subsequently identified through reverse genetics.

4.5.1. Organelle-nucleus Cross-talk Regulates PD Transport

The first two ise mutants, ise1 and ise2, each encode RNA helicases: ISE1 is a mitochondrial RNA helicase (Stonebloom et al. 2009) and ISE2 is a plastid RNA helicase (Burch-Smith et al. 2011b). Transcriptomic and ultrastructural analyses of ise1 and ise2 mutants demonstrated that both mutants impact chloroplast-to-nucleus retrograde signaling pathways, disrupting the expression of photosynthesis-associated nuclear genes (Burch-Smith et al. 2011b). Taking a reverse genetic approach to test this model, multiple groups demonstrated that PD transport rates are elevated in other mutants defective in chloroplast genome expression that trigger chloroplast-to-nucleus retrograde signaling, including a uL15c ribosomal protein mutant (Bobik et al. 2019) and a clpr2 protease mutant (Carlotto et al. 2016). Knocking down or knocking out expression of ISE1 or ISE2 increases the number of PD connecting adjacent cells, including a substantial increase in secondary PD that form after cytokinesis, suggesting that chloroplast-to-nucleus retrograde signals regulate PD transport, in part, by triggering de novo PD biogenesis (Burch-Smith and Zambryski 2010).

A parallel genetic screen for altered PD trafficking in Arabidopsis roots identified another mutant defective in chloroplast physiology, thioredoxin m3 (trx-m3) (Benitez-Alfonso et al. 2009). m-type thioredoxins participate in photosynthesis and regulate redox status within the chloroplast. Trx-m3 mutants generate large quantities of reactive oxygen species (ROS) that cause broad oxidative stress in the cell, triggering callose production at PD and drastically reducing PD transport. In contrast, ise2 mutants do not cause oxidative stress; instead, the redox potential of ise2 mutant chloroplasts is significantly reduced and does not strongly impact the cytosolic or mitochondrial redox potentials (Stonebloom et al. 2012). This discovery hinted that PD might be responsive to multiple chloroplast-associated physiological cues. Indeed, subsequent studies revealed that PD transport is dynamically regulated by diurnal light and dark cues and that rates of PD trafficking are coordinated by the circadian clock (Brunkard and Zambryski 2019). Moreover, light- and redox-sensitive stroma-containing tubular extensions from the chloroplast, called “stromules”, often associate with PD and could act as conduits for direct chloroplast-to-PD signaling (Brunkard et al. 2015c).

4.5.2. Target of Rapamycin Regulates PD Transport

The next two ise mutants, ise3 and ise4, revealed that PD transport is tightly coordinated by the Target of Rapamycin (TOR) eukaryotic metabolic signaling hub (Brunkard et al. 2020). TOR is a broadly conserved protein kinase that coordinates growth and development with nutrient availability and environmental cues (Brunkard 2020). When conditions are favorable (e.g., nutrients are plentiful), TOR is active and triggers kinase signaling cascades to promote growth; when conditions are unfavorable (e.g., nutrients are limiting), TOR becomes inactive and cells become quiescent, relying on nutrient recycling pathways, such as autophagy, to sustain life. TOR and its direct interacting partners, RAPTOR and LST8, are extremely well-conserved across all eukaryotic lineages; the upstream and downstream signaling pathways that converge on TOR are more variable in plants, animals, fungi, and protists. Although the mechanisms of nutrient-sensing may vary, one consistent feature of TOR signaling across eukaryotes is that TOR monitors the availability of amino acids, nucleotides, and ATP.

ise3 encodes a plant-specific subunit of the mitochondrial oxidative phosphorylation (OXPHOS) chain of unknown yet essential molecular function (Brunkard et al. 2020). ise4 encodes the plant orthologue of Reptin (sometimes called RuvBL2, RVB2, or Tip49b, among other names), a universally conserved ATPase distantly homologous to bacterial RuvB (Dauden et al. 2021). Reptin and its interacting partner, Pontin, play diverse roles in eukaryotic cells, including as scaffolds for chromatin remodelers in the nucleus, but the ATPase activities of Reptin and Pontin are thought to only be strictly required for their role as HSP90 co-chaperones in the R2TP prefoldin-like complex (Chatterjee et al. 2021). R2TP, which is composed of Reptin, Pontin, an HSP90-binding protein Spaghetti (sometimes called RPAP3 or Tah1), and sometimes PIH1, participates in the HSP90-dependent assembly of the TOR kinase complex. In mammals and possibly also in plants, Reptin and Pontin likely act as ATP sensors, only allowing the active TOR kinase complex to form when sufficient ATP is available (Kim et al. 2013). Although null alleles of reptin are lethal at the earliest stages of plant gametophyte development, ise4 specifically disrupts the ATP-binding Walker A motif of Reptin, which is proposed to limit its ability to activate TOR and thus interfere with plant growth and development (Brunkard et al. 2020).

