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Keywords:

  • ER;
  • Golgi;
  • reticulons;
  • Yop1;
  • RHD3

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

The endoplasmic reticulum (ER) is an interconnected network comprised of ribosome-studded sheets and smooth tubules. The ER plays crucial roles in the biosynthesis and transport of proteins and lipids, and in calcium (Ca2+) regulation in compartmentalized eukaryotic cells including plant cells. To support its well-segregated functions, the shape of the ER undergoes notable changes in response to both developmental cues and outside influences. In this review, we will discuss recent findings on molecular mechanisms underlying the unique morphology and dynamics of the ER, and the importance of the interconnected ER network in cell polarity. In animal and yeast cells, two family proteins, the reticulons and DP1/Yop1, are required for shaping high-curvature ER tubules, while members of the atlastin family of dynamin-like GTPases are involved in the fusion of ER tubules to make an interconnected ER network. In plant cells, recent data also indicate that the reticulons are involved in shaping ER tubules, while RHD3, a plant member of the atlastin GTPases, is required for the generation of an interconnected ER network. We will also summarize the current knowledge on how the ER interacts with other membrane-bound organelles, with a focus on how the ER and Golgi interplay in plant cells.

  • image

[ Huanquan Zheng (Corresponding author)]


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

The endoplasmic reticulum (ER) is an interconnected network of sheets and tubules that stretches throughout the cytoplasm. The ER is the largest membrane-bound organelle in eukaryotic cells. It has a number of important roles including the synthesis, folding, modification, and trafficking of both soluble and membrane proteins destined to the secretory and endocytic compartments (Shibata et al. 2010), the biosynthesis and distribution of phospholipids and steroids, and the detoxification of drugs and poisons (Meusser et al. 2005). The ER also plays a crucial role in the regulation of calcium (Ca2+) concentration through the sequestration and release of Ca2+ (Hong et al. 1999; Vandecaetsbeek et al. 2011). In addition, the ER is also frequently associated with other organelles including the plasma membrane, peroxisomes, mitochondria, Golgi, and in plant cells, chloroplasts. Reflecting these fundamental roles that the ER plays, the interconnected structure of sheets and tubules of the ER is evolutionarily conserved across animal and plant lineages (Figure 1A) (Sparkes et al. 2009a; Shibata et al. 2010; Zheng and Chen 2011). The ER sheets often have ribosomes attached to them, and are thus closely correlated to the rough ER (Meusser et al. 2005) responsible for the biosynthesis of membrane and secretory proteins. The long ER tubules are mainly ribosome-free, and are therefore closely correlated to the smooth ER (Meusser et al. 2005). They are involved in the synthesis and delivery of lipids, in detoxification, and in Ca2+ regulation.

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Figure 1. Morphology of the endoplasmic reticulum (ER) highlighted by GFP-HDEL.  (A) A spinning disc confocal image of a mature Arabidopsis thaliana leaf epidermal cell expressing GFP-HDEL. Note both sheet-like (arrowheads) and tubule-like (arrows) ER domains are visible. Scale bar = 5 μm.  (B–C) Schematic diagram showing sheet-like ER (B) and tubule-like ER (A). The curvature-stabilizing protein reticulons (blue colored) are enriched in the edges of the sheet (B) and in highly bent tubules (C). In contrast, the sheet-inducing protein Climp63 (red colored) is only found in the sheet of the ER.

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The ER was first described by electron microscopists in the 1950s (Porter et al. 1945). Befitting the different roles that the ER plays, the morphology of the ER is remarkably different across different cell types. For instance, during cell division in animal cells (Lu et al. 2009), and perhaps also in plant cells (Gupton et al. 2006; Sheahan et al. 2007), the ER undergoes a prominent tubule-to-sheet shape transition. In developing plant cells such as root tip cells (Ridge et al. 1999) and growing root hairs (Figure 2A), the ER is more sheet-like, whereas in mature plant cells, tubular ER is more commonly found (Figure 2B; Ridge et al. 1999). In addition, the ER sheets and tubules undergo notable transitions to each other in response to developmental (Ridge et al. 1999) or environmental cues (Quader et al. 1989; Takemoto et al. 2003). The ER can also dynamically change its polygonal network by changing branching patterns and moving network nodes (Sparkes et al. 2009a).

