Byung-Ho Kanga

Charles T Andersonb

Shin-Ichi Arimurac

Emmanuelle Bayerd

Magdalena Bezanillae

Miguel A Botellaf

Federica Brandizzig

Tessa M Burch-Smithj

Kent D Chapmank

Kai Dünses

Yangnan Gul

Yvon Jaillaism

Helmut Kirchhoffn

Marisa S Oteguio

Abel Rosadop

Yu Tangl

Jürgen Kleine-Vehns

Pengwei Wangq

Bethany Karlin Zolmanr

A Glossary of Plant Cell Structures: current insights and future questions

In this glossary of plant cell structures, we asked experts to summarize a present-day view of plant organelles and structures, including a discussion of outstanding questions. In the following short reviews, authors discuss the complexities of the plant cell endomembrane system, exciting connections between organelles, novel insights into peroxisome structure and function, dynamics of mitochondria, and the mysteries that need to be unlocked from the plant cell wall. These discussions are focused through a lens of new microscopy techniques. Advanced imaging has uncovered unexpected shapes, dynamics and intricate membrane formations. With a continued focus in the next decade, these imaging modalities coupled with functional studies are sure to begin to unravel mysteries of the plant cell.

However, it remains to be determined whether other PNL components, such as INM proteins, also contribute to this process.

Nuclear interior organization

Within the nucleus, the genome is organized three-dimensionally with chromosomes occupying specific territories and active and inactive chromatin regions separated from each other. Most heterochromatic regions and chromocenters are typically positioned near the nuclear periphery.

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Recent genome-wide high throughput chromosome conformation capture (Hi-C) analyses in both diploid and polyploid plant species revealed extensive inter-and intra-chromosomal interactions that define higher-order chromosomal packing during interphase (Bi et al.,

Introduction

For the Cell Biology Special Focus Issue, we wanted readers to have a modern view of plant cell structures that are sure to come up in research articles and other reviews. We gathered experts in the field to share their striking images as well as their intellectual insights on how plant cell structures are viewed today. The journey through the plant cell begins at the nucleus, where new insights into nuclear connections with the cytoplasm are beginning to decipher how the nucleus is organized and linked to cytoplasmic status. The nucleus is also the site of many phase separated structures, making the plant nucleus an ideal place to study this exciting new cell biological phenomena. Continuous with the nuclear envelope, the endoplasmic reticulum (ER) in all of its spatial and temporal complexity holds many unresolved questions. The Golgi, of central importance in the polysaccharide biosynthesis for building the plant body, has several plant-specific features. The trans Golgi network and endosomes comprise a nexus of membrane intricate compartments with vastly different shaping mechanisms ultimately linking trafficking to the plasma membrane or the vacuole. The vacuole is the largest organelle in mature plant cells, playing multiple roles from cellular homeostasis, storage, development, to response against biotic/abiotic stresses. Long known to be a reservoir for lipids, lipid droplets are emerging as important for responding to environmental stress. New insights into the molecular mechanisms driving lipid droplet formation from the ER are discussed. Recent imaging of peroxisomes in developing seedlings revealed strikingly complex membrane topologies. Imaging of live mitochondria demonstrates how dynamic plant mitochondria are with many fusion and fission events occurring to generate a syncytial mitochondria network within cells. High resolution imaging of chloroplasts during developmental transitions underscores the structural complexity of these organelles and provides new models for populating the essential thylakoid membranes. Contact sites couple organelles to each other creating a mechanism to communicate status across different subcellular structures. Plasmodesmata connect cells to each other providing long-distance communication. Finally, the cell wall not only patterns the cell but builds the plant body and protects the plant from both abiotic and biotic stressors.

Plant nucleus: a giant in the organelle galaxy

The nucleus can be viewed as a gigantic organelle and is defined by a double-layered membrane structure called the nuclear envelope (NE). The NE sequesters the nuclear genome and spatially separates transcription from translation, an evolutionary invention that enables remarkable functions and regulations in the eukaryotic cell. Here, we briefly summarize current views in key aspects of the plant nucleus, including structure, composition, dynamics, and function, from the surface to the interior.

Nuclear envelope protein composition and function

The NE is thought to have evolved from the endoplasmic reticulum (ER) and coated vesicles and is composed of the outer nuclear membrane (ONM) and the inner nuclear membrane (INM), both of which harbor distinct collections of proteins that make the NE a platform for versatile functions and communications. Plant NE proteins have been reported to function in nuclear calcium signaling [START_REF] Capoen | Nuclear membranes control symbiotic calcium signaling of legumes[END_REF][START_REF] Charpentier | Nuclear-localized cyclic nucleotide-gated channels mediate symbiotic calcium oscillations[END_REF], chromatin organization and dynamics [START_REF] Pawar | A novel family of plant nuclear envelope-associated proteins[END_REF][START_REF] Gumber | MLKS2 is an ARM domain and F-actin-associated KASH protein that functions in stomatal complex development and meiotic chromosome segregation[END_REF], immune activation [START_REF] Gu | Nuclear Pore Permeabilization Is a Convergent Signaling Event in Effector-Triggered Immunity[END_REF], cell cycle progression (Wang et al., 2019c), mechanical shielding [START_REF] Goswami | Mechanical Shielding in Plant Nuclei[END_REF], and more. Among them, the linker of nucleoskeleton and cytoskeleton (LINC) complex is one of the best characterized. The LINC complex is composed of the INM-localized Sad1/UNC48 homology (SUN) protein and the ONM-localized Klarsicht/ANC-1/Syne Homology (KASH) protein, with the former associated with the nucleoskeleton and chromatin and the latter bound with cytoskeleton and motor proteins. SUN and KASH physically interact in the perinuclear region and establish a molecular structure that enables the translation of cytoplasmic mechanical forces into the nuclear movement and chromatin activities. The plant LINC complexes have been shown to play critical roles in stomatal development and responses to light and hormone signals [START_REF] Gumber | MLKS2 is an ARM domain and F-actin-associated KASH protein that functions in stomatal complex development and meiotic chromosome segregation[END_REF]Biel et al., 2020a, b), male gametophyte development (Tamura et al., 2013;[START_REF] Varas | Absence of SUN1 and SUN2 proteins in Arabidopsis thaliana leads to a delay in meiotic progression and defects in synapsis and recombination[END_REF]Zhou et al., 2015;[START_REF] Moser | LINC-complex mediated positioning of the vegetative nucleus is involved in calcium and ROS signaling in Arabidopsis pollen tubes[END_REF], and plant-microbe interactions [START_REF] Griffis | Nuclei in motion: movement and positioning of plant nuclei in development, signaling, symbiosis, and disease[END_REF][START_REF] Newman-Griffis | Medicago LINC Complexes Function in Nuclear Morphology, Nuclear Movement, and Root Nodule Symbiosis[END_REF]. Nonetheless, compared with animals and yeast, we still lack a comprehensive understanding of the NE proteome and function in plants. Recent applications of advanced proteomic tools (e.g. proximity labeling proteomics) empowered the identification of novel NE components [START_REF] Goto | Comprehensive nuclear proteome of Arabidopsis obtained by sequential extraction[END_REF][START_REF] Huang | Proximity labeling proteomics reveals critical regulators for inner nuclear membrane protein degradation in plants[END_REF] and NE-specific biological processes in plants (e.g. INM-associated membrane protein degradation [START_REF] Huang | Proximity labeling proteomics reveals critical regulators for inner nuclear membrane protein degradation in plants[END_REF], opening brand new avenues to greatly expand our view of the global protein landscape of the plant NE and to unravel both eukaryote-conserved and plant-specific NE functions.

Nuclear pore complex, more than a conduit for nucleocytoplasmic transport

As a special membrane compartment, the nucleus evolved a sophisticated communication system that allows the remarkably efficient but highly selective exchange of materials across the NE. The ONM and the INM fuse at numerous sites to form physical openings, ~120 nm each in diameter, termed nuclear pores. The surface of individual plant nuclear pores is covered by ~1,000 nucleoporin proteins of ~40 different kinds, which are assembled into a structurally conserved mega protein complex named the nuclear pore complex (NPC) [START_REF] Ueda | Myosin-dependent endoplasmic reticulum motility and F-actin organization in plant cells[END_REF][START_REF] Mosalaganti | In situ architecture of the algal nuclear pore complex[END_REF]. The central channel of the NPC is filled with a protein meshwork made by intrinsically disordered phenylalanine-glycine (FG)-rich nucleoporins, which are capable of interacting with nuclear transport receptors (importin and exportin) and mediate selective transport of cargo molecules. Besides a conserved role in mediating nucleocytoplasmic transport, individual plant nucleoporins have been reported to play specific roles in regulating flowering time, hormone signaling, and activation of abiotic and biotic stress responses, suggesting that the NPC may function as a versatile signaling platform in addition to a conserved trafficking apparatus in plants [START_REF] Meier | Cell Biology of the Plant Nucleus[END_REF][START_REF] Gu | The nuclear pore complex: a strategic platform for regulating cell signaling[END_REF][START_REF] Li | Structural and functional insight into the nuclear pore complex and nuclear transport receptors in plant stress signaling[END_REF].

Nucleoskeleton

Underneath the INM deploys the plant nucleoskeleton assembled by long coiled-coil lamin-like proteins (e.g. CRWNs) and CRWN-associated proteins (e.g. KAKU4), which bear no sequence homology with animal lamin proteins. They are required for proper nuclear morphology [START_REF] Wang | Arabidopsis CROWDED NUCLEI (CRWN) proteins are required for nuclear size control and heterochromatin organization[END_REF][START_REF] Goto | The Novel Nuclear Envelope Protein KAKU4 Modulates Nuclear Morphology in Arabidopsis[END_REF][START_REF] Mckenna | Maize (Zea mays L.) Nucleoskeletal Proteins Regulate Nuclear Envelope Remodeling and Function in Stomatal Complex Development and Pollen Viability[END_REF] and potentially interact extensively with membranebound INM proteins to form the plant nuclear lamina (PNL). CRWNs were recently shown to interact with histone modifiers and necessary for tethering chromatin to the INM to suppress stress-related gene expression [START_REF] Hu | Plant lamin-like proteins mediate chromatin tethering at the nuclear periphery[END_REF][START_REF] Mikulski | The Chromatin-Associated Protein PWO1 Interacts with Plant Nuclear Lamin-like Components to Regulate Nuclear Size[END_REF][START_REF] Choi | The role of CRWN nuclear proteins in chromatin-based regulation of stress response genes[END_REF][START_REF] Sakamoto | Subnuclear gene positioning through lamina association affects copper tolerance[END_REF][START_REF] Wang | Characterization of a Plant Nuclear Matrix Constituent Protein in Liverwort[END_REF]. These studies suggest a critical role of the PNL in maintaining heterochromatin organization and repression at the nuclear rim, similar to what was found in animals.

Open questions on the network structure of the plant endoplasmic reticulum

The plant ER network is composed of interconnected tubules and cisternae that form a highly dynamic membrane network, which is anchored to the plasma membrane (PM), similar to a spider web hanging off surfaces (Figure 2). ER tubules connect with other tubules and cisternae forming small, triangular sheets that are called "three-way junctions" [START_REF] Shemesh | A model for the generation and interconversion of ER morphologies[END_REF]. The bulk of the plant ER is distributed at the cell cortex where it is sandwiched between the PM and the tonoplast, in continuum with the nuclear envelope and the transvacuolar strands (TVSs). TVSs form a tightly packed meshwork of ER tubules and cisternae that connect distal portions of the ER across the cell through tonoplast invaginations. The nature of the plant ER cisternae is unknown: they may be continuous membrane sheets and tightly packed tubules or perforated sheets of membranes, as described in non-plant species [START_REF] Nixon-Abell | Increased spatiotemporal resolution reveals highly dynamic dense tubular matrices in the peripheral ER[END_REF][START_REF] Schroeder | Dynamic nanoscale morphology of the ER surveyed by STED microscopy[END_REF].

The ER network undergoes continuous remodeling, through processes that include homotypic fusion of ER tubules and interconversion of ER tubule and cisternae, due to the action of ER shapers, cytoskeleton and associated motors, and ER-cytoskeleton connectors [START_REF] Brandizzi | Maintaining the structural and functional homeostasis of the plant endoplasmic reticulum[END_REF]. Together, these processes and ER shapers contribute to the overall movement or streaming of the ER, which is distinct from the movement of other organelles (e.g., peroxisomes, mitochondria, endosomes), which translocate across the cytoplasm. The relative abundance of ER tubules and cisternae varies during cell growth. As cells expand, the ER shape transitions from a more predominantly cisternal form, typical of non-expanded cells, to a more tubular form that is visible in mature cells [START_REF] Ridge | Developmental transitions and dynamics of the cortical ER of Arabidopsis cells seen with green fluorescent protein[END_REF][START_REF] Lai | ER stress signaling requires RHD3, a functionally conserved ER-shaping GTPase[END_REF], through mechanisms that are yet to be established.

In vitro and in vivo experiments have demonstrated that the ER membrane-associated GTPase Root

Hair Defective 3 (RHD3) is responsible for the homotypic fusion of the ER membrane [START_REF] Chen | Arabidopsis RHD3 mediates the generation of the tubular ER network and is required for Golgi distribution and motility in plant cells[END_REF]Stefano et al., 2012;[START_REF] Zhang | ROOT HAIR DEFECTIVE3 family of dynamin-like GTPases mediates homotypic endoplasmic reticulum fusion and is essential for Arabidopsis development[END_REF][START_REF] Ueda | Phosphorylation of the C Terminus of RHD3 Has a Critical Role in Homotypic ER Membrane Fusion in Arabidopsis[END_REF], similarly to the mammalian and yeast homologs: atlastins and Sey1p, respectively [START_REF] Mcnew | GTP-dependent membrane fusion[END_REF]; however, the mechanisms underlying the fast and dynamic interconversion of ER tubules and cisternae are yet to be discovered.

A redistribution of membrane curvature-inducing proteins, such as the conserved membrane-inducing reticulons (Tolley et al., 2008;[START_REF] Sparkes | The plant endoplasmic reticulum: a cell-wide web[END_REF] and the three-way junction-stabilizing Lunapark proteins (Lnp1 and Lnp2) [START_REF] Kriechbaumer | Arabidopsis Lunapark proteins are involved in ER cisternae formation[END_REF][START_REF] Ueda | Endoplasmic Reticulum (ER) Membrane Proteins (LUNAPARKs) are Required for Proper Configuration of the Cortical ER Network in Plant Cells[END_REF]Sun et al., 2020a), is likely responsible for the dynamic interconversion of ER forms, but the underlying regulatory mechanisms are yet largely unknown.

The biological function of the reshaping of the plant ER is still unclear. Confocal microscopy analyses have demonstrated that the ER movement increases during cell growth concomitant with an increase in the streaming of other organelles with whom the ER is in close association, such as Golgi stacks, mitochondria, peroxisomes, and endosomes. Furthermore, defects in ER network structure due to the loss of RHD3 compromise cell expansion as well as the streaming of the ER and closely associated organelles [START_REF] Lai | ER stress signaling requires RHD3, a functionally conserved ER-shaping GTPase[END_REF]Stefano et al., 2015). Therefore, the ER contributes to the dynamics and spatial organization of other organelles, possibly through ER-organelle contact sites, and this may be necessary for organelles' function. This is supported by the evidence that in an rhd3 loss-offunction mutant, the streaming of endosomes is reduced and clathrin-mediated endocytosis is compromised (Stefano et al., 2015). These results support the conceivable hypothesis that the streaming of the ER and closely associated organelles is ultimately important for cell growth, but this has yet to be experimentally demonstrated.

A double loss-of-function mutant of the two Arabidopsis Lnps shows an increased abundance of ER sheets with dense fenestration and ER conglomerates [START_REF] Kriechbaumer | Arabidopsis Lunapark proteins are involved in ER cisternae formation[END_REF][START_REF] Ueda | Endoplasmic Reticulum (ER) Membrane Proteins (LUNAPARKs) are Required for Proper Configuration of the Cortical ER Network in Plant Cells[END_REF]Sun et al., 2020a). Combined, the evidence that the localized distribution of Lnps in the ER depends on the cellular availability of their interacting protein RHD3, and that Lnps antagonize the ER shaping role of RHD3 and induce RHD3 degradation via the proteasome pathway (Sun et al., 2020a) mechanistically support a dependence of certain ER shapers onto the abundance of other ER shapers for their distribution and function in the ER. Curiously, the loss of RHD3 alone is viable and causes only limited phenotypic defects in plant growth (Stefano et al., 2012); however, the loss of RHD3 with either member of the RHD3 family of proteins, RHD3-like 1 or RHD3-like 2, is either lethal or causes reproductive defects [START_REF] Zhang | ROOT HAIR DEFECTIVE3 family of dynamin-like GTPases mediates homotypic endoplasmic reticulum fusion and is essential for Arabidopsis development[END_REF]. Conversely, the loss of both Lnps is viable, and causes only minor defects in plant growth (Sun et al., 2020a). Therefore, certain ER shapers may have a more relevant role in the life of the cell compared to others, either because associated ER shaping events are essential compared to others or because the shapers carry out other functions, in addition to ER reshaping. For example, it has been demonstrated that maize reticulons 1 and 2 function in shaping the ER but also as autophagy receptors, and are involved in degradation of the ER through the regulated process known as ER-phagy [START_REF] Zhang | Reticulon proteins modulate autophagy of the endoplasmic reticulum in maize endosperm[END_REF]. Furthermore, RHD3 has been found to interact with Ark1, an armadillo-repeat containing kinesin, which has been suggested to pull an ER tubule toward another tubule (Sun et al., 2020b). While these findings support earlier discovery that the remodeling of a subset of ER tubules depends on their sliding on pre-existing microtubules [START_REF] Hamada | Microtubules contribute to tubule elongation and anchoring of endoplasmic reticulum, resulting in high network complexity in Arabidopsis[END_REF], they also highlight additional functions of RHD3 besides a fusogenic activity of the plant ER membranes.

