SPECIAL ISSUE PAPER
The plant ER–Golgi interface: a highly structured and
dynamic membrane complex
Patrick Moreau1,*, Federica Brandizzi2, Sally Hanton2, Laurent Chatre1,2, Su Melser1, Chris Hawes3and
Be ´atrice Satiat-Jeunemaitre4
1Laboratoire de Biogene `se membranaire, UMR 5200 CNRS-Universite ´ de Bordeaux II, case 92,
146 rue Le ´o-Saignat, F-33076 Bordeaux-Cedex, France
2Department of Biology, 112 Science Place, University of Saskatchewan, Saskatoon, SK, Canada S7N 5E2
3Research School of Biological and Molecular Sciences, Oxford Brookes University, Oxford OX3 0BP, UK
4Laboratoire de Dynamique de la Compartimentation Cellulaire, Institut des Sciences du Ve ´ge ´tal,
CNRS UPR 2355, Avenue de la Terrasse, F-91400 Gif sur Yvette-Cedex, France
Received 20 January 2006; Accepted 26 July 2006
As compared with other eukaryotic cells, plants have
developed an endoplasmic reticulum (ER)–Golgi in-
terface with very specific structural characteristics. ER
to Golgi and Golgi to ER transport appear not to be
dependent on the cytoskeleton, and ER export sites
have been found closely associated with Golgi bodies
to constitute entire mobile units. However, the molec-
ular machinery involved in membrane trafficking
seems to be relatively conserved among eukaryotes.
Therefore, a challenge for plant scientists is to de-
termine how these molecular machineries work in a
different structural and dynamic organization. This
review will focus on some aspects of membrane
dynamics that involve coat proteins, SNAREs (soluble
N-ethylmaleimide-sensitive factor attachment receptor
proteins), lipids, and lipid-interacting proteins.
Key words: Coat proteins, endoplasmic reticulum (ER), Golgi
apparatus (GA), lipids, membrane biogenesis, membrane
Eukaryotic cells have developed a complex organization
of biological membranes defining their intracellular
compartments. In plant cells, membrane biology has to
be adapted to specific requirements. For instance, it has
to organize the photosynthetic pathways in chloroplasts,
the formation of specific organelles for protein, glycan, or
lipid storage, permit rapid changes in vacuolar content and
membrane composition in response to osmotic or ionic
stresses whilst maintaining the correct turgor pressure, and
even organize de novo compartmentation during symbiotic
processes. Plant cell endomembranes also participate in
the organization of the transport and delivery of specific
secretory molecules (transport and delivery of cell wall
components, polarized distribution of specific plasma
membrane transporters, etc). As in any eukaryotic cells,
the plant cell secretory pathway is made up of a group
of discrete membrane-bound organelles [i.e. endoplasmic
reticulum (ER), Golgi apparatus (GA), endosomes/pre-
vacuoles, and vacuoles], working along a secretory gradient
to organize the processing, sorting, and delivery of cargo
molecules to their final destination (Neumann et al., 2003;
Hawes and Satiat-Jeunemaitre, 2005; Boutte ´ et al., 2006).
The functional and structural identity of each membrane-
bound compartment is related to the architecture/composi-
tion of its constitutive membranes (Moreau et al., 1998).
However, early views of membrane-bound compartments as
fixed structures are now being replaced by the concept that
they are highly dynamic (Brandizzi et al., 2004). It is now
* To whom correspondence should be addressed. E-mail: email@example.com
Abbreviations: ARF, ADP-ribosylation factor; CFP, cyan fluorescent protein; ER, endoplasmic reticulum; ERES, endoplasmic reticulum export site; GA, Golgi
apparatus; GAP, GTPase-activating protein; GEF, guanine-nucleotide exchange factor; GFP, green fluorescent protein; OSBP, oxysterol-binding protein;
PA, phosphatidic acid; PLA, phospholipase A; PLD, phospholipase D; SNARE, soluble N-ethylmaleimide-sensitive factor attachment receptor protein; TMD,
trans-membrane domain; YFP, yellow fluorescent protein.