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Figure 2. Morphological difference in the endoplasmic reticulum (ER) in a growing and elongated root hair cell.  (A) The morphology of the ER in a growing root hair cell. Note that the condensed sheet-like ER with small membrane-free areas is seen in the tip region, but the ER in the region further back is more tubule-like.  (B) The morphology of the ER in an elongated root hair cell. Note that ER with open reticulum is seen throughout the root hair cell.  Scale bar in (B) for (A, B)= 20 μm.

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Clearly, the morphology of the ER is actively shaped, and undergoes continuous rearrangement to properly execute its function. It is implicit that mechanisms must exist in shaping the unique morphology of the ER and in regulating the dynamics of the ER. In this review, we will focus on new understandings of how an interconnected ER network is formed, and its interactions with other membrane-bound organelles, with a focus on plant cells. We will also discuss the possible roles of the interconnected ER in cell polarity.

Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

As previously mentioned, a typical ER network often appears as an extended network of interconnected tubules and sheets. It has been recently shown that two families of ER membrane proteins, the reticulons and DP1/Yop1, are able to shape ER tubules in yeast and mammalian cells (Voeltz et al. 2006) as well as in proteoliposomes (Hu et al. 2008). The reticulons and the DP1/Yop1 proteins have a conserved reticulon homology domain that contains transmembrane domains (Shibata et al. 2008). This membrane-embedded segment is responsible for the localization of the reticulons and DP1/Yop1 to the tubular ER, where it forms hairpins (Zurek et al. 2011) and is organized into a W-shaped structure (Figure 1B, C) in the ER lipid bilayer (Shibata et al. 2008). It is assumed that the hairpin transmembrane domains could expand the area of the outer leaflet relative to the inner leaflet of the lipid bilayer, to generate the membrane curvature found in an ER tubule (Shibata et al. 2008; Tolley et al. 2010). Although reticulons and DP1/Yop1 do not share any primary sequence homology (Voeltz et al. 2006), they interact with each other in the formation of ER tubules (Shibata et al. 2008).

The Arabidopsis genome contains at least 21 reticulon homology domain proteins, termed reticulon-like proteins B1–21 (RTNLB1–21) (Sparkes et al. 2010). The overexpression of RTNLB13 reduces the lumenal diameter of ER tubules (Tolley et al. 2008), suggesting that the membrane curvature in ER tubules is increased. A topology prediction based on TOPCONs (Bernsel et al. 2008) suggested that the transmembrane domains in Arabidopsis RTNLB proteins, which have been shown to be important for plant reticulons to shape ER tubules in vivo (Tolley et al. 2010), may be organized into a W-shaped structure (Sparkes et al. 2010). In addition to RTNLB13, the ER location, topology, and ER membrane-shaping abilities of RTNLB1–4 have also been experimentally confirmed (Sparkes et al. 2010). In Arabidopsis, homologs of DP1/Yop1 are identified as HVA22, whose expression can be induced by abscisic acid (ABA) (Chen et al. 2009). The Arabidopsis genome contains at least 11 HVA22 proteins (Chen et al. 2009). However, their role in the formation of ER tubules has not been demonstrated. When HVA22d is depleted by RNAi, autophagy is enhanced (Chen et al. 2009). It is known that an ER defect often invokes autophagy (Yorimitsu et al. 2006). It is possible that the enhanced autophagy observed in HVA22d-depleted cells is in fact a result of altered ER.