Future characterization of the broader roles of the plant ER shapers may provide opportunities to establish how physiologically and developmentally relevant processes are connected to ER network integrity. For example, the loss of RHD3 leads to an attenuation of signaling in the unfolded protein response [START_REF] Lai | ER stress signaling requires RHD3, a functionally conserved ER-shaping GTPase[END_REF], a conserved cytoprotective pathway that is designed to attenuate proteotoxic stress in the ER [START_REF] Pastor-Cantizano | Functional Diversification of ER Stress Responses in Arabidopsis[END_REF]. While these results support that the homeostasis of the ER network structure is critical for cell health, a challenge for the future is to establish a mechanistic framework to connect ER shape integrity with essential signaling pathways.

Despite a functional conservation of shapers, such as RHD3, reticulons and Lnps, the plant ER structure depends on plant-unique factors. For example, a minor role of microtubules in ER reshaping is consistent with a predominant role of actin in this process (Sparkes et al., 2009a); this is markedly different from the dependence of the ER network shaping on microtubules in mammalian cells [START_REF] Waterman-Storer | Endoplasmic reticulum membrane tubules are distributed by microtubules in living cells using three distinct mechanisms[END_REF]English et al., 2009). The existence of plant-unique ER-actin interactors (i.e., SYP73 and NETWORKED 3B; [START_REF] Cao | SYP73 Anchors the ER to the Actin Cytoskeleton for Maintenance of ER Integrity and Streaming in Arabidopsis[END_REF][START_REF] Wang | NETWORKED 3B: a novel protein in the actin cytoskeleton-endoplasmic reticulum interaction[END_REF], plantspecific molecular motors (i.e. Myosin XI family) [START_REF] Peremyslov | Class XI myosins are required for development, cell expansion, and F-Actin organization in Arabidopsis[END_REF][START_REF] Ueda | Myosin-dependent endoplasmic reticulum motility and F-actin organization in plant cells[END_REF], and the absence in the plant genome of CLIMP63, the connector of the mammalian ER to microtubules and a spacer of the cisternal lumen [START_REF] Klopfenstein | Subdomain-specific localization of CLIMP-63 (p63) in the endoplasmic reticulum is mediated by its luminal alpha-helical segment[END_REF][START_REF] Shibata | Mechanisms determining the morphology of the peripheral ER[END_REF], further support that plants have developed specific mechanisms of ER shaping across kingdoms. An obvious challenge for the future is to determine the nature or such mechanisms through the identification of additional players. For example, proteomics of cellular compartments or targeted proteomics based on pulldowns of ER shapers have yielded opportunities to identify proteins making up the plant ER [START_REF] Dunkley | Mapping the Arabidopsis organelle proteome[END_REF][START_REF] Kriechbaumer | Arabidopsis Lunapark proteins are involved in ER cisternae formation[END_REF], but the challenge ahead is to define a functional pipeline to identify proteins specifically involved in ER structure. Forward genetics screen based on confocal microscopy analyses of Arabidopsis seedlings expressing fluorescent markers to identify mutants with defective organization of secretory organelles (Faso et al., 2009;[START_REF] Nakano | GNOM-LIKE1/ERMO1 and SEC24a/ERMO2 are required for maintenance of endoplasmic reticulum morphology in Arabidopsis thaliana[END_REF]Takagi et al., 2013) offer a realistic opportunity to identify mutations that compromise the ER, although an innate limitation of these screens is their labor-intensive nature. Automation of this type of screens alongside the implementation of software capable to quantitatively analyze the dynamics of the ER (Pain et al., 2019) is likely to offer a platform for the fast identification of modifiers of the ER shape and dynamics.

Plant Golgi stacks -Versatile glycosylation factories on the move

The Golgi lies at the center of the secretory pathway, importing cargoes from the endoplasmic reticulum (ER), adding glycosyl groups, and exporting to the post-Golgi compartments or to the extracellular space [START_REF] Alberts | Intracellular Membrane Traffic[END_REF]. Its role as a processing trader is illustrated in its polarized stack architecture where entry (cis) and exit (trans) sides are discerned (Figure 3 A-C) (Farquhar and Palade, 1981;Moore et al., 1991). The plant Golgi also synthesizes non-cellulosic cell wall polysaccharides [START_REF] Zhang | Functional compartmentation of the Golgi apparatus of plant cells : immunocytochemical analysis of high-pressure frozen-and freeze-substituted sycamore maple suspension culture cells[END_REF][START_REF] Carpita | The Cell Wall[END_REF]. The Golgi in plant cells consists of many discrete stacks (Figure 3B) and each stack is thought to function independently [START_REF] Nebenfuhr | Mobile factories: Golgi dynamics in plant cells[END_REF]. The stacks travel in the cytosol at speeds up to several microns per second and the movement is dependent on myosin motors [START_REF] Boevink | Stacks on tracks: the plant Golgi apparatus traffics on an actin/ER network[END_REF][START_REF] Madison | Class XI Myosins Move Specific Organelles in Pollen Tubes and Are Required for Normal Fertility and Pollen Tube Growth in Arabidopsis[END_REF]. The decentralized organization contrasts with the mammalian Golgi in which stacks are stitched to form a ribbon or a complex next to the nucleus [START_REF] Marsh | Organellar relationships in the Golgi region of the pancreatic beta cell line, HIT-T15, visualized by high resolution electron tomography[END_REF]. Therefore, ER-to-Golgi transport and post-Golgi secretion requires long-distance vesicular trafficking to and from the Golgi (Gillingham and Munro, 2016). Mobile Golgi stacks in plants, instead, can visit ER export sites (ERESs), concentrate to sites of secretion, and redistribute for cell division [START_REF] Nebenfuhr | Redistribution of Golgi stacks and other organelles during mitosis and cytokinesis in plant cells[END_REF][START_REF] Yang | Dynamics of COPII Vesicles and the Golgi Apparatus in Cultured Nicotiana tabacum BY-2 Cells Provides Evidence for Transient Association of Golgi Stacks with Endoplasmic Reticulum Exit Sites[END_REF][START_REF] Ndinyanka Fabrice | Efficient preparation of Arabidopsis pollen tubes for ultrastructural analysis using chemical and cryo-fixation[END_REF].

Transport through the plant Golgi

Golgi stacks move slowly when they become associated with ERES and the ER-to-Golgi transport is mediated by COPII-type vesicles [START_REF] Nebenfuhr | Stop-and-go movements of plant Golgi stacks are mediated by the acto-myosin system[END_REF] ] [START_REF] Yang | Dynamics of COPII Vesicles and the Golgi Apparatus in Cultured Nicotiana tabacum BY-2 Cells Provides Evidence for Transient Association of Golgi Stacks with Endoplasmic Reticulum Exit Sites[END_REF] [START_REF] Kang | ER-to-Golgi transport by COPII vesicles in Arabidopsis involves a ribosome-excluding scaffold that is transferred with the vesicles to the Golgi matrix[END_REF]. In mammalian cells, the ER-to-Golgi intermediate compartment (ERGIC) assembles at ERES and ER-resident proteins are retrieved from ERGIC before they reach the perinuclear Golgi complex [START_REF] Appenzeller-Herzog | The ER-Golgi intermediate compartment (ERGIC): in search of its identity and function[END_REF]. The plant Golgi lacks ERGIC compartments as COPII vesicles are directly transferred to the cis-Golgi. It was demonstrated that biosynthetic enzymes are localized to the medial Golgi after recycling of ER proteins is finished in the cis-Golgi [START_REF] Donohoe | Cis-Golgi cisternal assembly and biosynthetic activation occur sequentially in plants and algae[END_REF], suggesting that the cis cisternae functionally replaces ERGIC in plants [START_REF] Ito | Differentiation of Trafficking Pathways at Golgi Entry Core Compartments and Post-Golgi Subdomains[END_REF].

Among the models explaining the intra-Golgi transport, the cisternal progression/maturation model has been supported by transmission electron microscopy (TEM) studies of the plant Golgi [START_REF] Robinson | Plant Golgi ultrastructure[END_REF]. It is evident from electron micrographs of plant Golgi stacks that Golgi cisternae are peeled off from the trans-side, indicating that Golgi cisternae are transient entities, disposed of from the stack once they reach the trans-end [START_REF] Day | A three-stage model of Golgi structure and function[END_REF]. Electron tomography (ET) analysis has shown assembly intermediates of new cisternae on the cis-side that exhibit highly varying sizes and shapes [START_REF] Donohoe | Cis-Golgi cisternal assembly and biosynthetic activation occur sequentially in plants and algae[END_REF]. Cell wall polysaccharides were detected in the cisternal lumen but not in COPI-type vesicles at the cisternal margins that are thought to retain Golgi resident proteins against the cisternal membrane flux [START_REF] Donohoe | Identification and characterization of COPIa-and COPIb-type vesicle classes associated with plant and algal Golgi[END_REF]. On the trans-side, trans-Golgi network (TGN) compartments arise from the trans-most cisternae. The transformation involves a significant reduction in the membrane amounts, suggesting that Golgi resident proteins are retrieved from the TGN [START_REF] Kang | Shrinkage and fragmentation of the trans-Golgi network in non-meristematic plant cells[END_REF][START_REF] Kang | Electron Tomography of RabA4b-and PI-4Kβ1-Labeled Trans Golgi Network Compartments in Arabidopsis[END_REF].

The aforementioned transports occur within a ribosome-excluding matrix that encloses COPII vesicles to TGN cisternae (Figure 3D) [START_REF] Staehelin | Nanoscale architecture of endoplasmic reticulum export sites and of Golgi membranes as determined by electron tomography[END_REF]. They probably correspond to a dense network of proteins involved in Golgi membrane assembly, maturation, TGN formation, and fastening cisternae into a stack. Golgins are Golgi-localized long coiled-coil proteins and some of them are tethering factors [START_REF] Latijnhouwers | Holding it all together? Candidate proteins for the plant Golgi matrix[END_REF] and; given their rod-like shape, they constitute scaffolds for the matrix. Mammalian Golgins are required for the Golgi integrity, vesicular trafficking to the Golgi, and protein glycosylation [START_REF] Wong | Membrane trafficking. The specificity of vesicle traffic to the Golgi is encoded in the golgin coiled-coil proteins[END_REF], [START_REF] Liu | Loss of the golgin GM130 causes Golgi disruption, Purkinje neuron loss, and ataxia in mice[END_REF] [START_REF] Witkos | GORAB scaffolds COPI at the trans-Golgi for efficient enzyme recycling and correct protein glycosylation[END_REF].

Arabidopsis Golgins have been shown to play roles in COPII vesicular transport [START_REF] Kang | ER-to-Golgi transport by COPII vesicles in Arabidopsis involves a ribosome-excluding scaffold that is transferred with the vesicles to the Golgi matrix[END_REF] and interaction of the cis-Golgi with ERES [START_REF] Osterrieder | Stacks off tracks: a role for the golgin AtCASP in plant endoplasmic reticulum-Golgi apparatus tethering[END_REF].

Biosynthesis in the plant Golgi

Production and export of cell wall matrix polysaccharides distinguish the plant Golgi from its animal counterpart. It has been shown that the reaction cascades for polysaccharide synthesis are arranged sequentially over the stack from the cis-to-trans direction. As amounts of glycosyltransferases and sugar transporters are small per stack, their localization within the Golgi has been investigated with overexpressor lines or by localizing reaction products [START_REF] Chevalier | Subcompartment localization of the side chain xyloglucan-synthesizing enzymes within Golgi stacks of tobacco suspension-cultured cells[END_REF][START_REF] Meents | Organization of Xylan Production in the Golgi During Secondary Cell Wall Biosynthesis[END_REF].

Cell wall polysaccharides secreted constitutively and several mechanisms for retaining Golgi proteins from the bulk flow have been characterized [START_REF] Brandizzi | The Destination for Single-Pass Membrane Proteins Is Influenced Markedly by the Length of the Hydrophobic Domain[END_REF][START_REF] Schoberer | A signal motif retains Arabidopsis ER-alpha-mannosidase I in the cis-Golgi and prevents enhanced glycoprotein ERAD[END_REF].

TGN cisternae consist of distinct domains where secretory and vacuolar cargoes are separately packaged [START_REF] Shimizu | Cargo sorting zones in the trans-Golgi network visualized by super-resolution confocal live imaging microscopy in plants[END_REF]. Electron tomography imaging of Golgi/TGN complexes revealed that varying ratios of different vesicle buds in a TGN cisterna, suggesting that biosynthetic functions of each Golgi stack are not uniform in a plant cell [START_REF] Staehelin | Nanoscale architecture of endoplasmic reticulum export sites and of Golgi membranes as determined by electron tomography[END_REF]. Golgi stacks appear to be versatile factories of which activities are determined by the protein that they received from the ERES.

Golgi stacks enriched with enzymes for synthesizing cell wall polysaccharides would give rise to more secretory vesicles than those that received proteins destined for the vacuole.

Future research perspectives

The structure of plant Golgi varies in different cell types (Staehelin et al., 1990) but the molecular mechanisms governing the remodeling remain elusive. For example, small Golgi stacks in root meristem cells exhibit structural-functional modifications in parallel with the differentiation of the meristem cells into columella and eventually into peripheral/border cells in the root cap (Wang et al., 2017c). Since several cell-specific markers for the Arabidopsis root cap have been identified [START_REF] Kamiya | Control of root cap maturation and cell detachment by BEARSKIN transcription factors in Arabidopsis[END_REF], it would be possible to identify genes involved in Golgi-mediated secretion or glycosylation in columella or peripheral/border cells after single-cell sequencing of root cap isolates [START_REF] Shaw | Single-Cell Transcriptome Analysis in Plants: Advances and Challenges[END_REF].

Correlative light and electron microscopy refers to protocols in which macromolecules are first localized with fluorescence microscopy and the volume enclosing the macromolecules is imaged with EM. The correlative approach will be useful for analyzing organelles composed of heterogeneous members (Wang et al., 2019a). After classifying Golgi stacks associated with specific markers, each group could be investigated for characterizing its nanoscale architectures and interaction with other organelles. The dynamics of Golgi subpopulations under stress conditions will help understanding how the secretory pathway reorganizes in response to threats from outside. As the export from the Golgi is mediated by the trans-Golgi network, this study should be combined with TGN dynamics [START_REF] Uemura | A Golgi-Released Subpopulation of the Trans-Golgi Network Mediates Protein Secretion in Arabidopsis[END_REF].

Advances in cryo-electron microscopy and sample processing technology allowed for ET analysis of frozen-hydrated cells to visualize macromolecular complexes in situ [START_REF] Otegui | Electron tomography in plant cell biology[END_REF].

Due to the limitation in section thickness for EM, frozen cells need to be either sliced or thinned with focused ion beam milling (FIB). Golgi vesicles and intraluminal filaments were delineated in Chlamydomonas cells by cryo-ET [START_REF] Engel | In situ structural analysis of Golgi intracisternal protein arrays[END_REF]. Although intact plant tissues are too thick for FIB, in vitro germinated pollen tube tips are amenable to FIB thinning [START_REF] Liu | Membrane imaging in the plant endomembrane system[END_REF]. As Golgi stacks produce numerous secretory vesicles to sustain pollen tube tip growth, it would be exciting to examine plant Golgi stacks with cryo-ET to uncover novel features not observed in plastic-embedded electron microscopy samples.

Acknowledgements: This work was supported by grants from the Research Grants Council of Hong Kong (GRF14121019, GRF 14113921, C4012-16E, C4002-17G, and AoE/ M-05/12) to B-HK. Special thanks to Drs. Pengfei Wang, Zizhen Liang, and Peng Gao (Chinese University of Hong Kong) for their help in preparing electron microscopy image panels. We apologize to colleagues for not citing their excellent work on the plant Golgi, due to the limitation in article lengths.

Plant Endosomes: The protein sorting masters.

The ability to regulate the composition of the plasma membrane and the endomembrane system is critical for cell survival. Endosomes have a central role in this process by regulating protein and lipid (cargo) trafficking in the endomembrane system, both through the anterograde and retrograde pathways. As part of the anterograde pathways, that is, transport from the site of synthesis to the place of residence and function, proteins and lipids synthesized in the endoplasmic reticulum (ER) are typically transported in vesicles to the Golgi, to the Trans-Golgi Network (TGN), and from there, either to the plasma membrane (exocytosis) or to the vacuole. Retrograde pathways mediate the transport of cargo or trafficking factors in the opposite direction to the anterograde pathway, usually back to their original donor compartments. Proteins removed from the plasma membrane in vesicles through endocytosis are delivered to early endosomes, where they can be either recycled to the plasma membrane or carried to multivesicular endosomes (MVEs) for further sorting into intralumenal vesicles and subsequent degradation in the vacuolar lumen [START_REF] Valencia | Endocytosis and Endosomal Trafficking in Plants[END_REF]. In plants, the TGN functions as the early endosome since it is the first compartment that receives endocytosed cargo [START_REF] Dettmer | Vacuolar H+-ATPase activity is required for endocytic and secretory trafficking in Arabidopsis[END_REF][START_REF] Lam | Rice SCAMP1 defines clathrin-coated, trans-golgilocated tubular-vesicular structures as an early endosome in tobacco BY-2 cells[END_REF]. Thus, in contrast to animal cells, plant cells do not have separate early endosomes but combine both endocytic and biosynthetic sorting at the TGN (Viotti et al., 2010).

TGNs and MVEs, the two types of plant endosomes, arise, mature, and are consumed as part of their membrane trafficking function. Therefore, both types of organelles are in continuous flux and can be found as subpopulations at different maturation stages.

The TGN

As part of the endosomal pathway, the TGN receives plasma membrane cargo, which is either recycled back to the plasma membrane or retained for further sorting in MVEs and degradation in vacuoles. As part of the secretory pathway, the TGN produces both secretory vesicles carrying protein, membrane lipids, and cell wall polysaccharides to the plasma membrane and vesicles containing vacuolar cargo [START_REF] Rosquete | The Plant Trans-Golgi Network: Not Just a Matter of Distinction[END_REF] and mediates retrograde recycling back to the Golgi and ER through COPI- [START_REF] Bykov | The structure of the COPI coat determined within the cell[END_REF] and retromer-mediated traffic [START_REF] Niemes | Retromer recycles vacuolar sorting receptors from the trans-Golgi network[END_REF].