ª The Author . Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: firstname.lastname@example.org
Journal of Experimental Botany, Vol. 58, No. 1, pp. 49–64, 2007
Intracellular Compartmentation: Biogenesis and Function Special Issue
doi:10.1093/jxb/erl135 Advance Access publication 21 September, 2006
by guest on November 6, 2015
clear that membrane composition and membrane-bound
structures evolved to fit cellular requirements, and that
membrane synthesis (and the turnover of membrane com-
ponents) is a highly regulated process. It is well known that
there can be a continuous exchange of membrane between
the compartments of the secretory pathway, alongside some
recycling processes to maintain compartment identity. The
complexity of the mechanisms involved in such membrane
dynamics and processing is increased further by the recent
findings that chloroplast proteins may also transit through
the GA, thus suggesting a possible membrane continuity
between what was once considered to be two entirely
independent organelles (Villarejo et al., 2005).
To understand the specificities of plant membrane
biology along the secretory pathway, several questions
must be addressed. (i) How is membrane identity acquired
for each compartment? (ii) How is membrane composition
maintained despite the vectorial membrane flow from ER to
plasma membrane? (iii) How is the membrane composition
adapted for differential developmental and environmental
programmes? (iv) What are the molecular machineries
regulating the passage of membrane from one compartment
to the other along the secretory pathway?
Here, an overview of the recent data gained from studies
on the ER–GA complex is provided, illustrating the
complexity of the processes involved in the exchange
between two distinct membrane interfaces. Moreover, since
components of the molecular machinery regulating these
processes appear to be conserved between eukaryotes,
unravelling the exact roles of these molecular structures
in the plant ER–GA interface, which appears to be
structurally different from that of animal and yeast cells,
is a tremendous challenge (Hawes and Satiat-Jeunemaitre,
2005). The requirement for specific sets of proteins and
lipids that ensure efficient transit of cargo from the ER to
the Golgi whilst maintaining the identities of the two
organelles will be discussed. In this context, the focus is on
the role of coat proteins, specific SNAREs (soluble N-ethyl-
maleimide sensitive factor attachment receptor proteins)
involved in membrane fusion, and the machinery that
maintains the lipid identity of the two compartments.
The plant ER–GA interface: a dynamic channel
From ER to GA: All proteins, membrane-bound or soluble,
thatare pre-destinedforexocytosis orstorage in intracellular
compartments of the endomembrane system, have to enter
the secretory pathway co-translationally via the Sec61 chan-
nel in the ER (Nicchita, 2002). Some may then undergo a
first set of glycosylation events, and the quality of their
processing will be checked by ER molecular chaperones
(Trombetta and Parodi, 2003). Most will then have to reach
the GA compartments, and transit through the Golgi stack
before reaching their final destination (Munro, 2005).
The ER and GA are very distinct organelles in plant
cells, having their own morphological, physiological, and
biochemical features (Vitale and Denecke, 1999; Hawes
and Satiat-Jeunemaitre, 2005). The ER is generally orga-
nized into a tubular reticulate network with occasional
cisternae, although the precise 3D organization may appear
to vary with the visualization techniques used and the
cell types observed (Satiat-Jeunemaitre et al., 1999). The
GA is usually made up of hundreds of discrete micron
size stacks of membrane-bound cisternae, usually termed
Golgi bodies, Golgi stacks, or dictyosomes. Direct con-
nections between cisternae are often observed (Hawes and
Satiat-Jeunemaitre, 2005; Ke ´pe `s et al., 2005). Moreover,
membrane-like continuity between ER and GA has been
observed using electron microscopy (see references in
Ke ´pe `s et al., 2005), and the ER has to participate in the
biogenesis of the GA (membrane synthesis, processing, and
sorting of Golgi-resident proteins). Conversely, the biology
of the GA will condition the structure of the ER as shown
by some drug-induced effects of GA disruption on ER
structure and function (Satiat-Jeunemaitre et al., 1996).