As expected, therefore, TOR activity is substantially reduced in both the ATP synthesis-deficient ise3 mutant and R2TP-deficient ise4 mutant (Brunkard et al. 2020). Chemical and reverse genetic approaches confirmed that metabolic cues upstream of TOR, including glycolysis, OXPHOS, and the Spaghetti subunit of R2TP all regulate PD transport. Moreover, directly disrupting TOR signaling in lst8 mutants, knocking down LST8 expression, or attenuating TOR activity with several classes of selective kinase inhibitors all increase transport through PD in leaves and/or embryos. This demonstrates that the ancestral TOR signaling network has evolved a new role in land plants as a key regulator of PD trafficking. The mechanism(s) of TOR-PD signaling and the consequences of TOR-restricted PD transport are currently under investigation, with apparent implications for the reallocation of photo-synthesized sugars from “source” leaves to actively growing “sink” tissues, like root tips, shoot tips, and young leaves (Brunkard et al. 2020; Brunkard 2020).

4.5.3. Phytohormone Regulation of PD Trafficking

Plant development and physiology are regulated by small signaling molecules analogous to mammalian hormones, called “phytohormones”. These include auxins, cytokinins, ethylene, gibberellins, abscisic acid, jasmonic acid, salicylic acid, brassinosteroids, and strigolactones, among many others. Phytohormones act both locally and over distances to modulate responses to external cues and establish developmental patterns. Phytohormones move between cells using multiple routes, including through the cytosolic sleeve of PD (remaining in the symplast) or through plasma membrane exporters and importers (moving through the apoplast, sometimes called “carrier-mediated transport”) (Band 2021; Rutschow et al. 2011). PD trafficking is generally passive and directionless, allowing phytohormones to spread beyond the site of their synthesis, but also potentially “diluting” a phytohormone’s local concentration to limit its efficacy. Plasma membrane exporters and importers often directionally traffic phytohormones to specific locations; for example, the auxin PIN-FORMED-like exporters show polar subcellular localization that drives directional trafficking and concentration of auxins (Wiśniewska et al. 2006; Ung et al. 2022). Thus, depending on the context, dynamic changes in PD transport rates may enhance or repress phytohormone signaling; similarly, many phytohormones regulate PD transport as a potential feedback mechanism to amplify or curb their local efficacy.

Auxin signaling is a prime example of the tension between symplastic and apoplastic phytohormone trafficking. Auxins accumulate at the site of lateral root initiation through carrier-mediated transport (i.e., via the apoplast), stimulating cell divisions that are critical for the lateral root to establish and emerge (Thomas et al. 2008; Simpson et al. 2009). During lateral root initiation, large callose deposits accumulate at PD and restrict intercellular transport to these differentiating cells (Maule et al. 2013; Benitez-Alfonso et al. 2013). Analysis of mutants over- or under-expressing the callose-degrading enzymes, β−1,3-glucanases, and the callose deposition-promoting proteins, PDCBs, demonstrated that localized callose deposition is critical for the initiation of lateral roots. Moreover, when callose deposition is disrupted, auxin signaling responses become diffuse in the root, suggesting that the symplastic isolation of initiating lateral roots is necessary to allow carrier-mediated transport to concentrate auxin in these differentiating cells without “backflow” transport through the symplast. Later studies expanded on this discovery by showing that auxin activates the expression of callose synthases to locally restrict PD transport, effectively amplifying auxin concentration in cells during developmental patterning (Han et al. 2014).