The TGN forms largely through cisternal maturation of the trans-most Golgi cisterna (Golgi-associated TGN or GA-TGN) but eventually detaches from the Golgi becoming an independent organelle (free or also called Golgi-independent TGN or GI-TGN) that fragments into vesicles (Toyooka et al., 2009;[START_REF] Kang | Electron Tomography of RabA4b-and PI-4Kβ1-Labeled Trans Golgi Network Compartments in Arabidopsis[END_REF][START_REF] Uemura | Dynamic behavior of the trans-golgi network in root tissues of Arabidopsis revealed by super-resolution live imaging[END_REF][START_REF] Uemura | A Golgi-Released Subpopulation of the Trans-Golgi Network Mediates Protein Secretion in Arabidopsis[END_REF] (Figure 4). There are approximately 35 Golgi stacks and GA-TGNs in an interphase Arabidopsis meristematic cell (Segui-Simarro and [START_REF] Segui-Simarro | Cell cycle-dependent changes in Golgi stacks, vacuoles, clathrin-coated vesicles and multivesicular bodies in meristematic cells of Arabidopsis thaliana: a quantitative and spatial analysis[END_REF]. In Arabidopsis root cells, as the trans-most cisterna matures into the TGN, it develops numerous vesicle buds, loses 30-35% of its total membrane surface area, and becomes enriched in the Rab GTPases RAB-A2a and RAB-A4b, the phosphatidylinositol 4-kinase PI4Kb1, the vacuolar V-ATPase subunit VHA1a, and the SNAREs (Soluble N-ethylmaleimide sensitive factor Attachment protein Receptor) SYP61, SYP43 , VAMP721, VAMP722, and VAMP727 [START_REF] Dettmer | Vacuolar H+-ATPase activity is required for endocytic and secretory trafficking in Arabidopsis[END_REF][START_REF] Chow | Rab-A2 and Rab-A3 GTPases define a trans-golgi endosomal membrane domain in Arabidopsis that contributes substantially to the cell plate[END_REF][START_REF] Kang | Electron Tomography of RabA4b-and PI-4Kβ1-Labeled Trans Golgi Network Compartments in Arabidopsis[END_REF][START_REF] Zhang | Arabidopsis R-SNARE proteins VAMP721 and VAMP722 are required for cell plate formation[END_REF]. As the GA-TGN detaches from the Golgi stacks to become free/GI-TGNs, the budding profiles become more abundant.

These GA-and GI-TGN subpopulations play distinct trafficking functions [START_REF] Renna | TGNap1 is required for microtubule-dependent homeostasis of a subpopulation of the plant trans-Golgi network[END_REF][START_REF] Uemura | A Golgi-Released Subpopulation of the Trans-Golgi Network Mediates Protein Secretion in Arabidopsis[END_REF][START_REF] Ito | Differentiation of Trafficking Pathways at Golgi Entry Core Compartments and Post-Golgi Subdomains[END_REF]. For example, GA-TGN but not free/GI-TGNs label with the endocytic tracer FM4-64 [START_REF] Uemura | A Golgi-Released Subpopulation of the Trans-Golgi Network Mediates Protein Secretion in Arabidopsis[END_REF], suggesting that endosomal function is carried out by the GA-TGN whereas free GI/TGNs seem to be primarily involved in exocytosis. The different trafficking functions of the TGN are spatially separated in subdomains that differ both in their protein and membrane lipid composition [START_REF] Wattelet-Boyer | Enrichment of hydroxylated C24-and C26-acyl-chain sphingolipids mediates PIN2 apical sorting at trans-Golgi network subdomains[END_REF]. Thus, within the GA-TGN, there are at least two "zones", the secretory-trafficking zone that generates exocytic vesicles and is enriched in the SNARE VAMP721, the adaptor complex AP-1, the accessory protein EPSIN1, and clathrin and the vacuolar trafficking zone enriched in VAMP727, the adaptor complex AP-4, and accessory protein MODIFIED TRANSPORT TO THE VACUOLE1 (MTV1) [START_REF] Heinze | EPSIN1 and MTV1 define functionally overlapping but molecularly distinct trans-Golgi network subdomains in Arabidopsis[END_REF][START_REF] Shimizu | Cargo sorting zones in the trans-Golgi network visualized by super-resolution confocal live imaging microscopy in plants[END_REF]. In addition, a plant-specific TRAPPII complex has been hypothesized to mediate the recruitment/tethering of endocytosed vesicles to subdomains of the TGN [START_REF] Rosquete | AtTRAPPC11/ROG2: A Role for TRAPPs in Maintenance of the Plant Trans-Golgi Network/Early Endosome Organization and Function[END_REF].

The TGN not only has subdomains for exocytic, endocytic, and vacuolar trafficking, but also it associates with protein complexes that control the trafficking of specific cargo proteins. Thus, for example, the TGN-localized protein ECHIDNA controls the secretion of only a subset of plasma membrane proteins, such as the auxin influx carrier AUX1 [START_REF] Boutte | ECHIDNA-mediated post-Golgi trafficking of auxin carriers for differential cell elongation[END_REF]. In contrast, a module formed by seven transmembrane domain-containing proteins (7TM) and components of the guanine nucleotide-binding (G) protein signaling work together at the Golgi and TGN to regulate exocytosis of cellulose synthases, but not endocytosis or general exocytosis of both soluble and plasma membrane cargo [START_REF] Mcfarlane | A G protein-coupled receptor-like module regulates cellulose synthase secretion from the endomembrane system in Arabidopsis[END_REF].

MVEs

MVEs arise from membranes derived from the TGN and are characterized by a rounded shape, the presence of intralumenal vesicles, and their association with RAB-F GTPases such as ARA6, ARA7, and RHA1 [START_REF] Haas | The Arabidopsis AAA ATPase SKD1 is involved in multivesicular endosome function and interacts with its positive regulator LYST-INTERACTING PROTEIN5[END_REF]. There are approximately 17-20 MVEs in interphase meristematic cells and are usually found in close proximity to the GA-TGN (Segui-Simarro and [START_REF] Segui-Simarro | Cell cycle-dependent changes in Golgi stacks, vacuoles, clathrin-coated vesicles and multivesicular bodies in meristematic cells of Arabidopsis thaliana: a quantitative and spatial analysis[END_REF]. Plasma membrane proteins targeted for degradation are usually ubiquitinated at the plasma membrane, internalized by endocytosis, and delivered first to the TGN and then to MVEs. At the MVE limiting membrane, the ESCRT (Endosomal Sorting Complex Required for Transport) machinery binds, clusters, and sorts the ubiquitinated cargo into membrane domains that bend away from the cytoplasm, that is membrane bending in the reverse (negative) topology of the better understood cases of vesiculation, such as clathrin-mediated endocytosis. Failure to properly sort plasma membrane components into intralumenal vesicles results in the accumulation of plasma membrane proteins in the vacuolar membrane (Figure 4), which leads to severe alterations in development and most frequently, to lethality. Although it has long been assumed that ESCRTs orchestrate the formation and release of one single endosomal vesicle at a time, studies performed in Arabidopsis thaliana have shown that at least in plants, these vesicles do not bud off individually but form in concatenated networks [START_REF] Buono | ESCRT-mediated vesicle concatenation in plant endosomes[END_REF][START_REF] Goodman | ESCRT components ISTL1 andLIP5 are required for tapetal function and pollen viability[END_REF].

In general, ESCRT proteins are well conserved across organisms, from Archaea [START_REF] Makarova | Evolution of diverse cell division and vesicle formation systems in Archaea[END_REF][START_REF] Dobro | Electron cryotomography of ESCRT assemblies and dividing Sulfolobus cells suggests that spiraling filaments are involved in membrane scission[END_REF][START_REF] Pulschen | Live Imaging of a Hyperthermophilic Archaeon Reveals Distinct Roles for Two ESCRT-III Homologs in Ensuring a Robust and Symmetric Division[END_REF] to Eukarya. In fungi and metazoans, five multimeric ESCRT complexes have been identified: ESCRT-0 to III and the triple AAA ATPase SKD1 (SUPPRESSOR OF K + TRANSPORT GROWTH DEFECT 1) with its activator LIP5. Plants contain putative orthologs for most of the ESCRT proteins originally identified in metazoans and fungi [START_REF] Spitzer | The Arabidopsis elch mutant reveals functions of an ESCRT component in cytokinesis[END_REF][START_REF] Haas | The Arabidopsis AAA ATPase SKD1 is involved in multivesicular endosome function and interacts with its positive regulator LYST-INTERACTING PROTEIN5[END_REF][START_REF] Spitzer | The ESCRT-related CHMP1A and B proteins mediate multivesicular body sorting of auxin carriers in Arabidopsis and are required for plant development[END_REF][START_REF] Kalinowska | Arabidopsis ALIX is required for the endosomal localization of the deubiquitinating enzyme AMSH3[END_REF][START_REF] Buono | Role of SKD1 Regulators LIP5 and IST1-LIKE1 in Endosomal Sorting and Plant Development[END_REF][START_REF] Yu | ESCRT-I Component VPS23A Affects ABA Signaling by Recognizing ABA Receptors for Endosomal Degradation[END_REF]Wang et al., 2017a) with the exception of ESCRT-0 (Winter and Hauser, 2006), which is an early acting complex that binds phophoinositide-3-phosphate (PI3P), a lipid enriched in endosomal membranes and critical for recruitment of ESCRT proteins to endosomes. However, a group of proteins called TOL (TOML1-LIKE) are likely to play the role of ESCRT-0 in plants [START_REF] Korbei | Arabidopsis TOL Proteins Act as Gatekeepers for Vacuolar Sorting of PIN2 Plasma Membrane Protein[END_REF][START_REF] Moulinier-Anzola | TOLs Function as Ubiquitin Receptors in the Early Steps of the ESCRT Pathway in Higher Plants[END_REF].

How do ESCRT proteins mediate intralumenal vesicle formation and sequestration of cargo proteins?

ESCRT-0, -I, and -II contain ubiquitin-binding domains and contribute to the clustering of ubiquitinated cargo on the endosomal membrane and to membrane deformation [START_REF] Liese | Protein crowding mediates membrane remodeling in upstream ESCRT-induced formation of intraluminal vesicles[END_REF]. Deubiquitinating enzymes remove the ubiquitin on cargo before their final sequestration into intralumenal vesicles. Critical for the final steps in vesicle formation are the presence of membrane cargo [START_REF] Chiaruttini | Relaxation of Loaded ESCRT-III Spiral Springs Drives Membrane Deformation[END_REF] as well as ESCRT-III and ESCRT-III-associated proteins, which are able to trigger membrane deformation and neck constriction [START_REF] Hanson | Plasma membrane deformation by circular arrays of ESCRT-III protein filaments[END_REF][START_REF] Fyfe | Association of the endosomal sorting complex ESCRT-II with the Vps20 subunit of ESCRT-III generates a curvature-sensitive complex capable of nucleating ESCRT-III filaments[END_REF][START_REF] Mccullough | Membrane fission reactions of the mammalian ESCRT pathway[END_REF][START_REF] Chiaruttini | Relaxation of Loaded ESCRT-III Spiral Springs Drives Membrane Deformation[END_REF].

Plants commonly contain several isoforms for each ESCRT subunits and even have evolved plantspecific ESCRT proteins, such as PROS (Positive Regulator of SKD1) that enhances SKD1 activity [START_REF] Reyes | A novel endosomal sorting complex required for transport (ESCRT) component in Arabidopsis thaliana controls cell expansion and development[END_REF], FREE1/FYVE1 [START_REF] Gao | A unique plant ESCRT component, FREE1, regulates multivesicular body protein sorting and plant growth[END_REF], and FYVE4 (Liu et al., 2021a). Interestingly, both proteins contain FYVE domains able to bind PI3P. FREE1 interacts with ESCRT-I subunits and is essential for endosomal sorting [START_REF] Gao | A unique plant ESCRT component, FREE1, regulates multivesicular body protein sorting and plant growth[END_REF] whereas FYVE4 is necessary for the recruitment of ESCRT-III subunits (Liu et al., 2021a).

Future perspectives

Our understanding of endosomal biogenesis and the molecular machinery mediating its multiple sorting functions have increased dramatically during the past decades. However, new regulatory and sorting components are being discovered and many more remain elusive, making it still challenging to comprehend how sorting functions are both segregated and integrated in TGNs and MVEs. The plant endosomes have many distinct features that make them different from their counterparts in other organisms. For example, whether the concatenation of MVE intralumenal vesicles in complex networks is unique to plants or is a universal mechanism in all eukaryotes is presently unknown. It is tempting to speculate that the evolution of unique ESCRT components and the drastic diversification of some ESCRT isoforms may have contributed to the unique features of intralumenal vesicle formation in plants.

Lipid Droplets: Specialized Subcellular Hydrophobic Compartments

Like all cells, plant cells accumulate storage lipids in their cytoplasm as discrete lipid droplets (LDs), most often consisting of a hydrophobic core of non-bilayer forming lipids such as triacylglycerols (TAGs) or sterol esters surrounded by an emulsifying monolayer of phospholipids (Pyc et al., 2017a;[START_REF] Huang | Plant Lipid Droplets and Their Associated Proteins: Potential for Rapid Advances[END_REF][START_REF] Ischebeck | Lipid droplets in plants and algae: Distribution, formation, turnover and function[END_REF]. Although less commonly considered, rubber particles of rubberproducing plant species with a polyisoprenoid hydrophobic core, share the same overall structure [START_REF] Yamashita | Molecular Mechanisms of Natural Rubber Biosynthesis[END_REF]. This thermodynamically stable structure was originally observed in TEM micrographs, and described by various terms as lipid bodies, oil bodies, spherosomes or oleosomes [START_REF] Wanner | Membranous appendices of spherosomes (oleosomes) : Possible role in fat utilization in germinating oil seeds[END_REF]. However, the contemporary, unifying terminology of "lipid droplets" emphasizes the evolutionary conservation of this compartment across kingdoms of life where there are increasing reports of functions beyond the efficient storage of carbon [START_REF] Lundquist | Lipid droplets throughout the evolutionary tree[END_REF].

In plants, LDs are most commonly associated with oilseeds and oleaginous fruits where they compartmentalize the well-known "vegetable oils" [START_REF] Chapman | Biogenesis and functions of lipid droplets in plants: Thematic Review Series: Lipid Droplet Synthesis and Metabolism: from Yeast to Man[END_REF]. However LDs are present in essentially all cell types in plants. Because the most abundant LD proteins in seeds-oleosins-are not produced in most plant cell types, recent efforts to identify LD proteins in non-seed tissues [START_REF] Horn | Identification of a new class of lipid droplet-associated proteins in plants[END_REF][START_REF] Brocard | Proteomic Analysis of Lipid Droplets from Arabidopsis Aging Leaves Brings New Insight into Their Biogenesis and Functions[END_REF][START_REF] Kretzschmar | PUX10 Is a Lipid Droplet-Localized Scaffold Protein That Interacts with CELL DIVISION CYCLE48 and Is Involved in the Degradation of Lipid Droplet Proteins[END_REF][START_REF] Fernandez-Santos | Protein Profiles of Lipid Droplets during the Hypersensitive Defense Response of Arabidopsis against Pseudomonas Infection[END_REF] have expanded the inventory of LD proteins in plant cells. These new proteins and their partners have begun to suggest previously unrecognized participants in LD formation, stability, turnover and functions.

Among the recently-recognized LD proteins are the so-called LDAPs (LIPID DROPLET-ASSOCIATED PROTEINs), which share homology with small rubber particle proteins from rubber producing plants. LDAPs were identified as prominent proteins in purified LDs isolated from avocado mesocarp [START_REF] Horn | Identification of a new class of lipid droplet-associated proteins in plants[END_REF], and have since become appreciated for their widespread occurrence throughout the plant kingdom [START_REF] Gidda | Lipid Droplet-Associated Proteins (LDAPs) Are Required for the Dynamic Regulation of Neutral Lipid Compartmentation in Plant Cells[END_REF][START_REF] Brocard | Proteomic Analysis of Lipid Droplets from Arabidopsis Aging Leaves Brings New Insight into Their Biogenesis and Functions[END_REF][START_REF] De Vries | Ties between Stress and Lipid Droplets Pre-date Seeds[END_REF] as well as for their induction by drought stress [START_REF] Kim | Arabidopsis Small Rubber Particle Protein Homolog SRPs Play Dual Roles as Positive Factors for Tissue Growth and Development and in Drought Stress Responses[END_REF]. The LDAPs are relatively small proteins without extended hydrophobic regions, and they have been shown to localize specifically to the LD surface perhaps through their extensive amphipathic helices. Screens for potential protein interactors, that might serve as protein recognition sites for LDAPs on the organelle surface, identified the protein LDIP (LDAP-INTERACTING PROTEIN), which also is widely-distributed in the plant kingdom (Pyc et al., 2017b;Coulon et al., 2020) (de Vries andIschebeck, 2020). LDAPs and LDIP are expressed in both seed and non-seed tissues of plants, and are suspected in playing broader roles in compartmentalization of neutral lipids in cells beyond those found in seed tissues.

Another recently-identified LD protein is the PLANT UBX DOMAIN-CONTAINING PROTEIN10 (PUX10). PUX10 localizes to LDs through a hydrophobic polypeptide sequence and recruits an AAAtype ATPase CELL DIVISION CYCLE48 (CDC48) protein to the LD surface [START_REF] Deruyffelaere | PUX10 Is a CDC48A Adaptor Protein That Regulates the Extraction of Ubiquitinated Oleosins from Seed Lipid Droplets in Arabidopsis[END_REF][START_REF] Kretzschmar | PUX10 Is a Lipid Droplet-Localized Scaffold Protein That Interacts with CELL DIVISION CYCLE48 and Is Involved in the Degradation of Lipid Droplet Proteins[END_REF]. This interaction is believed to support the selective extraction of LD surface proteins, like oleosins and LDAPs, for their ubiquitin-mediated protein degradation. This LDassociated degradation pathway likely operates in all cells of plants to repurpose the surface and/or contents of the LD compartment during development or in response to environmental stresses.