This tight functional and structural interaction between ER
and GA facilitates a mandatory passage between the two
compartments for secreted and membrane-bound proteins.
The next question to address is the mode of transfer of
cargo molecules between the two compartments.
Crossing the ER–GA interface: different transportation
systems?: Biochemical studies suggest that several distinct
mechanisms may effect the ER–GA passage of cargo
molecules (Moreau et al., 1998; see also additional
references in Me ´rigout et al., 2002), but their mechano-
structural support is still a question for further study.
Two main models, not mutually exclusive, have been
proposed to explain the passage of ER-processed molecules
to the GA. As electron microscopy studies have often
indicated membrane connections between the two compart-
ments in plant cells, it was first proposed that material
exchange could take place by tubular connections, either
transient or permanent. A major stream would be the
unspecific transport of cargo molecules (membrane-bound
or soluble), in so-called bulk flow (Denecke et al., 1990).
Alternatively, movement from one compartment to the
other could be selective and carried out by specific
structures such as transport vesicles (Contreras et al.,
2004b; see also above).
Arguments in favour of or against each of the proposed
models are outlined in many reviews, with comments on
themethodologicalapproaches, the biological material,and,
in some cases, the specific school of thought favoured in
this controversial debate (Neumann et al., 2003; Hanton
et al., 2005; Hawes, 2005; Ke ´pe `s et al., 2005). Therefore, it
will not be discussed further here. The recent development
of bioimaging techniques mainly using tobacco leaf
epidermal cells has contributed much to this debate, as
they demonstrate beautifully the close relationship between
the ER and GA (Boevink et al., 1998; Brandizzi et al.,
50 Moreau et al.
by guest on November 6, 2015
2002b, Saint-Jore et al., 2002; Chatre et al., 2005). These
bioimaging approaches on leaf epidermal cells or BY-2
cells have also revealed the amazing dynamics of the ER–
GA complex (Boevink et al., 1998; Nebenfu ¨hr et al., 1999).
Different types of movements of the ER–GA complex have
been observed: (i) rapid movement of protein on the ER
surface (Runions et al., 2006); (ii) organelle movements as
both ER and Golgi stacks can move with cytoplasmic
streams; (iii) more organized movements, as Golgi stacks
move over ER/actin tracks (Boevink et al., 1998); and (iv)
intraorganelle movement as ER contents can also move
within the lumen of the network (Hawes et al., 2001). Are
such movements necessary for the operation of ER to Golgi
transport? It has clearly been shown by photobleaching
experiments that Golgi movement is not necessary for the
passage of cargo molecules from the ER to the GA
(Brandizzi et al., 1992c), although transport can take place
towards moving Golgi (daSilva et al., 2004).
Asecond clear messagefromthese live-cell imaging data
is that the ER–GA interface can be crossed in both
directions, and these transport processes are independent
of a functional cytoskeleton (Saint-Jore et al., 2002).
Therefore, the regulation of ER–GA exchange is a subtle
balance between anterograde and retrograde transport, and
the regulation of such bidirectional movement of proteins
and lipids is mediated by distinct molecular machineries.
Quest for the molecular mechanisms associated with the
ER–GA interface: The ER–GA interface in plant cells
differs from the one described in animal cells. In the latter
case, an intermediate compartment is a cargo carrier
between ER and GA, and the role of microtubules in ER
to GA transport is well established (Murshid and Presley,
2004). Therefore, the molecular machineries associated
with the plant ER–GA interface must be plant specific at
least at the level of their fine tuning.