Many other phytohormones influence PD transport, including recent demonstrations that cytokinins and brassinosteroids are key regulators of PD trafficking. Comparative analysis of all four ise mutants hinted that cytokinin signaling is upregulated in all of these embryos with increased PD transport, which led to the hypothesis that cytokinins promote PD transport (Horner and Brunkard 2021). Indeed, treating leaves with cytokinins at physiologically relevant concentrations is sufficient to stimulate large increases in PD transport between cells, and genetically disrupting cytokinin signaling is sufficient to restrict intercellular trafficking, demonstrating that cytokinins regulate PD transport in leaves. Recent studies of shoot meristems in Arabidopsis have indicated that cytokinin signaling is upregulated when TOR is inactive (Kong et al. 2023) and that auxin can promote TOR activity (Li et al. 2017); moreover, it is well-established that cytokinins and auxins often act in opposition (Bishopp et al. 2011; Moubayidin et al. 2009). This hints that TOR, auxins, and cytokinins might all work in intersecting pathways to regulate PD transport during organogenesis in shoot meristems.

In Arabidopsis roots, the enzymes that are required for brassinosteroid biosynthesis are spatially separated into different cell types (Wang et al. 2023). Brassinosteroid precursors are synthesized in outer cell files of the root and then move via PD to the central stele, where an enzyme catalyzes the last step to make the active brassinosteroid, brassinolide. Brassinolide, in turn, promotes callose deposition and restricts PD transport between the inner stele and outer cell files, which is proposed to act as a negative feedback loop to regulate brassinosteroid accumulation. Brassinosteroids are monitored by cell surface receptors that signal through receptor-like kinase cascades to modulate growth and development; this discovery elegantly shows how PD can add regulatory layers to the metabolic synthesis of key regulatory molecules and fine-tune developmental patterning.

4.6. Conclusions

The repeated, independent evolution of multicellularity in eukaryotic lineages has followed different trajectories, influenced by the cellular structures and physiology of their ancestors. PD enabled the evolution of multicellularity in lineages with cellulosic cell walls, and PD have evolved to play complex roles in physiology and developmental patterning. Innovative, orthogonal approaches to investigate PD, including cutting-edge electron microscopy methods, improved mass spectrometry techniques to chemically define PD, and sophisticated forward genetic screens for regulators of PD transport have made huge progress in our understanding of PD over the last decade. Ongoing efforts to understand the diversity of PD within organisms and across species, the dynamic regulation of PD transport by processes beyond callose deposition in the cell wall, and the cross-talk between PD trafficking and cellular homeostasis, metabolism, and development will be crucial to deepen our fundamental understanding of plant biology and contribute to the social project of improving crop species that are urgently needed for a sustainable agricultural future (Lutt and Brunkard 2022; Springmann et al. 2018).