LD formation at the ER

Like in most eukaryotes, LD formation in plant cells originates in the ER where the enzymes for storage lipid assembly are present. Ultrastructural studies frequently reveal intimate connections of LDs with the ER [START_REF] Herman | Seed Oil Body Ontogeny[END_REF][START_REF] Brocard | Proteomic Analysis of Lipid Droplets from Arabidopsis Aging Leaves Brings New Insight into Their Biogenesis and Functions[END_REF] which can also be captured by confocal fluorescence laser scanning microscopy (Figure 6A). The process of LD proliferation can be capitulated at the subcellular level in Nicotiana benthamiana cells, where LDs are normally low in abundance, and transient expression of cDNAs encoding proteins suspected in LD formation can be readily studied (Figure. 6). A transient system for LD studies also has been developed with tobacco pollen tubes [START_REF] Muller | Tobacco pollen tubes -a fast and easy tool for studying lipid droplet association of plant proteins[END_REF], and this has been particularly useful for protein localization studies due to the large number of LDs normally present in these cells.

Existing models for LD formation suggest that newly-synthesized TAGs aggregate and form foci or "lipid lens" structures between the two leaflets of the ER bilayer (Figure 6B) (Pyc et al., 2017a). An oligomeric protein complex comprised of SEIPIN subunits in the ER bilayer coordinates these TAG foci as they grow [START_REF] Chapman | Mechanisms of lipid droplet biogenesis[END_REF]. SEIPIN proteins direct a bulge of newly accumulating neutral lipids to emerge into the cytoplasm covered with a monolayer of ER-derived phospholipid. Unlike fungi and metazoans, plants have multiple genes encoding SEIPIN isoforms [START_REF] Cai | Arabidopsis SEIPIN Proteins Modulate Triacylglycerol Accumulation and Influence Lipid Droplet Proliferation[END_REF]. In Arabidopsis SEIPIN 1 is expressed mostly in seed and seedling tissues, whereas SEIPIN 2 and SEIPIN 3 are expressed in essentially all issues. While loss-of-function mutants in a single SEIPIN gene in Arabidopsis resulted in negligible phenotypes, double and especially triple seipin mutants showed dramatic cellular disruptions in normal LD formation (Taurino et al., 2018). Seeds and pollen of sei1/sei2/sei3 mutants accumulated large aberrant-shaped LDs, sometimes observable in the nucleus and ER lumen in addition to the cytoplasm. These results indicate that SEIPINs play a critical and partially redundant role in the normal formation of LDs in plant cells. Structural models based on homology with known structures of Drosophila and human SEIPIN, suggests that the three Arabidopsis SEIPINS can form homo-oligomeric structures with different numbers of subunits [START_REF] Chapman | Mechanisms of lipid droplet biogenesis[END_REF], but future work is required to understand the functional interactions of the three SEIPIN proteins in plant cells, their potential for hetero-oligomeric interactions and their partners in LD biogenesis.

The loss-of-function of two other Arabidopsis genes revealed similar, large and aberrant LD phenotypes in seeds, reminiscent of seipin mutants. These two genes encode the VESICLE-ASSOCIATED MEMBRANE PROTEIN-ASSOCIATED PROTEIN 27-1 (VAP27-1) and LDIP proteins, respectively, which were shown to interact with SEIPINs and with LDs (Pyc et al., 2017b;[START_REF] Greer | SEIPIN Isoforms Interact with the Membrane-Tethering Protein VAP27-1 for Lipid Droplet Formation[END_REF]. In other work, higher order oleosin mutants also displayed aberrant formation of LDs during early seed development, and this was reported to result from alteration of fusion dynamics of very small LDs, not necessarily during LD formation at the ER [START_REF] Miquel | Specialization of oleosins in oil body dynamics during seed development in Arabidopsis seeds[END_REF]. LDAPs also occur in seeds but at lower amounts than oleosins [START_REF] Kretzschmar | PUX10 Is a Lipid Droplet-Localized Scaffold Protein That Interacts with CELL DIVISION CYCLE48 and Is Involved in the Degradation of Lipid Droplet Proteins[END_REF]. And while LDAPs are interactors of LDIP (Pyc et al., 2017b), their loss-of-function in ldap mutants did not result in dramatic alterations of LD morphology in seeds [START_REF] Gidda | Lipid Droplet-Associated Proteins (LDAPs) Are Required for the Dynamic Regulation of Neutral Lipid Compartmentation in Plant Cells[END_REF], although there may be some increase in LD size in leaves of ldap knockdowns [START_REF] Brocard | Proteomic Analysis of Lipid Droplets from Arabidopsis Aging Leaves Brings New Insight into Their Biogenesis and Functions[END_REF]. Future work will be required to piece together the mechanistic associations between SEIPINs, LDIP, VAP27-1, oleosins, LDAPs and other LD proteins; nevertheless, results to date support cooperative roles for these proteins in the cellular process of LD formation in plant cells.

Engineering the LD compartment

Because of their high energy density and caloric value, LDs have become a target compartment for metabolic engineering strategies to overproduce storage lipids in vegetative parts of plants. This process has met with remarkable success, leading to tobacco plants with lipid yields from their leaves equivalent to oil yields from oilseed crops. This overall engineering process has been described as the "push, pull and protect" concept for the efficient production and packaging of storage lipids in plant tissues [START_REF] Vanhercke | Step changes in leaf oil accumulation via iterative metabolic engineering[END_REF][START_REF] Vanhercke | Metabolic engineering for enhanced oil in biomass[END_REF]. Apparently the accumulation of lipids in leaves is at the expense of transient starch [START_REF] Chu | Shifting carbon flux from non-transient starch to lipid allows oil accumulation in transgenic tobacco leaves[END_REF], illustrating a plasticity in leaves for carbon storage that may ultimately be exploited for bioenergy and/or feed energy-densification applications.

In addition to bioenergy applications, LDs offer a stable compartment for sequestration of various hydrophobic compounds. As such, several recent reports indicated that manipulation of LD machinery can be exploited for the subcellular storage of secondary metabolites. For example, [START_REF] Sadre | Cytosolic lipid droplets as engineered organelles for production and accumulation of terpenoid biomaterials in leaves[END_REF] engineered the accumulation of sesquiterpenes (patchoulol) and diterpenes (abetadiene) into cytoplasmic LDs. Elsewhere, the expression of lipogenic proteins from mouse elevated LDs dramatically in leaves, and supported the increased accumulation of the sesquiterpene phytoalexin, capsidiol, along with TAGs [START_REF] Cai | Mouse lipogenic proteins promote the co-accumulation of triacylglycerols and sesquiterpenes in plant cells[END_REF]. With the preponderance of bioactive hydrophobic secondary metabolites, these studies illustrate the utility of engineering the cytoplasmic LD compartment in plants as a repository for high value isoprenoids in the future.

Future prospects for LD biology

In the last decade, increasing attention on cytoplasmic LDs has revealed an increasing inventory of proteins that support the formation, stability and turnover of this compartment in plant cells. Some proteins appear to have specific plant lineages, while others are conserved across kingdoms. The identification of this LD machinery will support a mechanistic examination of the interplay of these and other proteins in LD biogenesis, both in oilseeds and in non-seed tissues of plants. In addition, functions beyond neutral lipid storage continue to be revealed for LDs in different plant cell types including as a reservoir for membrane lipid remodeling [START_REF] Xu | Triacylglycerol Metabolism, Function, and Accumulation in Plant Vegetative Tissues[END_REF], a platform for the production of lipophilic signals [START_REF] Shimada | Leaf Oil Body Functions as a Subcellular Factory for the Production of a Phytoalexin in Arabidopsis[END_REF][START_REF] Fernandez-Santos | Protein Profiles of Lipid Droplets during the Hypersensitive Defense Response of Arabidopsis against Pseudomonas Infection[END_REF], and responses to environmental stress [START_REF] Yang | Functions of triacylglycerols during plant development and stress[END_REF] [START_REF] Lu | The Role of Triacylglycerol in Plant Stress Response[END_REF]. Further, an improved understanding of the cellular processes for LD formation, neutral lipid deposition and LD stability will accelerate and expand promising applications for lipid engineering. preparation of Figure 6, and to Dr. Eliot Herman, University of Arizona, for the EM images in Figure 6.

Thanks also to Charlene Case for assistance with manuscript preparation, and to Dr. Robert Mullen, University of Guelph, for comments and edits.

The dynamic nature of peroxisome structures, abundance, and subcellular interactions

Peroxisomes compartmentalize diverse oxidative reactions, allowing metabolic, signaling, and detoxification roles while limiting the potential for damage [START_REF] Kao | Peroxisome Function, Biogenesis, and Dynamics in Plants[END_REF][START_REF] Pan | Peroxisomes: versatile organelles with diverse roles in plants[END_REF].

Peroxisomes are a closed system, permeable only to small (300-400 Da) molecules [START_REF] Charton | Plant peroxisomal solute transporter proteins[END_REF][START_REF] Plett | Peroxisomal Cofactor Transport[END_REF]. Lipid substrates and ATP, NAD + , and CoA cofactors are imported via membrane transporters [START_REF] Charton | Plant peroxisomal solute transporter proteins[END_REF][START_REF] Plett | Peroxisomal Cofactor Transport[END_REF], whereas enzymes are imported by cytosolic receptors that recognize a peroxisomal targeting signal (PTS1/PTS2; [START_REF] Reumann | Prediction of Peroxisomal Matrix Proteins in Plants[END_REF][START_REF] Pan | Peroxisomes: versatile organelles with diverse roles in plants[END_REF]). Plant peroxisomes are indispensable during early development, when seedlings rely on lipid breakdown prior to photosynthetic initiation [START_REF] Graham | Seed storage oil mobilization[END_REF] for photorespiration in leaf cells, and reactive oxygen (ROS) and nitrogen species (RNS) metabolism throughout development and under changing conditions (Del Rio and Lopez-Huertas, 2016; [START_REF] Kao | Peroxisome Function, Biogenesis, and Dynamics in Plants[END_REF][START_REF] Corpas | Plant Peroxisomes: A Factory of Reactive Species[END_REF][START_REF] Pan | Peroxisomes: versatile organelles with diverse roles in plants[END_REF]Su et al., 2020).

Peroxisome abundance varies based on cell type, developmental stage, and environmental conditions. Peroxisome numbers increase in response to stress, including salt [START_REF] Mitsuya | Salt stress causes peroxisome proliferation, but inducing peroxisome proliferation does not improve NaCl tolerance in Arabidopsis thaliana[END_REF][START_REF] Fahy | Impact of salt stress, cell death, and autophagy on peroxisomes: quantitative and morphological analyses using small fluorescent probe N-BODIPY[END_REF][START_REF] Frick | Kinase MPK17 and the Peroxisome Division Factor PMD1 Influence Salt-induced Peroxisome Proliferation[END_REF], light [START_REF] Desai | Light induces peroxisome proliferation in Arabidopsis seedlings through the photoreceptor phytochrome A, the transcription factor HY5 HOMOLOG, and the peroxisomal protein PEROXIN11b[END_REF], and cadmium [START_REF] Rodriguez-Serrano | Peroxisomes Extend Peroxules in a Fast Response to Stress via a Reactive Oxygen Species-Mediated Induction of the Peroxin PEX11a[END_REF]Terron-Camero et al., 2020), and prior to cell division [START_REF] Lingard | Arabidopsis PEROXIN11c-e, FISSION1b, and DYNAMIN-RELATED PROTEIN3A cooperate in cell cycle-associated replication of peroxisomes[END_REF].

Peroxisome division occurs via fission or the budding of pre-peroxisomes from the ER [START_REF] Agrawal | De novo peroxisome biogenesis: Evolving concepts and conundrums[END_REF][START_REF] Kao | Peroxisome Function, Biogenesis, and Dynamics in Plants[END_REF][START_REF] Pan | Peroxisomes: versatile organelles with diverse roles in plants[END_REF]Su et al., 2020). Peroxisomes can be degraded via pexophagy, an organelle-specific autophagy [START_REF] Young | Pexophagy and peroxisomal protein turnover in plants[END_REF]Su et al., 2019), as part of a natural turnover ( [START_REF] Kao | Peroxisome Function, Biogenesis, and Dynamics in Plants[END_REF][START_REF] Yamauchi | Autophagy controls reactive oxygen species homeostasis in guard cells that is essential for stomatal opening[END_REF] or when excess organelles are not necessary following stress [START_REF] Calero-Munoz | Cadmium induces reactive oxygen species-dependent pexophagy in Arabidopsis leaves[END_REF] or developmental transitions [START_REF] Kim | Autophagy-related proteins are required for degradation of peroxisomes in Arabidopsis hypocotyls during seedling growth[END_REF].

Peroxisomes are small, measuring 1-2 𝜇m in Arabidopsis [START_REF] Rinaldi | The Roles of beta-Oxidation and Cofactor Homeostasis in Peroxisome Distribution and Function in Arabidopsis thaliana[END_REF] but with notable variability. Larger structures can be visualized 3-4 days post imbibition [START_REF] Rinaldi | The Roles of beta-Oxidation and Cofactor Homeostasis in Peroxisome Distribution and Function in Arabidopsis thaliana[END_REF], with some peroxisomes over 10 𝜇m in 4-day-old seedlings. This expansion is temporary, hypothesized to occur following an influx of seed-storage lipids [START_REF] Rinaldi | The Roles of beta-Oxidation and Cofactor Homeostasis in Peroxisome Distribution and Function in Arabidopsis thaliana[END_REF]. Although morphology can differ, peroxisomes are primarily spherical.

Since their identification, peroxisomes have been considered simple organelles, with typical definitions highlighting their small size, lack of a genome, and a single membrane surrounding a defined matrix. However, recent investigations by [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF] have led to an enhanced description of peroxisomes, one in which extensive internal membranes are present. The authors combined two high-sensitivity fluorescence reporters: an mRuby3-PTS to visualize the peroxisome interior and mNeonGreen tagged with an mPTS membrane peroxisomal targeting signal to label the membrane [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF]. This combination revealed the unexpected presence of internal structures, coined intralumenal vesicles (ILVs).

In 3-4 day-old samples, membrane reporters localized around the structures, but also within the interior of the organelles [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF]. Many peroxisomes contained numerous internal vesicles, which varied in size (Figure 7). As introduced above, 5-day-old seedlings showed expanded organelles that rapidly decreased in size. These size changes were concurrent with increasing ILVs and internalized membrane content. By 8 days, seedlings continued to show membrane reporters within the peroxisome lumen, with some images showing membrane signals throughout the entire structure. Watching this process suggested dense packing over time that precluded observation of individual vesicles, such that the membrane reporter appeared uniform within the lumen at this age [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF]. These seedling experiments suggest how peroxisomes mature, starting as larger, variable structures, but stabilizing at a smaller size as membranes are internalized and lipid metabolism slows.

These microscopic images led to an enhanced understanding of peroxisomal structures: peroxisomes have an outer membrane surrounding lumenal space that contains imported matrix proteins, as well as a dynamic number of membrane-bound vesicles clear of matrix proteins (Figure 7) [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF]. Further, two proteins with unique peroxisomal localization (SCO3/UP9; [START_REF] Albrecht | The cytoskeleton and the peroxisomal-targeted snowy cotyledon3 protein are required for chloroplast development in Arabidopsis[END_REF][START_REF] Quan | Proteome analysis of peroxisomes from etiolated Arabidopsis seedlings identifies a peroxisomal protease involved in beta-oxidation and development[END_REF] accumulated within a subset of ILVs that lacked matrix proteins. This apparent segregation yields at least three distinct spaces within peroxisomes, potentially housing unique proteins, substrates, cofactors, and/or environments.

Mutants disrupting β-oxidation showed alterations in ILV number, size, composition, and orientation, suggesting β-oxidation activity is required for inner membrane formation [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF].

Long-chain seed storage lipids are insoluble; [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF] hypothesize membrane internalization may reduce the solubility challenges associated with lipid mobilization in an aqueous matrix. Lipids could be degraded from the membrane, with subsequent release and degradation of the shorter, more soluble substrates, leading to the reduced organelle size common in older seedlings.

Beyond structures, imaging and biochemical studies have revealed the physical association of peroxisomes with lipid bodies, plastids, mitochondria, and the ER (Figure 7); [START_REF] Shai | No peroxisome is an island -Peroxisome contact sites[END_REF]) [START_REF] Oikawa | Re-evaluation of physical interaction between plant peroxisomes and other organelles using live-cell imaging techniques[END_REF]. Peroxisomal enzymes catalyze specific reactions within metabolic pathways, which often extend to two (or more) subcellular spaces. These organelle interactions are dynamic: peroxisomes in seedlings associate with oil bodies, for instance, whereas peroxisomes in leaves associate with chloroplasts and mitochondria [START_REF] Oikawa | Re-evaluation of physical interaction between plant peroxisomes and other organelles using live-cell imaging techniques[END_REF]. Interaction points would enhance the transfer efficiency of pathway intermediates. These sites also may facilitate transferring hydrogen peroxide and other reactive species from other organelles to peroxisomes for sequestration and degradation [START_REF] Shai | No peroxisome is an island -Peroxisome contact sites[END_REF]Su et al., 2019).

Many plant species contain oil bodies (also known as lipid droplets or lipid bodies) that store triacylglycerols for energy reserves [START_REF] Esnay | Lipid Droplet-Peroxisome Connections in Plants[END_REF]. Peroxisome-oil body association facilitates the efficient transfer of stored material for metabolism via fatty acid β-oxidation and the glyoxylate cycle. Extended interactions and peroxisomal clusters in proximity to oil bodies occur in β-oxidation mutants [START_REF] Hayashi | Direct interaction between glyoxysomes and lipid bodies in cotyledons of the Arabidopsis thaliana ped1 mutant[END_REF][START_REF] Rinaldi | The Roles of beta-Oxidation and Cofactor Homeostasis in Peroxisome Distribution and Function in Arabidopsis thaliana[END_REF], while exogenous sucrose reduces the association [START_REF] Cui | Sucrose Production Mediated by Lipid Metabolism Suppresses the Physical Interaction of Peroxisomes and Oil Bodies during Germination of Arabidopsis thaliana[END_REF], suggesting this interaction is mediated by cellular requirements for lipid mobilization.