Whatever the nature of the transport intermediate
(tubular or vesicular), several types of structural and
regulatory molecules would most probably be required to
ensure efficient crossing of the ER–GA interface. (i)
Components that will facilitate the deformation of ER
membrane into tubules or vesicles (incidentally, one may
note that budding profiles on ER membranes are scarcely
observed in higher plant cells; meanwhile tubular exten-
sions are commonly seen). (ii) A machinery that would
facilitate the recruitment of specific cargo molecules either
membrane-bound or soluble, unless bulk flow is the only
mechanism associated with cargo exit in plant cells. (iii)
Components that will mediate the docking and fusion of
ER-derived membranes to Golgi membranes.
Different families of molecules can be involved in such
actions, and here the focus is on three important groups: the
coat proteins COPI and COPII with their associated
GTPases (ARF1 and Sar1), which may be involved in
specific recruitment processes, the SNARE proteins, which
may have a role in mediating specific membrane fusion
events, and finally the protein–lipid interactions, where
proteins may induce lipid remodelling and membrane
deformation (phospholipases, acyltransferases). Upstream
molecules such as the Rab GTPases (Batoko et al., 2000),
involved in regulating membrane fusion events, and
putative membrane-tethering factors (Latijnhouwers et al.,
2005) will not be discussed here.
Specific molecular machineries recruited at
the ER and Golgi membranes
The COPI and COPII coats
It is generally assumed that in yeasts and mammals,
proteins are exported from the ER in transport intermedi-
ates, which then fuse with the GA and release their contents
into this organelle. A common hypothesis proposes that the
formation of these carriers is mediated by a protein coat
made upof several proteins(Sar1, Sec23/Sec24,andSec13/
Sec31), termed COPII (Barlowe et al., 1994). Although
direct evidence for the existence of COPII transport
intermediates in plants has not yet been presented, the
identification of plant homologues of COPII coat proteins
implies that this pathway is present (d’Enfert et al., 1992;
Bar-Peled and Raikhel, 1997; Takeuchi et al., 1998;
Movafeghi et al., 1999; Belles-Boix et al., 2000). Further-
more, COPII components have been shown to influence
protein transport between ER and Golgi (Takeuchi et al.,
1998, 2000; Andreeva et al., 2000; Phillipson et al., 2001;
daSilva et al., 2004; Yang et al., 2005). It is also generally
assumed that the anterograde route is paralleled by
a retrograde pathway that transports selected cargo and
membrane from the Golgi back toward the ER, and
mediated by the COPI protein coat. In contrast to COPII
carriers, plant COPI vesicles have been visualized in situ
(Pimpl et al., 2000), although their function in retrograde
transport remains to be proven and even their exact role in
mammalian cells has yet to be elucidated (for a review see
Duden et al., 2005). Disruption of one route affects the
other, meaning that interruption of COPI-mediated trans-
port in the retrograde direction can prevent anterograde
transport mediated by COPII (Lee et al., 2002; Ritzenthaler
et al., 2002; Takeuchi et al., 2002; Pimpl et al., 2003;
Stefano et al., 2006). Whether this occurs in a direct or an
indirect manner has yet to be established (Stefano et al.,
In both directions of transport, coat assembly is initiated
by the activation of a small GTPase, achieved by the
interaction of the GTPase with a guanine-nucleotide
exchange factor (GEF), and its subsequent recruitment to
the membrane from which the vesicle eventually buds. In
non-plant systems, it has been shown that COPII carriers
are formed in response to the activation of Sar1, followed
by the recruitment of the larger coat subunits Sec23/Sec24
The plant ER–Golgi interface51
by guest on November 6, 2015
and Sec13/Sec31, which form the structural components of
the coat (Barlowe et al., 1994). The ER domains where
Sar1 (and other COPII proteins) are recruited define ER
export sites (ERES), with the Sar1 exchange factor, Sec 12,
being distributed over the surface of the ER (daSilva et al.,
2004). ARF1 plays a similar role in COPI vesicle
formation, recruiting the coatomer complex, made up of
seven different subunits within the cytosol and targeting the
resulting complex to the cis-Golgi membrane (Waters
et al., 1991; Palmer et al., 1993). The coat may have
several roles in membrane exchange: impose a curvature,
and/or specifically select some cargo molecules. The coat
has to be removed to allow fusion (and possibly transport)
of the carrier to its final destination. It was thought that GTP
hydrolysis was necessary for dissociation of the coat from
the vesicle, thus exposing the membrane in order for fusion
with the target membrane to occur. However, a recent study
in yeast has shown that GTP hydrolysis is required in order
for COPII vesicle fission from the ER membrane to occur
(Bielli et al., 2005). This hydrolysis is instigated by the
interaction of a GTPase-activating protein (GAP) with the
GTPase. In the case of COPII, the GAP function can be
performed by Sec23, a structural part of the coat itself
(Yoshihisa et al., 1993). However, in the case of COPI,
a Golgi-localized protein known as GAP1 catalyses GTP
hydrolysis (Randazzo, 1997). The GAP activity of GAP1
appears to be partially dependent on the presence of
coatomer (Szafer et al., 2000, 2001), which would reduce
the possibility for unproductive cycling of ARF1 between
the GTP- and GDP-bound states.