References

  1. Azim MF, Burch-Smith TM (2020) Organelles-nucleus-plasmodesmata signaling (ONPS): an update on its roles in plant physiology, metabolism and stress responses. Curr Opin Plant Biol 58:48–59 [DOI] [PubMed] [Google Scholar]
  2. Band LR (2021) Auxin fluxes through plasmodesmata. New Phytol 231:1686–1692 [DOI] [PubMed] [Google Scholar]
  3. Bayer EM, Benitez-Alfonso Y (2024) Plasmodesmata: channels under pressure. Annu Rev Plant Biol 75(1). 10.1146/annurev-arplant-070623-093110 [DOI] [PubMed] [Google Scholar]
  4. Benitez-Alfonso Y et al. (2009) Control of Arabidopsis meristem development by thioredoxin-dependent regulation of intercellular transport. Proc Natl Acad Sci USA 106:3615–3620 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Benitez-Alfonso Y et al. (2013) Symplastic intercellular connectivity regulates lateral root patterning. Dev Cell 26:136–147 [DOI] [PubMed] [Google Scholar]
  6. Bishopp A et al. (2011) A mutually inhibitory interaction between auxin and cytokinin specifies vascular pattern in roots. Curr Biol 21:917–926 [DOI] [PubMed] [Google Scholar]
  7. Bobik K et al. (2019) The essential chloroplast ribosomal protein uL15c interacts with the chloroplast RNA helicase ISE2 and affects intercellular trafficking through plasmodesmata. New Phytol 221:850–865 [DOI] [PubMed] [Google Scholar]
  8. Bowles AMC, Williamson CJ, Williams TA, Lenton TM, Donoghue PCJ (2023) The origin and early evolution of plants. Trends Plant Sci 28:312–329 [DOI] [PubMed] [Google Scholar]
  9. Brunkard JO (2020) Exaptive evolution of target of Rapamycin signaling in multicellular eukaryotes. Dev Cell 54:142–155 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Brunkard JO, Zambryski PC (2017) Plasmodesmata enable multicellularity: new insights into their evolution, biogenesis, and functions in development and immunity. Curr Opin Plant Biol 35:76–83 [DOI] [PubMed] [Google Scholar]
  11. Brunkard JO, Zambryski P (2019) Plant cell-cell transport via plasmodesmata is regulated by light and the circadian CLOCK. Plant Physiol 181:1459–1467 [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Brunkard JO, Runkel AM, Zambryski PC (2013) Plasmodesmata dynamics are coordinated by intracellular signaling pathways. Curr Opin Plant Biol 16:614–620 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Brunkard JO, Runkel AM, Zambryski PC (2015a) The cytosol must flow: intercellular transport through plasmodesmata. Curr Opin Cell Biol 35:13–20 [DOI] [PubMed] [Google Scholar]
  14. Brunkard JO, Runkel AM, Zambryski PC (2015b) Comment on ‘a promiscuous intermediate underlies the evolution of LEAFY DNA binding specificity’. Science 347:621. [DOI] [PubMed] [Google Scholar]
  15. Brunkard JO, Runkel AM, Zambryski PC (2015c) Chloroplasts extend stromules independently and in response to internal redox signals. Proc Natl Acad Sci USA 112:10044–10049 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Brunkard JO et al. (2020) TOR dynamically regulates plant cell-cell transport. Proc Natl Acad Sci USA 117:5049–5058 [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Burch-Smith TM, Zambryski PC (2010) Loss of increased size exclusion limit (ise)1 or ise2 increases the formation of secondary plasmodesmata. Curr Biol 20:989–993 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Burch-Smith TM, Zambryski PC (2016) Regulation of plasmodesmal transport and modification of plasmodesmata during development and following infection by viruses and viral proteins. In: Plant-virus interactions: molecular biology, intra- and intercellular transport. Springer International Publishing, Cham, pp 87–122. 10.1007/978-3-319-25489-0_4 [DOI] [Google Scholar]
  19. Burch-Smith TM, Stonebloom S, Xu M, Zambryski PC (2011a) Plasmodesmata during development: re-examination of the importance of primary, secondary, and branched plasmodesmata structure versus function. Protoplasma 248:61–74 [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Burch-Smith TM, Brunkard JO, Choi YG, Zambryski PC (2011b) Organelle-nucleus cross-talk regulates plant intercellular communication via plasmodesmata. Proc Natl Acad Sci USA 108:E1451–E1460 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Carlotto N et al. (2016) The chloroplastic DEVH-box RNA helicase increased size exclusion limit 2 involved in plasmodesmata regulation is required for group II intron splicing. Plant Cell Environ 39:165–173 [DOI] [PubMed] [Google Scholar]