Peroxisomes and chloroplasts can form specific pairs that remain intact over time [START_REF] Oikawa | Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysis[END_REF]. Peroxisome shape alterations expand the surface area to increase chloroplast interactions. In the light, peroxisomes extend into an elliptical shape, versus a spherical shape in darkness. Tethering factors connecting peroxisomes and chloroplasts may facilitate this interaction [START_REF] Oikawa | Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysis[END_REF][START_REF] Gao | In Vivo Quantification of Peroxisome Tethering to Chloroplasts in Tobacco Epidermal Cells Using Optical Tweezers[END_REF], potentially including the PEX10 RING finger protein [START_REF] Schumann | AthPEX10, a nuclear gene essential for peroxisome and storage organelle formation during Arabidopsis embryogenesis[END_REF][START_REF] Sparkes | An arabidopsis pex10 null mutant is embryo lethal, implicating peroxisomes in an essential role during plant embryogenesis[END_REF][START_REF] Schumann | Requirement of the C3HC4 zinc RING finger of the Arabidopsis PEX10 for photorespiration and leaf peroxisome contact with chloroplasts[END_REF]. A dominant-negative PEX10 line had clustered peroxisomes that did not associate with chloroplasts; this line had phenotypes similar to photorespiration mutants [START_REF] Schumann | Requirement of the C3HC4 zinc RING finger of the Arabidopsis PEX10 for photorespiration and leaf peroxisome contact with chloroplasts[END_REF], consistent with a role for organelle association in efficient metabolic transfer.

Mitochondria also appear in close proximity to both peroxisomes and chloroplasts in the light, consistent with their interactive metabolic roles [START_REF] Oikawa | Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysis[END_REF]. Peroxisomes associate with mitochondria in stress conditions as well; increasing interactions are seen in cells exposed to high ROS and might be important for ROS neutralization [START_REF] Jaipargas | High Light Intensity Leads to Increased Peroxule-Mitochondria Interactions in Plants[END_REF][START_REF] Mathur | Organelle extensions in plant cells[END_REF].

Finally, peroxisomes show a close proximity with the ER (Barton et al., 2013;[START_REF] Oikawa | Re-evaluation of physical interaction between plant peroxisomes and other organelles using live-cell imaging techniques[END_REF].

Interestingly, one of the two mPTS signals used by [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF] revealed accumulation in peroxisomes and reticular membranes thought to be ER. This finding is consistent with a hypothesis that the membrane protein was trafficked through the ER or has a dual function at both sites [START_REF] Wright | Peroxisomes form intralumenal vesicles with roles in fatty acid catabolism and protein compartmentalization in Arabidopsis[END_REF].

Another shape alteration to peroxisomes are thin organelle protrusions known as peroxules [START_REF] Mathur | Organelle extensions in plant cells[END_REF]. These structures are up to 15 𝜇m, dramatically increasing the surface area [START_REF] Sinclair | Peroxule extension over ER-defined paths constitutes a rapid subcellular response to hydroxyl stress[END_REF]Barton et al., 2013). Formation of these organelle extensions is transient and dynamic [START_REF] Mathur | Organelle extensions in plant cells[END_REF]. The interactions between organelles described above may be mediated by peroxules, including proposed interactions with oil bodies (Thazar-Poulot et al., 2015), chloroplasts [START_REF] Schumann | Requirement of the C3HC4 zinc RING finger of the Arabidopsis PEX10 for photorespiration and leaf peroxisome contact with chloroplasts[END_REF], mitochondria [START_REF] Jaipargas | High Light Intensity Leads to Increased Peroxule-Mitochondria Interactions in Plants[END_REF], and ER [START_REF] Sinclair | Peroxule extension over ER-defined paths constitutes a rapid subcellular response to hydroxyl stress[END_REF]. Extended structures are seen following H 2 O 2 , UV-A, and hydroxyl radical stress, but retract when stress is minimized [START_REF] Sinclair | Peroxule extension over ER-defined paths constitutes a rapid subcellular response to hydroxyl stress[END_REF]. In addition, elongations are common during the constriction and fission steps of peroxisome division [START_REF] Sinclair | Peroxule extension over ER-defined paths constitutes a rapid subcellular response to hydroxyl stress[END_REF]Barton et al., 2013). Cadmium induces ROS and similarly leads to peroxule formation that results in division to increase peroxisome numbers [START_REF] Rodriguez-Serrano | Peroxisomes Extend Peroxules in a Fast Response to Stress via a Reactive Oxygen Species-Mediated Induction of the Peroxin PEX11a[END_REF]. Increased peroxule frequency in cells exposed to ROS leads to a hypothesis that extensions facilitate neutralization to prevent or reduce damage [START_REF] Sinclair | Peroxule extension over ER-defined paths constitutes a rapid subcellular response to hydroxyl stress[END_REF]Barton et al., 2013;[START_REF] Rodriguez-Serrano | Peroxisomes Extend Peroxules in a Fast Response to Stress via a Reactive Oxygen Species-Mediated Induction of the Peroxin PEX11a[END_REF]. Separately, peroxule-mediated contacts might assist in protein localization. The SDP1 lipase [START_REF] Eastmond | SUGAR-DEPENDENT1 encodes a patatin domain triacylglycerol lipase that initiates storage oil breakdown in germinating Arabidopsis seeds[END_REF] localizes to peroxisomal membranes and then the oil body, a transition concurrent with peroxule development (Thazar-Poulot et al., 2015) The refined visualization of peroxisome structures and our increasing understanding of organelle interactions has led to an enhanced view of peroxisomes compared to a previously simple model.

Many open questions about peroxisome biology remain. What is the mechanism of and importance for dynamic membrane changes for peroxisomes in adult tissues and under changing environmental conditions? How do peroxisome substructures form and how are membrane and matrix proteins sorted to create unique environments or provide specific functionality? Understanding such details about peroxisome structures, as well as the factors promoting and mediating peroxisome interactions with other organelles, will continue to increase our understanding of these dynamic organelles.

Plant mitochondria

In plants, mitochondria provide a large portion of the ATP in the cytosol through oxidative phosphorylation, and are the sites of some amino acids, nucleic acids, lipids, and plant hormone metabolism. Plant mitochondria also control redox balances when photosynthesis is on or off or fluctuating [START_REF] Noguchi | Interaction between photosynthesis and respiration in illuminated leaves[END_REF][START_REF] Finkemeier | Mitochondrial Regulation and Signalling in the Photosynthetic Cell: Principles and Concepts[END_REF], have roles in cellular signaling [START_REF] Huang | The Roles of Mitochondrial Reactive Oxygen Species in Cellular Signaling and Stress Response in Plants[END_REF][START_REF] Welchen | Cross-talk between mitochondrial function, growth, and stress signalling pathways in plants[END_REF][START_REF] Welchen | Cross-talk between mitochondrial function, growth, and stress signalling pathways in plants[END_REF], and in resistance to diseases [START_REF] Fuchs | Single organelle function and organization as estimated from Arabidopsis mitochondrial proteomics[END_REF]. In agriculture, cytoplasmic male sterility, which is caused by genes encoded in the mitochondrial genome, is used for production of F1 hybrid seeds in diverse crops and vegetables. The fine structure and dynamics of plant mitochondria are briefly reviewed here.

Mitochondria have two lipid bilayers that form the outer and inner membrane (Figure 8). Some parts of the inner membrane are invaginated to form sacs, called cristae, that increase the area of oxidative phosphorylation complexes. Five diverse eukaryotic-conserved complexes are embedded in the cristae membrane. In contrast, plant-specific proteins (alternative oxidases and extra NDH and NDPH dehydrogenases) for alternative respiration pathways mainly reside in the non-cristae parts of the inner-membrane [START_REF] Schwarzlander | Plant mitochondrial membranes: adding structure and new functions to respiratory physiology[END_REF]. Plant ATP synthase dimers (complex V) are in the cristae membrane, where they contribute to its curvature [START_REF] Zancani | Structural and functional properties of plant mitochondrial F-ATP synthase[END_REF] Complexes I to V have roles in oxidative phosphorylation. Some of them form super-complexes for functional efficiency and to regulate oxidative phosphorylation [Braun, 2020 #2342. Protein-protein interactions and metabolite channeling are observed also in the TCA cycle in the matrix [START_REF] Zhang | Nanoscale movements of cellulose microfibrils in primary cell walls[END_REF][Zhang, #2382)).

Additionally, glycolysis enzymes in the cytosol dynamically associate on the outer surface of mitochondria [START_REF] Giege | Enzymes of glycolysis are functionally associated with the mitochondrion in Arabidopsis cells[END_REF][START_REF] Graham | Glycolytic enzymes associate dynamically with mitochondria in response to respiratory demand and support substrate channeling[END_REF], probably to more efficiently transport metabolites.

The mitochondrial outer membrane has the most abundant protein in plant mitochondria, the Voltage-Dependent Anion Channel (VDAC1). A single mitochondrion contains 40,000 VDACs out of a total of 1.4 million proteins [START_REF] Fuchs | Single organelle function and organization as estimated from Arabidopsis mitochondrial proteomics[END_REF]. The outer membrane does not just encapsulate the inner membrane but also sometimes extends into the cytosol and other organelles (without extending the inner-membrane), and occasionally the extensions are pinched off to form small vesicle-like structures [START_REF] Yamashita | Formation of Mitochondrial Outer Membrane Derived Protrusions and Vesicles in Arabidopsis thaliana[END_REF]; Figure 8D). In mammals, mitochondria-derived vesicles which do not contain inner membranes are reported to be involved in the transport of specific proteins to peroxisomes, endosomes, and multivesicular bodies (Sugiura et al., 2014), and in the biogenesis of peroxisomes [Sugiura, 2017[Sugiura, #2376. . Each Arabidopsis leaf cell contains 300-450 mitochondria. Many plant mitochondria move along microfilaments at 0.05 -3 µm/sec [Doniwa, 2007[Doniwa, #2344]] [START_REF] Oikawa | Mitochondrial movement during its association with chloroplasts in Arabidopsis thaliana[END_REF], which is about an order of magnitude faster than mammalian and yeast mitochondria, which mainly move along microtubules. Some plant mitochondria stop and wiggle, as if they were anchored to the cytoskeleton or other organelles, such as plastids and peroxisomes [START_REF] Jaipargas | High Light Intensity Leads to Increased Peroxule-Mitochondria Interactions in Plants[END_REF][START_REF] Oikawa | Mitochondrial movement during its association with chloroplasts in Arabidopsis thaliana[END_REF][START_REF] Cui | Sucrose Production Mediated by Lipid Metabolism Suppresses the Physical Interaction of Peroxisomes and Oil Bodies during Germination of Arabidopsis thaliana[END_REF]. Moving plant mitochondria can change their speed and also can change their shapes from granular to linear to attach to other organelles in response to the presence of sucrose or light [START_REF] Jaipargas | High Light Intensity Leads to Increased Peroxule-Mitochondria Interactions in Plants[END_REF]. A single plant cell can have mitochondria with different shapes [START_REF] Jaipargas | Mitochondrial pleomorphy in plant cells is driven by contiguous ER dynamics[END_REF], different DNA contents (Arimura et al., 2004b;[START_REF] Preuten | Fewer genes than organelles: extremely low and variable gene copy numbers in mitochondria of somatic plant cells[END_REF], (Figure 8E) and transiently fluctuating membrane potentials [START_REF] Schwarzlander | Pulsing of membrane potential in individual mitochondria: a stress-induced mechanism to regulate respiratory bioenergetics in Arabidopsis[END_REF]. In addition, as shown in Figure 8B, differently colored groups of mitochondria in a cell achieve a unified color in two hours, indicating that mitochondria undergo frequent fusion and fission, resulting in the sharing of internal proteins. Mitochondria involved in such dynamic sharing of materials in a plant cell are referred to as a dynamic syncytium [START_REF] Lonsdale | The Plant Mitochondrial Genome -Homologous Recombination as a Mechanism for Generating Heterogeneity[END_REF], and the collective mitochondria in a cell are thought to exist as a discontinuous whole [START_REF] Logan | The dynamic chondriome[END_REF]. Fusion of mitochondria results in the formation of elongated and/or branched mitochondria in some meristematic tissues, such as shoot apical apices (Segui-Simarro and [START_REF] Segui-Simarro | Cell cycle-dependent changes in Golgi stacks, vacuoles, clathrin-coated vesicles and multivesicular bodies in meristematic cells of Arabidopsis thaliana: a quantitative and spatial analysis[END_REF], germinating seeds [START_REF] Paszkiewicz | Arabidopsis Seed Mitochondria Are Bioenergetically Active Immediately upon Imbibition and Specialize via Biogenesis in Preparation for Autotrophic Growth[END_REF], and dedifferentiating protoplasts [START_REF] Sheahan | Mitochondria as a connected population: ensuring continuity of the mitochondrial genome during plant cell dedifferentiation through massive mitochondrial fusion[END_REF][START_REF] Rose | New Beginnings: Mitochondrial Renewal by Massive Mitochondrial Fusion[END_REF].

Mitochondrial fission is achieved by dynamin-related proteins that are well-conserved in eukaryotes (e.g. DRP3A and 3B in Arabidopsis [START_REF] Arimura | A dynamin-like protein (ADL2b), rather than FtsZ, is involved in Arabidopsis mitochondrial division[END_REF]Arimura et al., 2004a;Arimura et al., 2004b;[START_REF] Fujimoto | Arabidopsis dynamin-related proteins DRP3A and DRP3B are functionally redundant in mitochondrial fission, but have distinct roles in peroxisomal fission[END_REF], Figure 8A), which polymerize to form ring-like structures outside mitochondria [START_REF] Ingerman | Dnm1 forms spirals that are structurally tailored to fit mitochondria[END_REF]. Plant-specific ELM1, an outer surface protein, localizes DRP3s from the cytosol [START_REF] Arimura | Arabidopsis ELONGATED MITOCHONDRIA1 is required for localization of DYNAMIN-RELATED PROTEIN3A to mitochondrial fission sites[END_REF]). An outer-membrane embedded protein that is conserved in eukaryotes (Fis1) functions as a molecular adapter for DRP in budding yeast [START_REF] Okamoto | Mitochondrial morphology and dynamics in yeast and multicellular eukaryotes[END_REF]. Fis1 had been thought to carry out the similar functions but is now thought to have only a rather minor and indirect role in mitochondrial fission in both mammals [START_REF] Otera | Mff is an essential factor for mitochondrial recruitment of Drp1 during mitochondrial fission in mammalian cells[END_REF][START_REF] Giacomello | The cell biology of mitochondrial membrane dynamics[END_REF] and plants [START_REF] Nagaoka | DRP3 and ELM1 are required for mitochondrial fission in the liverwort Marchantia polymorpha[END_REF][START_REF] Arimura | Fission and Fusion of Plant Mitochondria, and Genome Maintenance[END_REF]. Other factors may be involved in plant mitochondrial fission, such as factors involved in cold-induced fission [START_REF] Arimura | Cold Treatment Induces Transient Mitochondrial Fragmentation in Arabidopsis thaliana in a Way that Requires DRP3A but not ELM1 or an ELM1-Like Homologue[END_REF] or factors that are independent of DRP and specific to Brassicaceae [START_REF] Aung | The Arabidopsis tail-anchored protein PEROXISOMAL AND MITOCHONDRIAL DIVISION FACTOR1 is involved in the morphogenesis and proliferation of peroxisomes and mitochondria[END_REF]. On the other hand, no orthologues, factors or molecular mechanisms are known with certainty to be involved in mitochondrial fusion in plants. However, Friendly (FMT) is suggested to mediate inter-mitochondrial association before mitochondrial fusion because in fmt mutants, mitochondria gather together [START_REF] Logan | The genetic control of plant mitochondrial morphology and dynamics[END_REF] but do not fuse (El Zawily et al., 2014).

Mitochondrial-specific autophagy (mitophagy) has been extensively studied in mammals and yeasts [START_REF] Onishi | Molecular mechanisms and physiological functions of mitophagy[END_REF], in which it is involved in mitochondrial quality control. In these organisms, degraded mitochondria with low membrane potential could not fuse with other "healthy" mitochondria, but they are specifically recognized, captured and engulfed by autophagosome membranes (Figure 8C). The engulfed mitochondria are transported to the vacuole to be digested to prevent accidental ROS generation and/or other negative effects. Therefore, mitophagy, fission, and fusion are thought to function as a quality control system for all the mitochondria in a cell (Twig et al., 2008).

Mitochondrial-specific degradation in Arabidopsis has also been observed in several situations, including during leaf senescence [START_REF] Broda | Mitophagy: A Mechanism for Plant Growth and Survival[END_REF] greening of cotyledons [START_REF] Ma | Friendly mediates membrane depolarization-induced mitophagy in Arabidopsis[END_REF], after UV-irradiation [START_REF] Nakamura | Autophagy Contributes to the Quality Control of Leaf Mitochondria[END_REF] and after treating the inner membrane with ionophores (Ma et al., 2021). Orthologues of factors specific to mitophagy in mammals and yeasts have not yet been found in plant genomes, although FMT was recently reported to be involved in mitophagy in Arabidopsis [START_REF] Ma | Friendly mediates membrane depolarization-induced mitophagy in Arabidopsis[END_REF].

Super resolution microscopy is a promising new technique that can clarify the internal structures of mitochondria in more detail with their diverse physiology and functions. In addition, recent trials to understand molecular numbers of averaged single mitochondrion [START_REF] Fuchs | Single organelle function and organization as estimated from Arabidopsis mitochondrial proteomics[END_REF] or estimated single mitochondrion [START_REF] Moller | What is hot in plant mitochondria?[END_REF] will hopefully develop to the next stage of analysis of the exact individual mitochondria to answer the actual quantitative dynamics of molecules (or "factfulness") of diverse mitochondria. Until recently, transformation of mitochondrial genomes in multicellular plants has been impossible, but new genome editing methods [START_REF] Kazama | Curing cytoplasmic male sterility via TALEN-mediated mitochondrial genome editing[END_REF][START_REF] Arimura | Targeted gene disruption of ATP synthases 6-1 and 6-2 in the mitochondrial genome of Arabidopsis thaliana by mitoTALENs[END_REF] have opened the door to analyzing the functions of mitochondrial genes, as well as to regulate their expressions to breed crops with agriculturally important characteristics.