While the ER tubules are shaped by the reticulons and DP1/Yop1, a recent study by Shibata et al. (2010) indicated that the formation of sheet-like ER is related to the degree of the generation of membrane curvature in the ER. The reticulons and DP1/Yop1, which induce membrane curvature, are highly concentrated in sheet edges and tubules (Figure 1B, C); however, the formation of sheet-like ER requires an accumulation of coiled-coil membrane proteins in proliferated ER sheets, of which Climp63 serves as a “lumenal ER spacer” (Figure 1B). In the ER membrane, Climp63 and reticulons undergo a “tug-of-war” that in turn determines the ratio of sheets to tubules (Shibata et al. 2010). In Arabidopsis, however, no sequence homolog of Climp63 exists.

Making an Interconnected Network: The Role of Atlastin-1/RHD3

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

Shaping the ER membrane into either tubules or sheets is not sufficient to explain how an interconnected ER network is often formed in eukaryotic cells. Ultimately, different ER tubules need to be connected. It has been indicated that continuous homotypic fusion of ER tubules is required for the formation of an interconnected ER network (Dreier and Rapoport 2000). In mammals, several proteins have been shown to have an essential role in homotypic ER membrane fusion (Latterich et al. 1995; Vedrenne and Hauri 2006), but more recent work has highlighted the important role of the atlastin family of dynamin-like large GTPases, including Sey1 in yeasts and Atlastin-1 in animal cells, in the homotypic fusion of ER tubules (Hu et al. 2009; Orso et al. 2009; Anwar et al. 2012). Interestingly, in plant cells, RHD3, a plant member of the atlastin family of dynamin-like GTPases (Hu et al. 2009; Stefano et al. 2012), has been shown to play an important role in maintenance of an interconnected ER network (Chen et al. 2011). Similar to overexpression of Atlastin in Drosophila (Orso et al. 2009) and Atlastin-1 in COS-7 cells (Hu et al. 2009), overexpression of wild type RHD3 can also make more sheet-like ER (Zheng and Chen 2011), so it is possible that RHD3 in plant cells is also invovled in the fusion of ER tubules.

All atlastin proteins harbor an N-terminal GTP-binding domain, two closely spaced transmembrane domains near the C-terminus, and a middle coiled-coil domain that links the GTP and transmembrane domains (Figure 3A, B; Hu et al. 2009; Stefano et al. 2012). They are localized predominantly to the tubular ER (Hu et al. 2009; Orso et al. 2009; Chen et al. 2011; Anwar et al. 2012; Stefano et al. 2012). In plant cells, mutations in RHD3 or expression of either GTP- or GDP-locked RHD3 result in a less-branched ER (Figure 3C; Zheng et al. 2004; Chen et al. 2011). In Arabidopsis, there are three closely related RHD3s: RHD3, RHD3-like1 and RHD3-like2 (Chen et al. 2011). While both RHD3 and RHD3-like2 are ubiquitously expressed, RHD3-like1 is pollen-specific (Hu et al. 2003). However, it appears that they are functional isoforms, as the expression of RHD3-like2 can rescue the less-branched ER phenotype in cells expressing either GTP- or GDP-locked RHD3 (Chen et al. 2011).

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Figure 3. Predicted structure of RHD3 and the morphology of the endoplasmic reticulum (ER) in rhd3-1.  (A, B) The predicted primary structure (A) and membrane organization (B) of RHD3. A RHD3 molecule is shown in fuchsia-green-blue combination in a lipid bilayer (grey colored). The green ellipse represents the middle domain, while the fuchsia and blue boxes on each side represent the GTPase domain and the C-terminal transmembrane domains, respectively.  (C) A spinning disc confocal image of a leaf epidermal cell of rhd3-1 expressing GFP-HDEL. Note the less branched, unfused ER tubules that run parallel (arrowheads). Scale bar = 10 μm.