Interestingly, ARF1 may play several distinct roles
within the plant cell. It has also been shown to play a role
in a vacuolar sorting pathway in plants (Pimpl et al., 2003).
Fluorescent protein fusions of Arabidopsis thaliana ARF1
have been localized to compartments of unknown identity
derived from the GA (Stefano et al., 2006) and on putative
endocytic structures (Xu and Scheres, 2005). An ARF-GEF
that localizes to the endosomes has also been identified
(Geldner et al., 2003), indicating that different GEFs may
mediate alternative functions for GTPases within the cell. It
is not clear whether the function of the GEF is only to
recruit the GTPase to the membrane, or whether different
GEFs induce subtly different conformations of the GTPase,
resulting in modified functionality. No data have yet been
presented to suggest similar multiple functions for Sar1, but
this possibility cannot be ruled out.
Export mechanisms—diffusion (non-selective flow) or
Whatever the nature of the transport intermediate in plant
cells, the mechanism by which proteins are recruited into
this structure has long been a subject of intense debate and
may vary depending on the nature of the proteins in-
vestigated. It has been shown for instance that soluble
proteins can exit the ER via a bulk flow mechanism
(Denecke et al., 1990; Phillipson et al., 2001), although
the existence of receptors to concentrate and increase the
rate of export of certain soluble cargo molecules cannot be
ruled out. Saturation of specific transport steps along the
secretory pathway has also been observed, where high
levels of expression of a secretory cargo molecule can
reduce the efficiency by which it is transported (Phillipson
et al., 2001). Further investigations are required to discern
at which stage in the secretory pathway such saturation
occurs, as it may indicate the existence of a previously
unidentified cargo receptor. Additional evidence for the
bulk flow mechanism of ER export of soluble proteins is
provided by the existence of a retrieval mechanism for
rescuing soluble ER-resident proteins from the GA. A
tetrapeptide H/KDEL motif is found at the extreme C-
terminus of many soluble ER-residents protein (Denecke
et al., 1992) and is thought to interact with a receptor
molecule (most probably a yeast ERD2p homologue) in the
cis-Golgi, as described in other systems (Lewis et al., 1990;
Semenza et al., 1990). This interaction apparently triggers
the formation of COPI vesicles (Lewis and Pelham, 1992;
Aoe et al., 1997), leading to the transport of the cargo
molecule back to the ER.
In a similar way, some ER-resident transmembrane-
spanning proteins travel to the GA and are later retrieved
back to the ER. In these cases, a di-lysine motif is thought to
interact with the COPI coat at the cis-Golgi (Contreras et al.,
2004a). This retrieval system initially indicated that proteins
with transmembrane domains travel through the secretory
pathway by means of diffusion. Support for this model was
presented by Brandizzi et al. (2002a), who showed that the
length of the transmembrane domain plays a role in the final
destination of type I membrane-spanning proteins within the
secretory pathway. Short transmembrane domains restrict
such proteins to the ER, whereas longer ones permit export
todistallocations.Thisindicated thata diffusionmechanism
might exist, whereby a protein would travel through the
secretory pathway until it reached a membrane of sufficient
thickness to mask its hydrophobic transmembrane domain.