Chloroplast -A Plant's Powerhouse with Tunable Performance

A unique endosymbiotic event more than 900 Ma ago was the starting point for the evolution of the chloroplast from a free-living cyanobacterial precursor [START_REF] Sibbald | Genomic Insights into Plastid Evolution[END_REF]. Every second, the thylakoid membrane system of a modern chloroplast in Viridiplantae can convert energy from the sun in up to 80 million ATP and NADPH + H + molecules which fuels a number of anabolic reactions localized in the chloroplast stroma, like syntheses of sugars, lipids/fatty acids, amino acids, nucleotides, pigments, alkaloids, hormones, and vitamins [START_REF] Kirchhoff | Chloroplast ultrastructure in plants[END_REF]. Furthermore, a battery of membrane-embedded chloroplast envelope transporters makes the competency for photosynthetic energy transforming available to the entire cell and beyond [START_REF] Weber | Connecting the Plastid: Transporters of the Plastid Envelope and Their Role in Linking Plastidial with Cytosolic Metabolism[END_REF].

Chloroplast Lifecycle

During the last decade, electron tomography has provided detailed structural insights in the morphological transitions from an undifferentiated, non-photosynthetic proplastid to a mature chloroplast in the plant shoot apical meristem for illuminated shoots [START_REF] Adam | Biogenesis of thylakoid networks in angiosperms: knowns and unknowns[END_REF][START_REF] Charuvi | Gain and loss of photosynthetic membranes during plastid differentiation in the shoot apex of Arabidopsis[END_REF] or via the etioplast with its characteristic para-crystalline prolamellar body [START_REF] Kowalewska | Three-Dimensional Visualization of the Tubular-Lamellar Transformation of the Internal Plastid Membrane Network during Runner Bean Chloroplast Biogenesis[END_REF]. The correlation between the sequential appearance of proteins like photosystem I and II, lightharvesting complex II, CURT1 proteins, ATPase, protochlorophyllide oxidoreductase, and plastidial ribosomes on the one hand and structural developments of the plastid, on the other hand, provides insights on the role of particular proteins for the proplastid-chloroplast differentiation [START_REF] Kowalewska | Three-Dimensional Visualization of the Tubular-Lamellar Transformation of the Internal Plastid Membrane Network during Runner Bean Chloroplast Biogenesis[END_REF]Liang et al., 2018a;[START_REF] Floris | Molecular landscape of etioplast inner membranes in higher plants[END_REF]. Proplastid development requires a massive protein import from the cytoplasm into the chloroplast mainly by the TOC/TIC translocase system [START_REF] Aronsson | The chloroplast protein import apparatus, its components, and their roles[END_REF][START_REF] Ling | Chloroplast Biogenesis Is Regulated by Direct Action of the Ubiquitin-Proteasome System[END_REF] since ~95% of chloroplast proteins are nuclearencoded. Currently, two non-exclusive models are mainly discussed as to how hydrophobic nuclearencoded proteins along with lipids and pigments, that are synthesized at the plastid envelope membranes, are transported through the aqueous stroma to reach their thylakoid membrane destination: (1) invaginations of the inner envelope membrane/direct contact sites with thylakoids and

(2) vesicle transport [START_REF] Lindquist | Chloroplast vesicle transport[END_REF][START_REF] Mechela | A brief history of thylakoid biogenesis[END_REF]. Evidence exists that the invagination/direct contact site pathway is realized only in the proplastid-to-chloroplast transition whereas vesicle transport seems dominant in mature chloroplasts [START_REF] Vothknecht | Biogenesis and origin of thylakoid membranes[END_REF][START_REF] Andersson | Chloroplast membrane lipid biosynthesis and transport[END_REF][START_REF] Lindquist | Chloroplast vesicle transport[END_REF]. For the latter, the role of typical vesicle forming proteins like COPI, COPII, SNARE, or VIPP1 for plastid biogenesis has still to be determined [START_REF] Mechela | A brief history of thylakoid biogenesis[END_REF]. However, for cyanobacterial VIPP1 a structure-based molecular understanding was recently achieved [START_REF] Gupta | Structural basis for VIPP1 oligomerization and maintenance of thylakoid membrane integrity[END_REF]. In contrast to proplastids, mature chloroplasts propagate by binary fission [START_REF] Osteryoung | Division and Dynamic Morphology of Plastids[END_REF][START_REF] Yoshida | Insights into the Mechanisms of Chloroplast Division[END_REF]. The plastid division machinery is made of four physically connected supramolecular ring structures, two outside (an outer polyglucan plastid-dividing ring and a dynamin-related ring) and two inside the chloroplast (an inner plastid-dividing ring and a tubulin-like FtsZ-ring beneath the inner envelope membrane). In a concerted mechanism, the rings generate the mechanical force for plastid constriction and eventually division. An example for the crucial role of regulatory proteins for plastid morphogenesis is visualized in Fig. 1. Open questions in the field are the composition of the inner plastid-dividing ring, how thylakoid membranes divide, and how chloroplast division is coordinated with cell and other organelle divisions [START_REF] Osteryoung | Division and Dynamic Morphology of Plastids[END_REF][START_REF] Yoshida | Insights into the Mechanisms of Chloroplast Division[END_REF]. At the end of their life span, chloroplasts enter highly coordinated dismantling processes with the goal to minimize reactive oxygen production (ROS) and recycling of their abundant macromolecules to sink tissues of the plant (Avila-Ospina et al., 2014).,Strikingly chloroplasts hold ~80% of leaf nitrogen [START_REF] Makino | Effects of Nitrogen Nutrition on Nitrogen Partitioning between Chloroplasts and Mitochondria in Pea and Wheat[END_REF]. It seems that ROS-dependent retrograde signaling plays a key role for coordinating chloroplast degradation via multiple breakdown pathways including chlorophagy [START_REF] Woodson | Chloroplast stress signals: regulation of cellular degradation and chloroplast turnover[END_REF][START_REF] Dominguez | Chloroplast dismantling in leaf senescence[END_REF].

Current research focuses on elucidating how particular environmental conditions trigger specific dismantling pathways and deciphering the corresponding signal cascades.

Structural membrane dynamics as a means to control energy conversion

The fact that photosynthetic energy conversion has to integrate and balance significant fluctuations in both cell metabolism (including CO 2 availability) and energy input by sunlight in an oxidizing environment, calls for its strict regulation to minimize toxic ROS production. In the last decade, it turns out that a central regulatory element for tuning photosynthetic performance in plants is the dynamic adjustment of lateral and transversal geometric (grana) thylakoid dimensions that regulate electron transport, light-harvesting, and protein repair [START_REF] Kirchhoff | Dynamic control of protein diffusion within the granal thylakoid lumen[END_REF][START_REF] Puthiyaveetil | Compartmentalization of the protein repair machinery in photosynthetic membranes[END_REF][START_REF] Hepworth | Dynamic thylakoid stacking and state transitions work synergistically to avoid acceptor-side limitation of photosystem I[END_REF]. It is an open question how reversible protein phosphorylation, physicochemical membrane properties, and protein composition dynamics work together to control architectural thylakoid features and further on energy conversion.

Future perspectives: For the next five to ten years, the fast methodical and technological development in (cryo-)electron tomography is expected to provide detailed new insights into chloroplast structure-function relationships not only for the mature plastid but also for its biogenesis and dismantling. Furthermore, studying chloroplast diversity in non-model, more exotic species as well as in specialized plant tissues and organs (including transitions between different plastid types) gains increasing attention because it will uncover the metabolic plasticity and diversity of this organelle. Along these lines, current and future bioengineering tools for chloroplasts offer potential for improving crop plants by tuning for example non-photochemical quenching or photorespiratory pathways or for using the anabolic competence of the plastid to employ them as metabolic factory for valuable chemicals.

Plant Membrane Contact Sites -Questions from the membrane interface.

Membrane Contact sites: Does one definition fit all?

Membrane Contact Sites (MCS) are evolutionarily conserved structures where the close proximity between two or more membrane-bound organelles enables direct exchange of molecules and facilitates coordinated inter-organelle adaptive responses (Figure 10 A-B). Recent advances in plant cell imaging and the development of novel genetic and molecular tools have fueled an emerging field of research devoted to the investigation of their structural organization, dynamics, and physiological functions. This interest is uncovering plant-specific MCS structures and molecular mechanisms, but also exposing some limitations on the commonly accepted definitions and physiological functions inferred from different model organisms. As in yeast and animal cells, the plant endoplasmic reticulum (ER) is an interconnected organelle that establishes MCS with multiple cellular structures including the plasma membrane (PM), mitochondria, endosomes, peroxisomes, Golgi and trans-Golgi network (Barton et al., 2013;[START_REF] Lai | ER stress signaling requires RHD3, a functionally conserved ER-shaping GTPase[END_REF][START_REF] Wang | The Plant Cytoskeleton, NET3C, and VAP27 Mediate the Link between the Plasma Membrane and Endoplasmic Reticulum[END_REF][START_REF] Perez-Sancho | The Arabidopsis Synaptotagmin1 Is Enriched in Endoplasmic Reticulum-Plasma Membrane Contact Sites and Confers Cellular Resistance to Mechanical Stresses[END_REF][START_REF] Wang | Plant AtEH/Pan1 proteins drive autophagosome formation at ER-PM contact sites with actin and endocytic machinery[END_REF][START_REF] Brandizzi | Maintaining the structural and functional homeostasis of the plant endoplasmic reticulum[END_REF]. Unique to plants; however, are the functional interactions at ER-plastids MCS for lipid synthesis and transport [START_REF] Liu | Communications Between the Endoplasmic Reticulum and Other Organelles During Abiotic Stress Response in Plants[END_REF], the control of intercellular communication through plasmodesmata MCS-regulated intercellular bridges [START_REF] Perez-Sancho | Stitching Organelles: Organization and Function of Specialized Membrane Contact Sites in Plants[END_REF], and the MCS activities driven by a super-continuum that encompasses the cell wall, PM, ER, and cytoskeleton [START_REF] Wang | The Plant Cytoskeleton, NET3C, and VAP27 Mediate the Link between the Plasma Membrane and Endoplasmic Reticulum[END_REF][START_REF] Perez-Sancho | The Arabidopsis Synaptotagmin1 Is Enriched in Endoplasmic Reticulum-Plasma Membrane Contact Sites and Confers Cellular Resistance to Mechanical Stresses[END_REF][START_REF] Zang | A novel plant actin-microtubule bridging complex regulates cytoskeletal and ER structure at ER-PM contact sites[END_REF]. These plant-specific features are placing MCS research in plants at the forefront of discovery, broadening the MCS definition beyond yeast and animal systems.

In plants, MCS can be defined as environmentally and developmentally regulated microdomains with an intermembrane gap as small as 3 nm in plasmodesmata, and an arbitrarily defined upper limit of 80-100 nm. Plant MCS are enriched with a variety of protein-protein, and/or protein-cytoskeleton tethering assemblies that establish dynamic interactions with membrane phospholipids and/or the cell wall. Plant MCS carry out essential cellular functions including, but not restricted to maintenance of membrane lipid homeostasis, cell-to-cell communication, organelle biogenesis, autophagy, endocytosis, receptor kinase signaling, and the regulation of Ca 2+ -dependent stress responses (Figure 10 C-D), reviews in [START_REF] Perez-Sancho | Stitching Organelles: Organization and Function of Specialized Membrane Contact Sites in Plants[END_REF][START_REF] Wang | NETWORKED 3B: a novel protein in the actin cytoskeleton-endoplasmic reticulum interaction[END_REF][START_REF] Liu | Communications Between the Endoplasmic Reticulum and Other Organelles During Abiotic Stress Response in Plants[END_REF][START_REF] Petit | Dare to change, the dynamics behind plasmodesmata-mediated cell-to-cell communication[END_REF][START_REF] Rosado | Geometry and cellular function of organelle membrane interfaces[END_REF].

Lipid transfer at MCS: Is that what plant tethers do?

Due to their hydrophobicity the transport of lipids between organelles relies on either vesicle-mediated delivery mechanisms or MCS-localized lipid transport proteins (LTP) [START_REF] Scorrano | Coming together to define membrane contact sites[END_REF]. Most MCS-localized LTPs contain an internal hydrophobic cavity adapted to solubilize water-insoluble molecules (Wong and[START_REF] Wong | Lipid transfer proteins do their thing anchored at membrane contact sites . . . but what is their thing[END_REF][START_REF] Wong | Lipid transfer proteins do their thing anchored at membrane contact sites . . . but what is their thing[END_REF], are anchored to the ER by either transmembrane domains or stable interactions with ER-anchored proteins [START_REF] Scorrano | Coming together to define membrane contact sites[END_REF], and interact with the opposing membrane mainly through domains that bind anionic lipids [START_REF] Perez-Lara | Extended synaptotagmins (E-Syts): Architecture and dynamics of membrane contact sites revealed[END_REF]. In animal cells or yeast, direct lipid transport using MCS-localized LTPs may be one of the most notorious and documented MCS functions and lipid species transferred using this mechanism include sterols, ceramides, phosphatidylserine (PS), phosphatidylinositol 4-phosphate (PI4P), and diacylglycerol (DAG) [START_REF] Wu | Here, there, and everywhere: The importance of ER membrane contact sites[END_REF]. In plants, the emerging view is that MCS-localized LTPs similarly participate in the delivery of lipids between the ER and organelles not linked by vesicular traffic, (e.g. mitochondria and plastids) but also in the bulk transport of lipids between organelles connected by the secretory pathway. In a recent landmark study Ruiz-Lopez et al. showed that stress signals regulate the activity of two members of the Synaptotagmin (SYT) family of LTPs at ER-PM MCS (SYT1 and SYT3), and demonstrated their function as LTPs transferring DAG between the PM and the ER in vivo [START_REF] Ruiz-Lopez | Synaptotagmins at the endoplasmic reticulum-plasma membrane contact sites maintain diacylglycerol homeostasis during abiotic stress[END_REF]. The authors propose a geometrical model where SYT activities counteract the stress-induced build-up of conically shaped DAG at the PM and prevent the generation of areas of negative membrane curvature that could disrupt the stability of the PM during stress episodes.

MCS in motion: Who controls MCS plasticity and dynamics?

The molecular composition, geometry, and plasticity of inter-organelle junctions determine their ability to integrate and respond to cellular signals. Recent studies provide an emerging picture where MCS tethers do not act in isolation but interact with anionic lipids, cytoskeletal elements, and regulate the plasticity, function and dynamics of these cellular microdomains.

Anionic phospholipids represent only a few percent of the total lipids but they are critical biochemical and biophysical landmarks of membrane identity [START_REF] Noack | Functions of Anionic Lipids in Plants[END_REF]. Within the endomembrane system, they consist of the phosphoinositides (PIPs), phosphatidic acid (PA) and phosphatidylserine (PS), and determine the electrostatic potential of each membrane being highest at the PM, intermediate in endosomes and low in the ER [START_REF] Platre | A Combinatorial Lipid Code Shapes the Electrostatic Landscape of Plant Endomembranes[END_REF][START_REF] Dubois | Anionic phospholipid gradients: an uncharacterized frontier of the plant endomembrane network[END_REF].

In vitro or in silico data for MCS tethers such as the synaptotagmins (SYTs), Multiple C2 domains and transmembrane region (MCTPs), and Vesicle-associated membrane protein (VAMP)-associated proteins (VAPs) families, support the notion that anionic lipids profoundly impact the structure and function of MCSs by enabling protein-lipid interactions that regulate the association of the ER with the PM, TGN and early endosomes [START_REF] Perez-Sancho | The Arabidopsis Synaptotagmin1 Is Enriched in Endoplasmic Reticulum-Plasma Membrane Contact Sites and Confers Cellular Resistance to Mechanical Stresses[END_REF][START_REF] Renna | TGNap1 is required for microtubule-dependent homeostasis of a subpopulation of the plant trans-Golgi network[END_REF][START_REF] Brault | Multiple C2 domains and transmembrane region proteins (MCTPs) tether membranes at plasmodesmata[END_REF][START_REF] Ruiz-Lopez | Synaptotagmins at the endoplasmic reticulum-plasma membrane contact sites maintain diacylglycerol homeostasis during abiotic stress[END_REF]. Interestingly, these interactions appear to be mostly non-specific, the primary determinants being the negative charge carried by the anionic lipids and, in some cases, the presence of Ca 2+ [START_REF] Schapire | Arabidopsis Synaptotagmin 1 Is Required for the Maintenance of Plasma Membrane Integrity and Cell Viability[END_REF]. Accordingly, electrostatic interactions between phosphatidylinositol-4-phosphate (PI4P) and SYT1/SYT3 underpin the localization of SYT1/SYT3 to ER-PM MCS [START_REF] Ruiz-Lopez | Synaptotagmins at the endoplasmic reticulum-plasma membrane contact sites maintain diacylglycerol homeostasis during abiotic stress[END_REF], MCTP4 to plasmodesmata-MCS, [START_REF] Brault | Multiple C2 domains and transmembrane region proteins (MCTPs) tether membranes at plasmodesmata[END_REF], and the remodelling of SYT1 ER-PM MCS in response to rare-earth elements [START_REF] Lee | Rare earth elements induce cytoskeleton-dependent and PI4P-associated rearrangement of SYT1/SYT5 endoplasmic reticulum-plasma membrane contact site complexes in Arabidopsis[END_REF]. Similarly, the accumulation of phosphatidylinositol 4,5-biphosphate (PI(4,5)P 2 ) at the at the PM enables interactions with SYT1 and correlates with the rearrangement and expansion of ER-PM MCS in response to ionic stress [START_REF] Lee | Ionic stress enhances ER-PM connectivity via phosphoinositide-associated SYT1 contact site expansion in Arabidopsis[END_REF].

MCS plasticity is also controlled by components that crosslink the actin cytoskeleton at MCS and create trapping mechanisms that influence MCS architecture and expansion. In plants, this crosslinking seems to be carried out by a plant-specific complex that includes the actin associated NET3C protein, and the microtubule associated Kinesin light chain related (KLCRs) and IQ67-Domain (IQD) proteins [START_REF] Zang | A novel plant actin-microtubule bridging complex regulates cytoskeletal and ER structure at ER-PM contact sites[END_REF] (Figure 10E). Remarkably, in plants, the presence of cell walls underlies the formation of a plant-specific supramolecular assembly known as the MCS super-continuum. This super-continuum encompasses the cell wall, PM, ER, and cytoskeleton, and renders MCS with distinct kinetics, shapes, geometries, and functions [START_REF] Rosado | Geometry and cellular function of organelle membrane interfaces[END_REF][START_REF] Zang | A novel plant actin-microtubule bridging complex regulates cytoskeletal and ER structure at ER-PM contact sites[END_REF].