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Recent studies on the structure of human Atlastin-1 in the presence/absence of GTP revealed that Atlastin-1 has two conformations: a nucleotide-free “pre-fusion” conformation, and a GTP hydrolysis-stimulated “post-fusion” conformation (Bian et al. 2011; Byrnes and Sondermann 2011). It was proposed that, when two Atlastin-1 molecules in the apposing membranes are in proximity, they undergo a GTP-dependent dimerization to tether the membranes. Afterward, the GTP hydrolysis will trigger a conformational change that pulls the membranes together. The energy released during this process is believed to be used to promote lipid mixing and membrane fusion (Moss et al. 2011). It would be interesting to further examine if RHD3 is also involved in the fusion of ER tubules, and, if it is, whether or not a similar working mode applies to RHD3.

In mammalian cells, Atlastin-1 physically interacts with the reticulons and DP1/Yop1 (Hu et al. 2009) as well as Spastin, a microtubule severing factor (Park et al. 2010). However, the functional significance of these interactions in the formation of the interconnected ER network is still unclear. It is highly unlikely that RHD3 works alone in the generation and maintenance of an interconnected ER network in plant cells. When HVA22d and RHD3 are co-expressed, they co-localized to ER tubules and mark numerous ER punctae (Chen et al. 2011). These results suggest that RHD3 may work together with other reticulons and DP1/Yop1 in concert to establish ER shape and to control its dynamics in plant cells.

Moving the ER in Plant Cells: The Roles of the Actomyosin System

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

Although the shape of tubular ER and the generation of an interconnected ER network can occur in the absence of the cytoskeleton (Dreier and Rapoport 2000), in living cells, many ER tubules are formed on the basis of their physical linkage with the cytoskeleton. Studies in animal cells have revealed that, to guide ER formation and remodeling, the microtubule cytoskeleton is primarily used in non-dividing cells, and during mitosis, cells may switch to actin-mediated movement of the ER (Wollert et al. 2002). Two distinct mechanisms, molecular motor model and the tip attachment complex (TAC) model, have been proposed. Microtubule motors kinesin and dynein are believed to be able to pull out ER tubules from a membrane reservoir along microtubules to the cell periphery and the center of the cell, respectively (Wozniak et al. 2009). In the TAC model, an ER tubule is being attached to the plus end of the growing microtubule, and therefore grows with its microtubule partner. The integral ER membrane protein STIM1 has been proposed to interact with the microtubule plus end-binding protein EB1 to move ER tubules towards the plasma membrane (Grigoriev et al. 2008). However, as we will mention below, STIM1 only moves ER tubules to the plasma membrane upon Ca2+ depletion from the ER (Grigoriev et al. 2008).

It has also been generally accepted that ER formation and movement in higher plant cells is actin-dependent (Sparkes et al. 2009a; Yokota et al. 2011), although in dividing plant cells, perhaps also at the onset of cell elongation, ER movement may be microtubule-dependent (Sheahan et al. 2007; Foissner et al. 2009; Sparkes et al. 2009a). It is assumed that in plant cells, myosin motors provide the major motive forces for the formation and remodeling of the ER. The Arabidopsis genome contains at least 17 myosin genes (Peremyslov et al. 2010; Ueda et al. 2010) which can be classified into two classes: myosin VIII with 4 members, and myosin XI with 13 members (Peremyslov et al. 2010). Recent studies through overexpression of truncated variants of myosin XI-K lacking the myosin head domain, or immunodepletion of myosin XI-2, indicate that myosin XI proteins are involved not only in the motility of the ER in the cytoplasm, but also in the tubule growth of the ER (Sparkes et al. 2009a; Yokota et al. 2011). It was thus proposed that, in plant cells, ER tubule growth and retraction is driven by actin dynamics and myosin activity. ER tubules may be attached on anchor/growth sites and move along actin filaments, mainly through a myosin such as myosin XI-K, but also via actin polymerization (Sparkes et al. 2009a). However, the mechanisms required for an ER tubule to attach to an actin filament are unknown.