However, two recent publications have shown that certain
amino acid motifs in the cytosolic domains of transmem-
brane proteins can also influence ER export in vivo (Hanton
et al., 2005; Yuasa et al., 2005). Contreras et al. (2004b)
have shown that a di-hydrophobic motif interacts with
COPII coat subunits in vitro, although the relevance of such
an interaction has yet to be established in vivo. It has been
reported that a di-basic motif is involved in the ER–Golgi
transport of a prolyl hydroxylase–green fluorescent protein
(GFP) fusion(Yuasaetal.,2005).A rolefordi-acidic motifs
was addressed by Hanton et al. (2005) for different types of
Golgi-localized proteins. The authors demonstrated that not
only did the di-acidic motifs of cargoes influence their rate
of transport from ER to Golgi in plant cells, but also that
they were dominant over transmembrane domain length in
defining ER export. These findings indicate that although
52 Moreau et al.
by guest on November 6, 2015
diffusion is one way by which proteins can exit the ER,
other mechanisms also exist to ensure efficient export.
Cytosolic signals may also contribute to allow fast tracking
of certain proteins out of the ER, although it is not yet clear
how these signals facilitate export of proteins. Studies in
non-plant systems have suggested that COPII coat compo-
nents interact with certain motifs and induce the formation
of transport carriers. These findings are supported by the
data presented by Contreras et al. (2004b) regarding di-
hydrophobic signals, although further investigations are
required to elucidate the role of other signals in ER export.
If cytosolic export motifs induce formation of COPII
carriers, it seems likely that other transmembrane proteins
might enter these carriers by means of diffusion and exit the
ER by this process.
The SNARE machinery
General characteristics of SNAREs
SNAREs in eukaryotic cells favour the fusion of two
apposed lipid bilayers and, through their specificities,
contribute to the targeting of membrane components to
maintain membrane identity and functions (Hong, 2005).
Compared with Homo sapiens (35 SNAREs), Drosophila
melanogaster (20 SNAREs), and Saccharomyces cerevi-
siae (21 SNAREs), the Arabidopsis genome contains
a greater number of SNAREs, since at least 54 genes
have been identified (Sanderfoot et al., 2000; Pratelli et al.,
2004; Uemura et al., 2004). This large number of SNARE-
related proteins in plants may reflect a plant-specific
diversity in their cellular functions (for instance cell growth
and development, autophagy, gravitropism, stress re-
sponses, and resistance to pathogens) that can be related
or not to their fusogenic properties (Pratelli et al., 2004;
Surpin and Raikhel, 2004).
SNAREs are mostly anchored to the cytosolic side of the
membrane by a C-terminal trans-membrane domain (TMD)
or a prenyl group, and a few, such as SNAP25, are bi-
palmitoylated (Hong, 2005). SNAREs are characterized by
one or rarely two SNARE motifs (coiled-coil domains)
helical bundle (Hong, 2005), and such domains are
sufficient for triggering membrane fusion (Hu et al.,
2005a). SNAREs were first classified into v-SNAREs and
Depending on the presence of an arginine or a glutamine in
a central position of the helical bundle (the ‘zero layer’) of
the SNARE motifs, SNAREs have since been classified
Five subfamilies have since emerged: Qa-SNAREs (syn-
taxin, t-SNAREs with Q in the zero layer and a single
layer and a single SNARE motif similar to the N-terminal
SNARE motif of SNAP25), Qc-SNAREs (v-SNAREs or
syntaxin-like with Q in the zero layer and a single SNARE
motif similar to the C-terminal SNARE motif of SNAP25),
R-SNAREs (VAMPs, v-SNAREs with R in the zero layer
and a single SNARE motif), and finally the SNAP25 family
with two SNARE motifs (Hong, 2005). Rossi et al. (2004a)
have proposed dividing VAMPs into two additional sub-
families, RD-SNAREs and RG-SNAREs, according to the
conserved flanking amino acid (D or G) in the C-terminus
of their zero layer.