Recent studies propose that the MCS super-continuum serves as a nexus that limits the mobility of MCS tethering assemblies [START_REF] Wang | Plant VAP27 proteins: domain characterization, intracellular localization and role in plant development[END_REF][START_REF] Lee | Ionic stress enhances ER-PM connectivity via phosphoinositide-associated SYT1 contact site expansion in Arabidopsis[END_REF][START_REF] Zang | A novel plant actin-microtubule bridging complex regulates cytoskeletal and ER structure at ER-PM contact sites[END_REF], and control their activities. Examples of regulation mediated by this continuum may include the activity of receptor like kinases in pollen and/or stomatal cells [START_REF] Ho | Modulators of Stomatal Lineage Signal Transduction Alter Membrane Contact Sites and Reveal Specialization among ERECTA Kinases[END_REF][START_REF] Duckney | NETWORKED2-subfamily proteins regulate the cortical actin cytoskeleton of growing pollen tubes and polarised pollen tube growth[END_REF], and the regulation of phospholipase C mediated stress signals at the PM [START_REF] Ruiz-Lopez | Synaptotagmins at the endoplasmic reticulum-plasma membrane contact sites maintain diacylglycerol homeostasis during abiotic stress[END_REF]. Last, a unique type of regulation occurs at plasmodesmata MCS where the transfer of molecules is parallel to the membranes, as opposed to orthogonal to them. In those ER-PM MCS the intermembrane space may not be solely regulated by the tethers, lipids, and cytoskeleton elements in the super-continuum, but also by wall polymers (e.g. callose), which are locally synthesized around the PD structure [START_REF] Petit | Dare to change, the dynamics behind plasmodesmata-mediated cell-to-cell communication[END_REF].

Future MCS research: What's in the plant toolkit?

MCS are microdomains with an intermembrane distance below the resolution limit of conventional fluorescence microscopy, and with a dynamic behavior that requires the use of live-cell compatible techniques [START_REF] Mcfarlane | Multiscale Structural Analysis of Plant ER-PM Contact Sites[END_REF]. In recent years, advances in super-resolution microscopy (e.g. Total Internal Reflection Fluorescence (TIRF), and Structure illumination microscopy (SIM), (Figure 10F), and electron tomography (Figure 10G) are providing for the first time detailed high-resolution visualizations and 3D reconstructions of the MCS ultrastructure in plants (Baillie et al., 2020). In parallel, the use of optical laser tweezers to manipulate plant MCS in vivo is facilitating the characterization of putative MCS components such as the AtCASP tether identified at ER-Golgi MCSs [START_REF] Osterrieder | Stacks off tracks: a role for the golgin AtCASP in plant endoplasmic reticulum-Golgi apparatus tethering[END_REF] and the mitochondria-associated GTPase, Miro2 at ER-mitochondria contact sites [START_REF] White | Miro2 tethers the ER to mitochondria to promote mitochondrial fusion in tobacco leaf epidermal cells[END_REF]. Plant MCS research is also adopting genetically encoded tools such as synthetic tethers that bridge apposed membranes (e.g. MAPPER-GFP, [START_REF] Lee | Ionic stress enhances ER-PM connectivity via phosphoinositide-associated SYT1 contact site expansion in Arabidopsis[END_REF], or splitfluorescence systems (e.g. split super-folder (sp) GFP proteins, (Li et al., 2020) to visualize MCS contacts. These artificial systems, however, have limited application in functional studies as their expression could induce non-physiological changes in the MCS structure. Additional molecular tools with broad application such as inducible phosphoinositide depletion systems [START_REF] Doumane | Inducible depletion of PI(4,5)P2 by the synthetic iDePP system in Arabidopsis[END_REF], and phosphoinositide fluorescent markers [START_REF] Simon | A multi-colour/multiaffinity marker set to visualize phosphoinositide dynamics in Arabidopsis[END_REF] are being currently adopted for MCS research and represent promising avenues to elucidate the roles of anionic phospholipids in plant MCS function and dynamics.

We forecast that the combination of collaborative research, technical advances, and novel molecular tools in this quickly evolving field will provide breakthroughs that will transcend plant MCS research. In this context, we apologize to those whose important contributions to plant MCS research were not included in this brief review.

Diversity in plasmodesmata form and function

General PD structure Plasmodesmata (PD) evolved multiple times in the plant lineage and are present in some groups of algae and in all land plants [START_REF] Brunkard | The cytosol must flow: intercellular transport through plasmodesmata[END_REF]Azim andBurch-Smith, 2020), 2020). Generally, PD provide continuity of the plasma membranes (PMs) and cytoplasm across cell walls. In land plants and some algae, PD also include a central strand of endoplasmic reticulum (ER) [START_REF] Botha | Plasmodesmatal distribution, structure and frequency in relation to assimilation in C3 and C 4 grasses in southern Africa[END_REF], [START_REF] Ding | Substructure of Freeze-Substituted Plasmodesmata[END_REF][START_REF] Franceschi | Mechanism of Plasmodesmata Formation in Characean Algae in Relation to Evolution of Intercellular Communication in Higher-Plants[END_REF][START_REF] Cook | Comparative ultrastructure of plasmodesmata of Chara and selected bryophytes: toward an elucidation of the evolutionary origin of plant plasmodesmata[END_REF]. The cytoplasmic and membrane connectivity provided by the PD is the route for intercellular trafficking of numerous biomolecules, effectively rendering the plant a symplast. PD are therefore essential for plant growth, development and environmental responses. Some molecules traffic through PD by passive diffusion, and movement depends on the size of the molecules and the trafficking capacity of the pores. Others are targeted to PD using the endomembrane system [START_REF] Spiegelman | A role for the endoplasmic reticulum in the cell-to-cell movement of SHORT-ROOT[END_REF]. A typical cell wall is pervaded by hundreds or thousands of PD that are often clustered into groups (pitfields), and as such the continuity between adjacent cells can be extensive.

PD are nanopores with outer diameters, delimited by the PMs of connected cells, ranging from 25 to 50 nm depending on the tissue and species, and they extend for the length of cell wall thickness.

Thus, much of what is known about PD structure is derived from TEM (Figure 11). The center of PD of land plants is occupied by a central structure called the desmotubule (DT). The DT was observed to be continuous with the cortical ER of connected cells and the DT is now recognized as an intercellular strand of modified ER. The DT diameter is constrained to approximately 15-20 nm [START_REF] Ding | Substructure of Freeze-Substituted Plasmodesmata[END_REF][START_REF] Schulz | Plasmodesmal Widening Accompanies the Short-Term Increase in Symplasmic Phloem Unloading in Pea Root-Tips under Osmotic-Stress[END_REF], and so the desmotubule comprises the most tightly curved biological membranes described to date (Tilsner et al., 2011). The DT does not include a typical ER lumen and instead the space is largely occupied by proteins (Tilney et al., 1991), whose likely function is to enable the tight curvature of the membranes, e.g. the ER tubulating reticulon proteins (Tilsner et al., 2011;[START_REF] Knox | Putting the Squeeze on Plasmodesmata: A Role for Reticulons in Primary Plasmodesmata Formation[END_REF][START_REF] Kriechbaumer | Reticulomics: Protein-Protein Interaction Studies with Two Plasmodesmata-Localized Reticulon Family Proteins Identify Binding Partners Enriched at Plasmodesmata, Endoplasmic Reticulum, and the Plasma Membrane[END_REF]. The DT is tightly connected to the PD PM by structures originally described as spokes [START_REF] Ding | Substructure of Freeze-Substituted Plasmodesmata[END_REF]. The cytosol-filled space between the DT and PM is called the cytoplasmic sleeve or annulus and it is likely the main route for PD trafficking although the spoke proteins divide it into nanochannels 2-3 nm wide.

Analysis of PD isolated from Arabidopsis suspension cell culture identified 1,341 proteins as the putative PD proteome [START_REF] Fernandez-Calvino | Arabidopsis plasmodesmal proteome[END_REF]. Of those, 21% were membrane proteins and included proteins previously identified as PD resident, e.g. PDLP1 (Thomas et al., 2008) and ATBG_papp [START_REF] Levy | A plasmodesmata-associated beta-1,3-glucanase in Arabidopsis[END_REF]. In addition, several GPI-anchored proteins and proteins associated with the secretory pathway were identified. Further refinement of the PD proteome identified multiple C2 domains and transmembrane region proteins (MCTPs) as PD constituents and they have been designated as the likely spokes of PD [START_REF] Brault | Multiple C2 domains and transmembrane region proteins (MCTPs) tether membranes at plasmodesmata[END_REF]. The spokes control spacing between the DT and PM and this distance is correlated with PD developmental states (Nicolas et al., 2017a).

Interestingly, in Arabidopsis roots, PD lacking cytoplasmic sleeves apparently had a higher trafficking capacity than did PD with cytoplasmic sleeves (Nicolas et al., 2017a), raising questions about how trafficking via those PD is achieved. There are a few reports of trafficking through the DT lumen, although the DT membranes appear to provide a surface for cell-to-cell movement [START_REF] Guenoune-Gelbart | Tobacco mosaic virus (TMV) replicase and movement protein function synergistically in facilitating TMV spread by lateral diffusion in the plasmodesmal desmotubule of Nicotiana benthamiana[END_REF]Barton et al., 2011). The DT membranes are important conduits for transport of at least some viruses between cells [START_REF] Guenoune-Gelbart | Tobacco mosaic virus (TMV) replicase and movement protein function synergistically in facilitating TMV spread by lateral diffusion in the plasmodesmal desmotubule of Nicotiana benthamiana[END_REF].

The lipid composition of PD PMs is also distinct from the bulk PM. The PM of PD from Arabidopsis suspension cells is enriched in sterols and sphingolipids with saturated very long chain fatty acids [START_REF] Grison | Specific Membrane Lipid Composition Is Important for Plasmodesmata Function in Arabidopsis[END_REF], consistent with the presence of lipid microdomains akin to lipid rafts in PD [START_REF] Raffaele | Remorin, a solanaceae protein resident in membrane rafts and plasmodesmata, impairs potato virus X movement[END_REF]Tilsner et al., 2013) and GPI-anchored proteins in the PD proteome [START_REF] Fernandez-Calvino | Arabidopsis plasmodesmal proteome[END_REF]. Further biochemical treatments revealed that PD protein composition was dependent on PM lipid composition [START_REF] Grison | Specific Membrane Lipid Composition Is Important for Plasmodesmata Function in Arabidopsis[END_REF]. PD lipid composition is also important for PD function, as changes in lipids affect the ultrastructure and permeability of PD [START_REF] Yan | Sphingolipid biosynthesis modulates plasmodesmal ultrastructure and phloem unloading[END_REF][START_REF] Iswanto | Sphingolipids Modulate Secretion of Glycosylphosphatidylinositol-Anchored Plasmodesmata Proteins and Callose Deposition[END_REF][START_REF] Liu | Phytosphinganine Affects Plasmodesmata Permeability via Facilitating PDLP5-Stimulated Callose Accumulation in Arabidopsis[END_REF]. A modern view of PD combines its unique lipid and protein composition to describe PD as specialized membrane contact sites [START_REF] Brault | Multiple C2 domains and transmembrane region proteins (MCTPs) tether membranes at plasmodesmata[END_REF]Petit et al., 2019;[START_REF] Ishikawa | Structural and functional relationships between plasmodesmata and plant endoplasmic reticulum-plasma membrane contact sites consisting of three synaptotagmins[END_REF].

PD formation and distribution

PD are intrinsic components of the cell walls. Primary PD form at the end of cell division, during cytokinesis, when strands of ER become encased in the developing cell plate. The reticulon proteins, RTNLB3 and 6, and MCTPs are involved in this process [START_REF] Knox | Putting the Squeeze on Plasmodesmata: A Role for Reticulons in Primary Plasmodesmata Formation[END_REF][START_REF] Brault | Multiple C2 domains and transmembrane region proteins (MCTPs) tether membranes at plasmodesmata[END_REF]. The presence of substructures like the DT in newly formed PD is uncertain, as revealed by TEM [START_REF] Ehlers | FRIENDLY regulates mitochondrial distribution, fusion, and quality control in Arabidopsis[END_REF]. Secondary PD form across existing cell walls where cell division is not occurring.

The insertion of these new PD is likely necessary to establish or maintain symplastic connectivity as in graft unions or when cells divide and grow [START_REF] Ehlers | Primary and secondary plasmodesmata: structure, origin, and functioning[END_REF]. Studies in Arabidopsis trichomes suggest that new secondary PD form in close proximity to existing PD, as described in the multiple twinning model [START_REF] Faulkner | Peeking into pit fields: a multiple twinning model of secondary plasmodesmata formation in tobacco[END_REF]. It is proposed that PD divide by fission, although a mechanism for this is unclear.

PD may also be removed from existing cell walls by a still unknown process. Studies on cambial division and vascular differentiation have shown that PD numbers can increase and decrease over the lifespan of a given cell-cell interface, and this would necessarily involve the removal and insertion of PD at a given interface [START_REF] Ehlers | FRIENDLY regulates mitochondrial distribution, fusion, and quality control in Arabidopsis[END_REF][START_REF] Fuchs | Season-associated modifications in symplasmic organization of the cambium in Populus nigra[END_REF]. In other instances, PD can be drastically modified or even truncated to disrupt intercellular trafficking. For example, guard cell initials contain PD, and PD trafficking is critical for guard cell development [START_REF] Guseman | Dysregulation of cell-to-cell connectivity and stomatal patterning by loss-of-function mutation in Arabidopsis chorus (glucan synthaselike 8)[END_REF]. As stomata develop, however, PD are lost from the guard-cells, rendering them symplastically isolated [START_REF] Wille | Ultrastructural and histochemical studies on guard cells[END_REF]. It may be that PD removal is a more common occurrence in plant cell development and differentiation than previously reported. This aspect of PD biology awaits further exploration using advanced imaging approaches.

Structure-function relationships in PD

PD are often depicted as simple linear structures traversing the cell wall but PD structure is much more diversified. PD are often branched, consisting of multiple pores that connect in the vicinity of the cell wall middle lamella (Figure 11). The formation and origins of branched PD are unclear, but they likely arise through modification of existing simple PD [START_REF] Burch-Smith | Plasmodesmata during development: re-examination of the importance of primary, secondary, and branched plasmodesmata structure versus function[END_REF]. This diversity in structure suggests diversity in function. Exemplary studies of PD in tobacco leaves undergoing the sink-source transition demonstrated that simple PD were converted to branched PD contemporaneously with decreased import of fluorescent dye [START_REF] Oparka | Simple, but Not Branched, Plasmodesmata Allow the Nonspecific Trafficking of Proteins in Developing Tobacco Leaves[END_REF][START_REF] Roberts | Dynamic changes in the frequency and architecture of plasmodesmata during the sink-source transition in tobacco leaves[END_REF]. Another common variation of PD form is the dilation of PD pores away from the PD openings or the constriction of PD at their necks (region just below opening). This PD variation seems to correlate with PD maturation [START_REF] Nicolas | Architecture and permeability of post-cytokinesis plasmodesmata lacking cytoplasmic sleeves[END_REF] or with trafficking capacity [START_REF] Ding | Substructure of Freeze-Substituted Plasmodesmata[END_REF].

Mathematical modeling supports that dilation increases trafficking capacity as the cell wall thickens [START_REF] Deinum | From plasmodesma geometry to effective symplasmic permeability through biophysical modelling[END_REF], a correlation previously observed by TEM [START_REF] Nicolas | Architecture and permeability of post-cytokinesis plasmodesmata lacking cytoplasmic sleeves[END_REF]. Another PD form that has a specialized role in trafficking is the 'funnel PD' in sink root tissue [START_REF] Ross-Elliott | Phloem unloading in Arabidopsis roots is convective and regulated by the phloem pole pericycle[END_REF]. These PD have wide openings at the phloem sieve elements that narrows considerably as they cross the cell wall and open on the phloem-pole pericycle creating a 'funnel' shape. The specialized PD shape appears to facilitate the unloading of sucrose in the root phloem and mathematical modelling supports the need for this unusual PD form to allow phloem unloading at physiological sucrose concentrations. Specialized PD forms have also been reported at sites where sugars are loaded into the phloem in source tissues. For example, PD at the phloem parenchymacompanion cell interface in Arabidopsis leaf veins have many openings to the phloem parenchyma but only one to the companion cells [START_REF] Haritatos | Minor vein structure and sugar transport in Arabidopsis thaliana[END_REF]. These distinct PD forms correlate with specialized functions, and raise the possibility of PD sub-functionalization between tissues and even at a given cell-interface. Despite the importance of PD to plants, many aspects of their formation, structure and function remain unknown. Undoubtedly, comprehensive understanding of PD will enable novel approaches to engineering solutions for challenges in plant growth and development.

Acknowledgements:

Our apologies to colleagues whose work we could not cite due to this article's format. Thanks to Dr. Brandon Reagan for critical reading of the manuscript and helpful feedback. This work was supported by the National Science Foundation (MCB1846245).