Association of the ER with Other Organelles

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

ER-PM junction

As a unique network-shaped organelle, the ER is not only morphologically dynamic, but is also frequently associated with various organelles. One such association is the ER and plasma membrane (PM) connection, called the ER-PM junction (Figure 4A-a). In mammalian cells, the existence of the connection between the ER and the PM was first reported by Dr. Palade in muscle cells (Porter and Palade 1957). Although the structural components of ER-PM junctions are still largely unknown, STIM1, an ER membrane protein, has been found to be enriched in the ER-PM junctions upon Ca2+ depletion from the ER (Grigoriev et al. 2008). The junction is also enriched in negatively-charged PIP2, PI(3,4,5)P3, and possibly phosphatidylserine lipids (Carrasco and Meyer 2011). In mammalian cells, many researchers considered the ER-PM connection to be a unique feature of some specialized cells, providing a direct communication link between the ER and the PM in Ca2+ signaling and lipid transfer (Pichler et al. 2001; Liou et al. 2005). In plant cells, the ER has been frequently observed to be associated with the plasma membrane (Hepler et al. 1990), but there have been no studies on the functional significance of such an association. In mature plant cells, the cortical ER frequently goes through cell walls in structures known as plasmodesmata (Figure 4A-b), where the ER tubule is closely linked with the PM (Lucas et al. 2009). Perhaps plasmodesmata should be considered a specialized ER-PM junction whose major function is to facilitate cell-to-cell communication in plant cells. However, little is known about how the ER is linked with the PM in plasmodesmata.

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Figure 4. Schematic diagram showing association of the endoplasmic reticulum (ER) with other membrane-bound organelles.  Association of the ER with the plasma membrane (A-a and A-b), peroxisomes (B), mitochondria (C), chloroplasts (D) and Golgi (E). pER = peroxisomal ER; ERPIC = ER-peroxisome intermediate compartment; ERMES = ER-mitochondria encounter structure; PLAM = Plastid associated membranes; ERES = ER exit site.

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ER-Peroxisome association

In plant as well as in animal cells, various forms of peroxisomes have been observed to associate frequently with the ER (Huang et al. 1983; Kunau 1998; Sinclair et al. 2009). Peroxisomes were therefore thought to be formed from a specialized ER subdomain called the peroxisomal ER (pER) (Figure 4B) (Beevers 1979), whereby ER-derived membrane carriers fuse homotypically to create a new peroxisome (Lam et al. 2010). However, some proteins required for peroxisome biogenesis, for example, the Arabidopsis peroxins AtPEX2 and AtPEX10, were found to be made on free ribosomes and to be targeted directly to peroxisomes (Sparkes et al. 2005). Thus, it was proposed that peroxisomes are formed by the multiplication-by-division of preexisting peroxisomes, with their size increase from the posttranslational import of protein constituents from the cytosol (Lazarow and Fujiki 1985). But this model could not reconcile the observations that, in yeasts, mammals, and plants, at least some peroxisome membrane proteins are sorted indirectly to peroxisomes by way of the ER, and that in yeast cells, perixosomes can be formed de novo from the ER (Hu et al. 2012; Huber et al. 2012). Indeed, the biogenetic relationship between the ER and peroxisomes has been highly debated in recently years (Hu et al. 2012). The current working model for peroxisome biogenesis is that, a peroxisome can undergo an autonomous replication to generate daughter peroxisomes, but the maturation of the daughter peroxisomes requires the fusion of small preperoxisomal membrane vesicles derived from the ER, and perhaps the ER-peroxisome intermediate compartment (ERPIC) (Figure 4B) (Mullen and Trelease 2006).