  22. Chatterjee S, Xu M, Shemyakina EA, Brunkard JO (2021) Pontin/Reptin-associated complexes differentially impact plant development and viral pathology. bioRxiv. 10.1101/2021.11.29.470430 [DOI] [Google Scholar]
  23. Citovsky V, Knorr D, Schuster G, Zambryski P (1990) The P30 movement protein of tobacco mosaic virus is a single-strand nucleic acid binding protein. Cell 60:637–647 [DOI] [PubMed] [Google Scholar]
  24. Crawford KM, Zambryski PC (2000) Subcellular localization determines the availability of non-targeted proteins to plasmodesmatal transport. Curr Biol 10:1032–1040 [DOI] [PubMed] [Google Scholar]
  25. Crawford KM, Zambryski PC (2001) Non-targeted and targeted protein movement through plasmodesmata in leaves in different developmental and physiological states. Plant Physiol 125:1802–1812 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Currier HB (1957) Callose substance in plant cells. Am J Bot 44:478–488 [Google Scholar]
  27. Dauden MI, López-Perrote A, Llorca O (2021) RUVBL1–RUVBL2 AAA-ATPase: a versatile scaffold for multiple complexes and functions. Curr Opin Struct Biol 67:78–85 [DOI] [PubMed] [Google Scholar]
  28. Dettmer J et al. (2014) Choline transporter-like1 is required for sieve plate development to mediate long-distance cell-to-cell communication. Nat Commun 5:4276–4276 [DOI] [PubMed] [Google Scholar]
  29. Donoghue PCJ, Harrison CJ, Paps J, Schneider H (2021) The evolutionary emergence of land plants. Curr Biol 31:R1281–R1298 [DOI] [PubMed] [Google Scholar]
  30. Esau K (1939) Development and structure of the phloem tissue. Bot Rev 5:373–432 [Google Scholar]
  31. Esau K (1950) Development and structure of the phloem tissue II. Bot Rev 16:67–114 [Google Scholar]
  32. Faulkner C (2018) Plasmodesmata and the symplast. Curr Biol 28:R1374–R1378 [DOI] [PubMed] [Google Scholar]
  33. Fernandez-Calvino L et al. (2011) Arabidopsis plasmodesmal proteome. PLoS One 6:e18880–e18880 [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Fitzgibbon J, Bell K, King E, Oparka K (2010) Super-resolution imaging of plasmodesmata using three-dimensional structured illumination microscopy. Plant Physiol 153:1453–1463 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Gilkey JC, Staehelin LA (1986) Advances in ultrarapid freezing for the preservation of cellular ultrastructure. J Elec Microsc Tech 3:177–210 [Google Scholar]
  36. Gombos S et al. (2023) A high-confidence Physcomitrium patens plasmodesmata proteome by iterative scoring and validation reveals diversification of cell wall proteins during evolution. New Phytol 238:637–653 [DOI] [PubMed] [Google Scholar]
  37. Grison MS et al. (2015) Specific membrane lipid composition is important for plasmodesmata function in arabidopsis. Plant Cell 27:1228–1250 [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Han X et al. (2014) Auxin-callose-mediated plasmodesmal gating is essential for tropic auxin gradient formation and signaling. Dev Cell 28:132–146 [DOI] [PubMed] [Google Scholar]
  39. Harris BJ et al. (2022) Divergent evolutionary trajectories of bryophytes and tracheophytes from a complex common ancestor of land plants. Nat Ecol Evol 6:1634–1643 [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Hepler PK (1982) Endoplasmic reticulum in the formation of the cell plate and plasmodesmata. Protoplasma 111:121–133 [Google Scholar]
  41. Horner W, Brunkard JO (2021) Cytokinins stimulate plasmodesmatal transport in leaves. Front Plant Sci 12:674128–674128 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Ishiwatari Y et al. (1998) Rice phloem thioredoxin h has the capacity to mediate its own cell-to-cell transport through plasmodesmata. Planta 205:12–22 [DOI] [PubMed] [Google Scholar]
  43. Johnston MG et al. (2023) Comparative phyloproteomics identifies conserved plasmodesmal proteins. J Exp Bot 74:1821–1835 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Kim I, Hempel FD, Sha K, Pfluger J, Zambryski PC (2002) Identification of a developmental transition in plasmodesmatal function during embryogenesis in Arabidopsis thaliana. Development 129:1261–1272 [DOI] [PubMed] [Google Scholar]