So much more than bricks and mortar: plant cell walls as dynamic extracellular "organelles"

Much as the skin is the largest organ in the human body, the plant cell wall can be thought of as the largest organelle in the plant cell, even though the wall is not bound by a membrane but instead encases the plasma membrane-delimited protoplast that contains the canonical organelles. Our understanding of cell wall composition, structure, and mechanics has expanded rapidly over the past decade due to advances in high-resolution imaging [START_REF] Zeng | Visualizing chemical functionality in plant cell walls[END_REF][START_REF] Rydahl | Report on the Current Inventory of the Toolbox for Plant Cell Wall Analysis: Proteinaceous and Small Molecular Probes[END_REF][START_REF] Voiniciuc | Monitoring Polysaccharide Dynamics in the Plant Cell Wall[END_REF][START_REF] Zhao | Advances in Imaging Plant Cell Walls[END_REF], biochemical and spectroscopic analyses of wall polymers and their interactions down to single-molecule and nanoscale levels [START_REF] Voxeur | Oligogalacturonide production upon Arabidopsis thaliana-Botrytis cinerea interaction[END_REF][START_REF] Zhao | Solid-state NMR of plant and fungal cell walls: A critical review[END_REF][START_REF] Cai | A solid-state nanopore-based singlemolecule approach for label-free characterization of plant polysaccharides[END_REF], and new computational modeling methods that relate wall mechanics to the deformations, movements, and interactions of individual wall polymers [START_REF] Zhang | Molecular insights into the complex mechanics of plant epidermal cell walls[END_REF]. In contrast to its previous conception as simply "dead wood" that is the fossilized product of polysaccharide secretion by plant cells, the plant cell wall is now starting to be appreciated as a dynamic structure that changes over time and encompasses specialized metabolic processes, including wall polymer polymerization, coalescence, binding/unbinding, cleavage, and re-ligation that facilitate both plant growth and the processing of plant biomass for human use [START_REF] Obro | Enzymatic Modification of Plant Cell Wall Polysaccharides[END_REF].

The intercellular transport of nutrients, secreted peptides, hormones, and other metabolites across the wall [START_REF] Ramakrishna | Polarized transport across root epithelia[END_REF], the discovery of extracellular vesicles that can deliver small RNAs to silence virulence genes in plant pathogens [START_REF] Cai | Plants send small RNAs in extracellular vesicles to fungal pathogen to silence virulence genes[END_REF] and the ability of plants to sense pathogen-generated wall fragments [START_REF] Vaahtera | Cell wall integrity maintenance during plant development and interaction with the environment[END_REF] and other aspects of wall integrity [START_REF] Rui | A wall with integrity: surveillance and maintenance of the plant cell wall under stress[END_REF] together highlight how the apoplast, the extracellular compartment in which the cell wall exists, might also allow for previously unappreciated forms of trafficking and act as a molecular frontier in the interactions between plants and their abiotic and biotic environments.

Cell wall assembly and structure

Consider the plant cell wall as a futuristic building that can grow (and sometimes shrink) and protects and shapes its occupant, in this case the protoplast, and also helps sense and transduce important environmental and mechanical information. Cellulose, the most abundant biopolymer on Earth, forms the "girders" of the cell wall as its primary load-bearing component, and coincidentally has a tensile strength similar to that of steel. Cellulose is extruded directly into the apoplast by multi-subunit Cellulose Synthase Complexes [START_REF] Wilson | The molecular basis of plant cellulose synthase complex organisation and assembly[END_REF], which move through the plasma membrane along trajectories that are likely driven by the force of polymerization and are guided by either cortical microtubules or existing wall patterning [START_REF] Chan | Interaction between Autonomous and Microtubule Guidance Systems Controls Cellulose Synthase Trajectories[END_REF]. The estimated 18 catalytic subunits in each Cellulose Synthase Complex [START_REF] Nixon | Comparative Structural and Computational Analysis Supports Eighteen Cellulose Synthases in the Plant Cellulose Synthesis Complex[END_REF] produce strands of β-1,4-linked glucose that coalesce into cable-like microfibrils that are predicted to have 18-24 chains [START_REF] Yang | A density functional theory study on the shape of the primary cellulose microfibril in plants: effects of C6 exocyclic group conformation and H-bonding[END_REF].

Forming the "cross-beams" and "insulation" between the cellulose "girders" are matrix polysaccharides that include pectins and hemicelluloses. Pectins are acidic polysaccharides that are composed of homogalacturonan, rhamnogalacturonan-I, and rhamnogalacturonan-II domains [START_REF] Anderson | Pectic Polysaccharides in Plants: Structure, Biosynthesis, Functions, and Applications[END_REF], whereas hemicelluloses contain mostly neutral sugars and include xyloglucans, xylans, and mannans [START_REF] Scheller | Hemicelluloses[END_REF]. In growing cells, matrix polysaccharides initially interact with cellulose upon their secretion at the cell surface, following polymerization in the Golgi lumen and post-Golgi trafficking [START_REF] Hoffmann | Subcellular coordination of plant cell wall synthesis[END_REF] (Figure 12). Both pectins and hemicelluloses can associate with the surfaces of cellulose microfibrils, potentially preventing cellulose agglomeration and thus assembling a strong but deformable wall that also contains glycoproteins, enzymes, metabolites, ions, and water [START_REF] Cosgrove | Nanoscale structure, mechanics and growth of epidermal cell walls[END_REF]. In the secondary walls produced by certain cell types, a polyphenolic, hydrophobic compound called lignin is also deposited [START_REF] Dixon | Lignin biosynthesis: old roads revisited and new roads explored[END_REF]. In many cell types, the wall is deposited in layers with differing cellulose orientations, conferring multidirectional resistance to mechanical failure.

Dynamics and functions of plant cell walls

What happens to the strong but flexible wall as a plant cell grows? Atomic force microscopy [START_REF] Zhang | Nanoscale movements of cellulose microfibrils in primary cell walls[END_REF] and coarse-grained modeling [START_REF] Zhang | Molecular insights into the complex mechanics of plant epidermal cell walls[END_REF] indicate that cellulose microfibrils bend, bundle, unbundle, and slide during experimentally imposed or computationally simulated wall deformation, respectively; one remaining question is exactly how these processes occur in the growing cells of living plants, where wall deposition is typically ongoing, matrix polysaccharides also undergo reorganization [START_REF] Anderson | Metabolic click-labeling with a fucose analog reveals pectin delivery, architecture, and dynamics in Arabidopsis cell walls[END_REF], and wall-modifying enzymes can modulate cell growth [START_REF] Xiao | POLYGALACTURONASE INVOLVED IN EXPANSION1 Functions in Cell Elongation and Flower Development in Arabidopsis[END_REF]. Also unclear is the extent to which extracellular ATP and other energetic compounds might be used in wall metabolism, in addition to their functions as signaling molecules [START_REF] Pietrowska-Borek | New Insight into Plant Signaling: Extracellular ATP and Uncommon Nucleotides[END_REF]. Cell walls are highly diverse across plant tissues and taxa [START_REF] Hoffmann | Subcellular coordination of plant cell wall synthesis[END_REF] and allow cells to adopt myriad shapes and perform specialized functions that include nutrient and water absorption (e.g., root epidermal cells), transport (e.g., xylem and phloem), and secretion (e.g., aerial epidermal and nectary cells). Autodegradation of the plant cell wall allows for developmental processes that include organ abscission and pollen dehiscence, and might allow for recycling of some wall components to produce new wall polymers (Barnes and Anderson, 2018).

Overall, the plant cell wall is a fascinating biological environment, one for which we are only beginning to be able to understand well enough to be able to engineer ourselves. (Liang et al., 2018b). FLZ is a dynamin-like protein in which thylakoid fusion is inhibited [START_REF] Gao | FZL, an FZO-like protein in plants, is a determinant of thylakoid and chloroplast morphology[END_REF][START_REF] Findinier | The dynamin-like protein Fzl promotes thylakoid fusion and resistance to light stress in Chlamydomonas reinhardtii[END_REF]. Instead of a stroma-wide network, thylakoids form discrete spirals in the

Acknowledgements:

  Thank you to past and present members of the Zolman Lab for inspiring discussions on peroxisome structure and function. Peroxisome research in the lab is funded by the National Institutes of Health (1R15GM116090-01), the University of Missouri Research Board, and the University of Missouri -St. Louis. My apologies to those colleagues whose contributions could not be included within this short overview.

Figures 1249 1250 Figure 1 .

 12501 Figures 1249

Figure 2 .

 2 Figure 2. The plant ER forms a distinctive network of membranes at the cell cortex. Confocal microscopy image of a tobacco leaf epidermal cell transiently expressing the fluorescent lumenal marker, ER-mCherry (Nelson et al., 2007), which labels the lumen of the bulk ER network. Scale bar = 40 m.

Figure 3 .

 3 Figure 3. Structure and distribution of Golgi stacks in plant cells. (A) TEM micrograph showing a cluster of Golgi stacks in an Arabidopsis root tip cell. Golgi stack (G), trans-Golgi network (TGN), plastids (P), mitochondrion (M), multivesicular body (MVB), and vacuoles (V) are marked. (B) Confocal laser scanning micrograph of Arabidopsis root tip cells expressing a Golgi-localized green fluorescent protein. The cell wall is counterstained. (C) ET slice image of a Golgi stack. Cis-side, trans-side and TGN cisterna are marked. (D) ET model of an Arabidopsis Golgi stack associated with the endoplasmic reticulum (ER). The entire Golgi and TGN are encompassed by a ribosomeribosome excluding matrix (Golgi matrix). Scale bars in A, C, and D: 500 nm. Scale bar in B: 10 m.

Figure 4 .

 4 Figure 4. Plant endosomes. (A) Diagram of plant endosomes and main associated pathways, highlighting the effects on MVE mis-sorting in ESCRT mutants. (B) Tomographic reconstructions of a Golgi, Golgi-associated TGN (GA-TGN), and free/Golgi independent (GI-TGN) in an Arabidopsis embryo cells. (C-D) Confocal images of the MVE-localized RabF2a/RHA1-GFP (C) and the TGNlocalized VHAa1-GFP in Arabidopsis root cells. Scale bar = 200 nm in (B) and 5 mm in (C) and (D).

Figure 6 .

 6 Figure 6. Spatial association of LD with ER and a model of LD biogenesis (A) Enhancedresolution fluorescence imaging of the relationship of ER to LDs in leaf mesophyll cells of N. benthamiana infiltrated with ER marker and stained with LD-specific fluorescent dye BODIPY 493/503. The ER network was visualized as cyan color with the ER-lumen marker protein Kar2-CFP-HDEL and LDs are false-colored as yellow (white arrows). Normally in leaves, small LDs are observed associated with the ER (top row). In this system, LDs were induced to proliferate by expression of lipogenic factors to study LD proteins and their roles in LD formation (second row). Here this process is illustrated by expression in these leaves of the seed transcription factor LEAFY COTYLEDON 2 (LEC2) that promotes storage lipid synthesis and LD formation. Under semi-normal conditions the LD phenotype of tobacco shows few small LD's intimately connected to the ER. The CFP fluorophore was excited with a 405-nm laser with emission spectrum collected between 450 to 500 nm. LDs were visualized by excitation with a 488-nm laser, with emission spectra collected between 500 to 540 nm. Scale bars: 5µm. (B) TEM micrographs showing LDs emerging from the ER. Left to right-freeze-fracture; cryofixation; chemical fixation. Arrows mark ER-LD junctions. For scale, ribosomes on the ER membrane are approximately 20 nm in diameter. Electron micrographs are courtesy of Dr. Eliot Herman, University of Arizona. (C) Diagram illustrating the current, general model for LD biogenesis. Initial LD formation begins with the coalescence of the "lipid lens" within the ER bilayer. Recruitment of various LD associated proteins such as SEIPINs, LDIP, LDAPs, VAP27-1, oleosins (in seeds) and LDIP, which together facilitate the formation and stabilization of nascent LD as it emerges into the cytoplasm.Adapted in part from model presented in[START_REF] Greer | SEIPIN Isoforms Interact with the Membrane-Tethering Protein VAP27-1 for Lipid Droplet Formation[END_REF].

Figure 7 .

 7 Figure 7. Microscopic images of peroxisomal structures in Arabidopsis cells. (A) Microscopic image of 4-day-old Arabidopsis cotyledons expressing mNeonGreen with a membrane peroxisomal targeting signal (mNeonGree-mPTSPEX26; green) and mRuby with a matrix-bound peroxisomal targeting signal (mRuby3-PTS1; magenta) show the presence of ILVs in peroxisomes. The close-up image highlights the variable sizes of the vesicles. (B) Separate images of the membrane (green) and matrix (magenta) fluorescent molecules highlights the different substructures within the peroxisome, including ILVs with (yellow) or without (blue) matrix proteins and a separate area with denser membrane accumulation. Images in parts A and B are from Wright and Bartel (2020). (C) ET slice image of a young root cell detailing a peroxisome (P), lipid bodies (*), and other organelles in its proximity. Scale bar: 500 nm.

Figure 8 .

 8 Figure 8. Microscopy imaging of plant mitochondria dynamics. (A) Five consecutive frames showing mitochondria fission in a tobacco BY-2 cell. The mitochondria are stained red and dynaminrelated protein 3A is labeled with GFP (Arimura 2018). (B) Fusion of mitochondria in an onion bulb epidermal cell. The cell contains thousands of mitochondria. The mitochondria on the left and right sides of the cell were labeled green and red, respectively. The photos show the movement and mixing of the mitochondria after of 10 minutes (upper), one hour (middle), and two hours (bottom). Yellow mitochondria are the result of fusion between green and red mitochondria. (C) Progression of mitophagy in an Arabidopsis cell, in which the mitochondria were stained with MitoTracker Red and autophagosomes were visualized by expression of YFP-ATG8e. The autophagosome on the right (arrowhead) is shown engulfing a mitochondrion over a 300 s interval (Ma et al., 2021). (D) An apparent mitochondrial outer-membrane derived vesicle (MDV) (arrow) in an Arabidopsis cell. At right is a mitochondrion whose outer-membrane was stained with ELM1-GFP and whose matrix was stained with RFP. The MDV has only the outer membranes and no matrix (Yamashita et al., 2016). E. Heterogeneity of DNA contents in mitochondria. Mitochondria are stained red by MitoTracker Red and DNA stained by SYBR Green I. Green signals in red are shown in yellow. So red particles with yellow dots mean mitochondria having DNA, and red mitochondria without yellow mean having no DNA(Arimura et al., 2004). F. ET image of a mitochondrion in an Arabidopsis root meristematic cell. Scale bars.A, C, and D, 2μm; B, 40μm; E, 1μm; F, 500nm

Figure 9 .

 9 Figure 9. Chloroplast morphogenesis is a highly regulated process. (A) ET slice image of a normal-sized wild-type (WT) chloroplast with typical thylakoid differentiation into stacked (grana) and unstacked domains. (B) 3D model based on the chloroplast in A. (C) ET slice image of an oversized chloroplast (compare scale bars) with aberrant thylakoid membrane organization in a flz mutant

  mutant. (D) 3D model based on the chloroplast in C. Scale bars: 500 nm.

Figure 10 .

 10 Figure 10. Examples of Membrane Contact Sites in Plants. (A-B) ET slice images showing different MCS present in plant cells. an ER-PM contact site (A) and an ER-mitochondrion contact site (B) are shown. Arrowheads mark plasmodesmata. CW: cell wall, M: mitochondrion. Scale bars: 500 nm. (C-D) Co-localization of SYT1-GFP and VAP27-1-YFP with ER markers (RFP-HDEL or GFP-HDEL) highlights the presence of spatially separated and functionally distinct ER-PM MCS within the cortical ER. (E) Co-expression of NET3C, IQD2 and VAP27-1 highlights the interaction of plant ER-PM MCS with the cortical cytoskeleton. (F) The intermembrane distances at MCS are below the light diffraction limit which limits the information provided by conventional confocal microscopes (Laser Scanning / Spinning Disc), and requires the use of super-resolution techniques (TIRF /SIM). (G) Advances in electron tomography techniques are enabling accurate 3D reconstructions of PD MCS. In current functional models, the cytosolic space between the ER and the PM inside PD serves as a trafficking conduit for mobile molecules and the adjustment of its width is believed to regulate their flow rate, effectively controlling inter-cellular trafficking.

Figure 11 .

 11 Figure 11. Structure of branched PD in Arabidopsis tissue revealed by ET. (A) Representative four individual frames from a tomogram (1/4 -4/4). While the PM is readily visible in these images the desmotubule is difficult to discern. Central cavities are found in the vicinity of the middle lamella. (B) 3D models of PD generated by tracing the inner (yellow) and outer (blue) leaflets of the PM in the tomogram in A. The PD on the left consists of two pores in Cell 2 and one in Cell 1. The PD on the left has two openings to Cell 2 but three to Cell 1. Ribosomes (red) are shown for scale.

Figure 12 .

 12 Figure 12. Cellulose labeled with Pontamine Fast Scarlet 4B (S4B, magenta) and newly synthesized pectin labeled with fucose-alkyne and Alexa488-azide (green) in epidermal cells of the root differentiation zone in 5-day-old Arabidopsis seedling. Note oblique, punctate labeling of Alexa488 signal and predominantly longitudinal labeling of S4B signal, and variation in intensity of Alexa488 signal between different cells.

Figure 5. The plant vacuole (A)

Acknowledgements:

We apologize to those colleagues whose work was not included in this review due to space limitations. This work was funded primarily by the National Institutes of Health (GM136637), National Science Foundation (MCB1727362), the Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, US Department of Energy (award number DE-FG02-91ER20021), and AgBioResearch (MICL02598) to FB. Acknowledgements: Sincere apologies to those whose work on endosomal trafficking could not be discussed due to the brevity of this overview. Research on endosomal trafficking in the Otegui Lab is supported by grant NSF MCB 2114603 to MSO. Acknowledgements: Sincere apologies to those whose important work on LDs could not be cited in this brief overview. Special thanks to Payton Whitehead for acquiring confocal images and for Acknowledgements: This work was supported by the JSPS (19KK0391, 19H02927 & 20H05680) to SA. Acknowledgements: The nature of the glossary format requires a highly selective literature survey. I sincerely apologize to my colleagues who were not cited. Research in the Kirchhoff lab is mainly supported by grants from NSF (MCB-BSF-1953570) and DOE (DE-SC0017160).

Acknowledgments: The authors would like to thank the different funding agencies supporting their basic research projects on plant membrane contact sites. We apologize to colleagues whose important contributions to the field were not featured due to length constraints.

Acknowledgements

: Thanks to members of the Anderson Lab and the Center for Lignocellulose Structure and Formation for inspiring discussions, and apologies to those whose important contributions could not be cited. This work was supported as part of The Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Basic Energy Sciences under Award # DE-SC0001090 to CTA.

This work was supported by the USDA National Institute of Food and Agriculture (HATCH project CA-B-PLB-0243-H), National Science Foundation (MCB-2049931), and start-up funds from the University of California Berkeley and the Innovative Genomics Institute (to YG).

We apologize to colleagues whose work was not included due to the limit in article length.