ER-Mitochondrion association

The ER and the mitochondria are other organelles that have a privileged association in animal cells. The membranes of both organelles can often be observed in close apposition by electron microscopy (Csordas et al. 2006). In mammalian cells, the isolation of subcellular mitochondria fractions usually contains contaminating ER membranes, and these contaminating ER membranes have a molecular composition that is slightly different from that of the general ER (Vance 1990). One role for the ER-mitochondrion connection (ER-mitochondrion encounter structure/ERMES, Figure 4C) is to facilitate lipid exchange between both organelles (Daum and Vance 1997). The ERMES may also promote Ca2+ exchange between the ER and the mitochondria (Rizzuto et al. 1993). In plant cells, there has been little research on the ER-Mitochondrion connection, but the association between the ER and the mitochondria is also observed in plant cells (Morre et al. 1971; Wang et al. 2010). Indeed, genetic data in Arabidopsis also indicates that specific phospholipid transfer from the ER to the mitochondria exists in plants (Nerlich et al. 2007).

ER-Chloroplast association

Chloroplasts are plant-specific organelles that are responsible for trapping solar energy into carbon-rich compounds. Association between the ER and chloroplasts as well as stromules that connect different chloroplasts has been regularly reported (McLean et al. 1988; Schattat et al. 2011). A role for a connection between the ER and the chloroplast membranes (Plastid associated membranes/PLAMs, Figure 4D) in the biogenesis of the thylakoid and the development of chloroplasts has been proposed (Kjellberg et al. 2000; Awai et al. 2006). Indeed, it is well established that the development of chroloplasts requires lipid precursors (for example, diacylglycerol), fatty acyl CoAs (Kjellberg et al. 2000) that are synthesized in the ER (Zheng et al. 2005). The direct ER-to-chloroplast association has been thought to be one of the possible transport routes for lipid transfer from the ER to the chloroplast membrane (Xu et al. 2008). It was found that, TGD4, an ER membrane protein, plays an important role in lipid transfer from the ER to the chloroplasts (Xu et al. 2008).

Interplay Between the ER and Golgi in Plant Cells

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

In the secretory pathway, the Golgi apparatus is an organelle that is functionally (and in plant cells, physically) associated with the ER. The Golgi apparatus is the central organelle that links the ER and the plasma membrane or lysosome/vacuole in the secretory pathway. Proteins received from the ER are usually modified and sorted in the Golgi. In mammalian cells, the Golgi is clustered near the microtubule organization center (Scales et al. 1997). In contrast to this, in plant cells, the Golgi stacks appear as scattered units that are distributed throughout the cytoplasm (Boevink et al. 1998; Nebenfuhr et al. 1999). They move rapidly along the ER (daSilva et al. 2004; Sparkes et al. 2009b). Live cell imaging based on the expression of fluorescent protein markers has indicated that Golgi bodies are intimately associated with the ER (daSilva et al. 2004). When individual Golgi bodies are pulled around the cortical cytoplasm by laser tweezers, Golgi bodies can drag the tubules of the ER behind them (Sparkes et al. 2009b), suggesting that at least a population of the Golgi stacks in plant cells are physically linked with the ER.

It is a well established fact that in mammalian and yeast cells, COPII/I vesicles are involved in protein transport between the ER and the Golgi (Scales et al. 1997). However, the evidence for COPII/I vesicle mediated transport in plants is still limited, though in Arabidopsis, genes required for the initiation, formation, transport, tethering and fusion of COPII/I vesicles exist (Robinson et al. 2007; Bassham et al. 2008). Based on observations that the motility of Golgi stacks often alternate between a random slow “wiggling” motion and fast movement along a linear track (Nebenfuhr et al. 1999), a “stop-and-go” model for protein transport from the ER to the Golgi was established. It is assumed that ER vesicles are generated from ER exit sites (ERESs, Figure 4E). When a stop signal is sent out from an ERES, the nearby Golgi stack disassociates from the actin track and displays a random “wiggling” motion around the ERES to pick up the vesicles. Once the vesicles are received, the Golgi resumes its fast movement along the cytoskeleton to optimize the membrane trafficking afterwards (Nebenfuhr et al. 1999). However, in the presence of dominant-negative RHD3 and in rhd3-7, the ER becomes less branched, and the fast directed movement of Golgi bodies is affected (Chen et al. 2011; Stefano et al. 2012); however, the ER-Golgi trafficking is not inhibited. Therefore, Golgi motility is not critical to protein transport from the ER to the Golgi. This conclusion is also supported by fluorescence recovery after photobleaching studies on fluorescent Golgi markers in cells treated with actin depolymerizing agents (Brandizzi et al. 2002). In plant cells, the ERES and the Golgi stacks often move along the ER together, and they may therefore act as a single functional unit (daSilva et al. 2004). Furthermore, some regions of the ER are known to be physically linked to the Golgi cisternae by tubules (Sparkes et al. 2009b). Thus, it is possible that there is a continuous flow of cargo from the ER to the Golgi stacks via direct tubular connections (Sparkes et al. 2009b).