  45. Kim SG et al. (2013) Metabolic stress controls mTORC1 lysosomal localization and dimerization by regulating the TTT-RUVBL1/2 complex. Mol Cell 49:172–185 [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Knox JP, Benitez-Alfonso Y (2014) Roles and regulation of plant cell walls surrounding plasmodesmata. Curr Opin Plant Biol 22:93–100 [DOI] [PubMed] [Google Scholar]
  47. Knox K et al. (2015) Putting the squeeze on plasmodesmata: a role for reticulons in primary plasmodesmata formation. Plant Physiol 168:1563–1572 [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Kong S et al. (2023) DRMY1 promotes robust morphogenesis by sustaining translation of a hormone signaling protein. Preprint at 10.1101/2023.04.07.536060 [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Kraner ME et al. (2017) Choline transporter-like1 (CHER1) is crucial for plasmodesmata maturation in Arabidopsis thaliana. Plant J 89:394–406 [DOI] [PubMed] [Google Scholar]
  50. Kriechbaumer V et al. (2015) Reticulomics: protein-protein interaction studies with two plasmodesmata-localized reticulon family proteins identify binding partners enriched at plasmodesmata, endoplasmic reticulum, and the plasma membrane. Plant Physiol 169:1933–1945 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Lee JY et al. (2011) A plasmodesmata-localized protein mediates crosstalk between cell-to-cell communication and innate immunity in arabidopsis. Plant Cell 23:3353–3373 [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Li X et al. (2017) Differential TOR activation and cell proliferation in Arabidopsis root and shoot apexes. Proc Natl Acad Sci USA 114:2765–2770 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Lutt N, Brunkard JO (2022) Amino acid signaling for TOR in eukaryotes: sensors, transducers, and a sustainable agricultural fuTORe. Biomol Ther 12:387–387 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Maule AJ, Gaudioso-Pedraza R, Benitez-Alfonso Y (2013) Callose deposition and symplastic connectivity are regulated prior to lateral root emergence. Commun Integr Biol 6:e26531. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Meeuse A (1941) Plasmodesmata. Bot Rev 7:249–262 [Google Scholar]
  56. Meng L, Wong JH, Feldman LJ, Lemaux PG, Buchanan BB (2010) A membrane-associated thioredoxin required for plant growth moves from cell to cell, suggestive of a role in intercellular communication. Proc Natl Acad Sci USA 107:3900–3905 [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Moubayidin L, Mambro RD, Sabatini S (2009) Cytokinin–auxin crosstalk. Trends Plant Sci 14:557–562 [DOI] [PubMed] [Google Scholar]
  58. Niklas KJ, Newman SA (2013) The origins of multicellular organisms. Evol Dev 15:41–52 [DOI] [PubMed] [Google Scholar]
  59. Okawa R, Hayashi Y, Yamashita Y, Matsubayashi Y, Ogawa-Ohnishi M (2023) Arabinogalactan protein polysaccharide chains are required for normal biogenesis of plasmodesmata. Plant J 113:493–503 [DOI] [PubMed] [Google Scholar]
  60. Ostermeyer GP, Jensen KH, Franzen AR, Peters WS, Knoblauch M (2022) Diversity of funnel plasmodesmata in angiosperms: the impact of geometry on plasmodesmal resistance. Plant J 110:707–719 [DOI] [PubMed] [Google Scholar]
  61. Paterlini A, Belevich I (2022) Serial Block Electron Microscopy to Study Plasmodesmata in the Vasculature of Arabidopsis thaliana Roots. In: Benitez-Alfonso Y, Heinlein M (eds) Plasmodesmata, vol 2457. Springer; US, New York, NY, pp 95–107 [DOI] [PubMed] [Google Scholar]
  62. Paterlini A et al. (2022) Enzymatic fingerprinting reveals specific xyloglucan and pectin signatures in the cell wall purified with primary plasmodesmata. Front Plant Sci 13:1020506. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Paultre DSG, Gustin MP, Molnar A, Oparka KJ (2016) Lost in transit: long-distance trafficking and phloem unloading of protein signals in arabidopsis homografts. Plant Cell 28:2016–2025 [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Peters WS, Jensen KH, Stone HA, Knoblauch M (2021) Plasmodesmata and the problems with size: interpreting the confusion. J Plant Physiol 257:153341. [DOI] [PubMed] [Google Scholar]
  65. Reagan BC, Ganusova EE, Fernandez JC, McCray TN, Burch-Smith TM (2018a) RNA on the move: the plasmodesmata perspective. Plant Sci 275:1–10 [DOI] [PubMed] [Google Scholar]
  66. Reagan BC, Kim PJ-Y, Perry PD, Dunlap JR, Burch-Smith TM (2018b) Spatial distribution of organelles in leaf cells and soybean root nodules revealed by focused ion beam-scanning electron microscopy. Funct Plant Biol 45:180–191 [DOI] [PubMed] [Google Scholar]
  67. Reagan BC, Dunlap JR, Burch-Smith TM (2022) Focused ion beam-scanning electron microscopy for investigating plasmodesmal densities. In: Benitez-Alfonso Y, Heinlein M (eds) Plasmodesmata, vol 2457. Springer; US, New York, NY, pp 109–123 [DOI] [PubMed] [Google Scholar]
  68. Rutschow HL, Baskin TI, Kramer EM (2011) Regulation of solute flux through plasmodesmata in the root meristem. Plant Physiol 155:1817–1826 [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Sankoh AF, Burch-Smith TM (2021) Approaches for investigating plasmodesmata and effective communication. Curr Opin Plant Biol 64:102143. [DOI] [PubMed] [Google Scholar]
  70. Schermelleh L et al. (2019) Super-resolution microscopy demystified. Nat Cell Biol 21:72–84 [DOI] [PubMed] [Google Scholar]
  71. Simpson C, Thomas C, Findlay K, Bayer E, Maule AJ (2009) An arabidopsis GPI-anchor plasmodesmal neck protein with callose binding activity and potential to regulate cell-to-cell trafficking. Plant Cell 21:581–594 [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Springmann M et al. (2018) Options for keeping the food system within environmental limits. Nature 562:519–525 [DOI] [PubMed] [Google Scholar]
  73. Stonebloom S et al. (2009) Loss of the plant DEAD-box protein ISE1 leads to defective mitochondria and increased cell-to-cell transport via plasmodesmata. Proc Natl Acad Sci USA 106:17229–17234 [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Stonebloom S et al. (2012) Redox states of plastids and mitochondria differentially regulate intercellular transport via plasmodesmata. Plant Physiol 158:190–199 [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Tee EE, Faulkner C (2024) Plasmodesmata and intercellular molecular traffic control. New Phytol:nph.19666. 10.1111/nph.19666 [DOI] [PubMed] [Google Scholar]
  76. Thomas CL, Bayer EM, Ritzenthaler C, Fernandez-Calvino L, Maule AJ (2008) Specific targeting of a plasmodesmal protein affecting cell-to-cell communication. PLoS Biol 6:0180–0190 [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Ung KL et al. (2022) Structures and mechanism of the plant PIN-FORMED auxin transporter. Nature 609:605–610 [DOI] [PMC free article] [PubMed] [Google Scholar]
  78. Wang X et al. (2013) Salicylic acid regulates plasmodesmata closure during innate immune responses in Arabidopsis. Plant Cell 25:2315–2329 [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Wang Y et al. (2023) Plasmodesmata mediate cell-to-cell transport of brassinosteroid hormones. Nat Chem Biol 19:1331–1341 [DOI] [PMC free article] [PubMed] [Google Scholar]
  80. Wegner L, Porth ML, Ehlers K (2023) Multicellularity and the need for communication—a systematic overview on (algal) Plasmodesmata and other types of symplasmic cell connections. Plan Theory 12:3342. [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Wiśniewska J et al. (2006) Polar PIN localization directs auxin flow in plants. Science 312:883–883 [DOI] [PubMed] [Google Scholar]
  82. Yan D et al. (2019) Sphingolipid biosynthesis modulates plasmodesmal ultrastructure and phloem unloading. Nat Plants 5:604–615 [DOI] [PMC free article] [PubMed] [Google Scholar]
  83. Yuan C, Lazarowitz SG, Citovsky V (2016) Identification of a functional plasmodesmal localization signal in a plant viral cell-to-cell-movement protein. MBio 7:e02052–e02015 [DOI] [PMC free article] [PubMed] [Google Scholar]
  84. Yuan C, Lazarowitz SG, Citovsky V (2018) The plasmodesmal localization signal of TMV MP is recognized by plant Synaptotagmin SYTA. MBio 9:e01314–e01318 [DOI] [PMC free article] [PubMed] [Google Scholar]
  85. Zavaliev R, Ueki S, Epel BL, Citovsky V (2011) Biology of callose (β−1,3-glucan) turnover at plasmodesmata. Protoplasma 248:117–130 [DOI] [PMC free article] [PubMed] [Google Scholar]
  86. Zavaliev R, Dong X, Epel BL (2016) Glycosylphosphatidylinositol (GPI) modification serves as a primary plasmodesmal sorting signal. Plant Physiol 172:1061–1073 [DOI] [PMC free article] [PubMed] [Google Scholar]

RESOURCES