As mentioned above, in Arabidopsis, genes required for the initiation, formation, transport, tethering and fusion of COPII/I vesicles exist. Interestingly, many of these trafficking-related genes have expanded during evolution quite differently from animal lineages (Bassham et al. 2008). In animal and yeast cells, Rab1/Ypt1 GTPases are known to regulate the tethering of COPII vesicles in ER-to-Golgi trafficking (Allan et al. 2000). Rab1-related proteins in higher plants are divided into two subclasses: Rab-D1 and Rab-D2. RAB-D2a has been shown to regulate ER-to-Golgi protein transport (Batoko et al. 2000). Although both RAB-D1 and RAB-D2a target to the same punctate structures that can be marked by Golgi as well as trans-Golgi-network markers, they could act at distinct biosynthetic trafficking pathways, since the dominant negative effect in cells expressing either RAB-D1(N121I) or RAB-D2a(N121I) can only be restored by expression of the respective wild-type form of the protein (Pinheiro et al. 2009).

Another good example of independent elaboration of conserved trafficking-related genes in plants is the p24 proteins, a family of small (24 kDa) type I integral membrane proteins. In yeast and animal systems, p24 proteins have four subfamilies: α, δ, β and γ (Strating and Martens 2009). They serve as cargo receptors for COPII vesicles (Schimmoller et al. 1995), and are required for the proper formation of COPI vesicles (Gommel et al. 2001), as well as being involved in the furnishing of the various subdomains of the ER-Golgi interface for selective transport of proteins between the ER and the Golgi (Strating and Martens 2009). In Arabidopsis, there are at least 11 putative p24 proteins (Chen et al. 2012), but characterization of these proteins began only recently (Langhans et al. 2008; Chen et al. 2012; Montesinos et al. 2012). Unlike in animal cells, the 11 Arabidopsis p24 proteins fall into only two subfamilies: δ with 9 members, and β with only 2 members (Chen et al. 2012). While the p24δ proteins are largely localized to the ER, and are sometimes also present in Golgi, the two p24β proteins reside largely in Golgi (Chen et al. 2012; Montesinos et al. 2012). There is also a coupled transport between p24δ and p24β at the ER-Golgi interface (Chen et al. 2012; Montesinos et al. 2012). Because of the different organization of the ER-Golgi interface in plant cells, it would be illustrative to exam how the transport of different cargo proteins is regulated by different p24 proteins in plant cells.

Future Directions

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

The actively-shaped ER network supports segregated functions of the ER. Despite significant progress in research on the structure, dynamics and function of the ER in plant cells in recent years, many questions remain. The reticulons and DP1/Yop1 have expanded spectacularly during plant evolution (Chen et al. 2009; Sparkes et al. 2010), it is likely that they may play some plant-specific roles. Does RHD3 also mediate the fusion of two lipid bilayers? If so, how is it involved? RHD3 may potentially interact with an array of proteins. As a first step, the interaction between RHD3, RTNLB, and HVA22 proteins should be examined.

As an extended network of interconnected tubules and sheets, the shape of the ER often undergoes drastic changes in response to both developmental cues and outside influences (Quader et al. 1989; Ridge et al. 1999; Takemoto et al. 2003). Thus, the action of the reticulons and RHD3 must be tightly regulated. It seems that proteins that act upstream of these proteins should hold the key to modulating the structure, dynamics, and function of the ER. In this regard, it is interesting to note that rhd2-1 is epistatic to rhd3-1 (Schiefelbein and Somerville 1990). RHD2 is an NADPH oxidase responsible for localized production of ROS in root hairs (Foreman et al. 2003). RHD2-derived ROS is known to stimulate Ca2+ influx into the cytoplasm (Foreman et al. 2003). Whether or not ER organization in plants is regulated by Ca2+ is unknown, but the application of Ca2+ in mammalian cells induces ER restructuring (Subramanian and Meyer 1997). Therefore, it would be interesting to examine if and how RHD2 is involved in regulating the function of RHD3.

RHD3 is a protein isolated in a genetic screen for mutants defective in root hair development (Wang et al. 1997). Similar to short and wavy axons of corticospinal neurons found in humans [a medical condition called hereditary spastic paraplegia (HSP)] whose Atlastin-1 is improperly altered, in rhd3 mutants, the root hairs are short and wavy (Wang et al. 1997). Clearly, root hairs of Arabidopsis rhd3 and Atlastin-defective HSPs are remarkable examples of the importance of ER organization in polarized cell growth. The ER is the port of entry for all membrane proteins and secretory proteins. Interestingly, in cells expressing dominant negative RHD3, the bulk flow of protein secretion is not prevented (Chen et al. 2011). However, Golgi stacks tend to aggregate, and many of them undergo a slow wiggling motion along the unbranched ER tubules (Chen et al. 2011). In root hair tip growth, cell wall materials need to be targeted to the growing dome (Vincent et al. 2005; Zhang et al. 2010). The agglomeration of Golgi stacks is known to affect the positioning, but not the transport of CESA6 (a subunit of the plasma membrane localized cellulose synthase) to the plasma membrane (Crowell et al. 2009). Thus, it is possible that some proteins (for example, CSLD3, a root hair tip-localized cellulose synthase-like protein required for cellulose synthesis in apical plasma membranes of growing root hairs) (Park et al. 2011) may be misplaced in rhd3.

In the apical dome of growing root hairs, tip-focused cytoplasmic calcium oscillates in response to tip growth (Monshausen et al. 2008). The ER plays an important role in regulating cytoplasmic Ca2+ distribution (Hong et al. 1999). Thus, it is possible that the Ca2+ gradient at the tip of the rhd3 root hairs is perturbed. Considering the remarkable functional similarity between RHD3 and Atlastin-1 (Hu et al. 2009; Chen et al. 2011; Stefano et al. 2012), further research on RHD3 in root hairs will not only shed light on how RHD3 works inside root hairs, but will also elucidate how Atlastin-1 may act in axonal neuron growth.

Finally, although the morphology of the plant ER-Golgi interface is different from that in mammalian cells, it appears that the molecular mechanisms governing the organization of the ER and Golgi are relatively conserved (Bassham et al. 2008). However, the mode of action of many trafficking relative genes or proteins in plant cells is different. It is thus pivotal to understand how these genes act in the context of the structure, dynamics and function of the ER-Golgi interface in plant cells.

(Co-Editor: Jianping Hu)

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References

This work was supported by a discovery grant from the National Science and Engineering Research Council of Canada to H.Z., and a grant from the Central Public-interest Scientific Institution Basal Research Fund (RIF2012–02) to J.C.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Tubules vs. Sheets: Roles of Reticulons and DP1/Yop1
  5. Making an Interconnected Network: The Role of Atlastin-1/RHD3
  6. Moving the ER in Plant Cells: The Roles of the Actomyosin System
  7. Association of the ER with Other Organelles
  8. Interplay Between the ER and Golgi in Plant Cells
  9. Future Directions
  10. Acknowledgements
  11. References