Finally, SNAREs can also be classified according to their
N-terminal domains. The most complex organization is
found in the syntaxins which have three helical structures
(Ha, Hb, and Hc) preceded by an N-terminal low complex-
ity domain (Hong, 2005). R-SNAREs/VAMPs are also
subdivided into short VAMPs with a very short N-terminus
(brevins) or long VAMPs with a longer N-terminus
(longins). The longin domains are structurally related to
profilin (Rossi et al., 2005b).
Plant ER–Golgi SNAREs
The subcellular localizations of putative or established
SNAREs of the ER–Golgi interface, based on protein
expression, effects of brefeldin A treatments, and in vivo
light microscopy analyses of fluorescent protein fusions
(Uemura et al., 2004; Chatre et al., 2005), are given in
Table 1. The fusogenic potential of plant SNAREs has
rarely been studied in vitro, and the published studies are
restricted to SNAREs from the trans-Golgi and the Golgi/
prevacuolar compartments (Sanderfoot et al., 2001; Chen
et al., 2005). In this review, the focus is on SNAREs
predicted to function at the ER–Golgi interface and within
the Golgi cisternae.
Some of these SNAREs are remarkable in the compo-
sition of their zero layer. Indeed AtBet11 (BS14a, Qc-
SNARE) contains a histidine in the zero layer instead of
a glutamine (Tai and Banfield, 2001) and AtSec22 (R-
SNARE) contains a valine instead of an arginine. The same
observation can be made for the yeast SNARE Bet1 which
harbours a serine at the zero layer (Tai and Banfield, 2001),
and for the mammalian Vti1a (Qb-SNARE) and Slt1 (Qc-
SNARE) which have an aspartate (Hong, 2005). Therefore,
SNARE interactions are either not affected by such changes
in the nature of this residue or these ER–GA SNAREs
perform their function by interacting differently.
Compared with mammals where SNARE expression can
be highly tissue specific (Rossi et al., 2004b), it appears that
SNAREs are widely expressed in plant tissues, and this is
particularly true for the SNAREs of the ER–Golgi interface
(Uemura et al., 2004). However, since the number of
SNAREs in plants is higher and considering that these
proteins may have very different functions in various tissues
(Pratelli et al., 2004; Surpin and Raikhel, 2004), it may be
expected that post-transcriptional or post-translational reg-
ulations confer tissue-specific activities to these proteins.
The plant ER–Golgi interface53
by guest on November 6, 2015
In plants, the role of the SNAREs has only recently been
determined in vacuolar transport, cell surface assembly, and
cell-plate formation duringcytokinesis (Pratelli et al.,2004;
Surpin and Raikhel, 2004; Ju ¨rgens, 2005). However, the
role of the different SNAREs in the early secretory pathway
of plants has yet to be established. Due to their homology
with mammalian and yeast proteins, and their location in A.
thaliana suspension-cultured cells (Uemura et al., 2004),
the SNAREs AtSec22, AtMemb11, and AtSYP31/AtSed5
(related to the yeast syntaxin Sed5) were expected to act at
the early ER–Golgi step. Figure 1 shows the intracellular
location of AtSec22–YFP and AtMemb11–YFP in tobacco
leaf epidermal cells and the effects of brefeldin A on their
distribution, suggesting that these SNAREs are at the ER–
Golgi interface (Chatre et al., 2005). Overexpression of
AtSec22-C–YFP or AtMemb11-C–YFP [SNAREs transla-
tionally fused to either the YFP or CFP] in this system
affected the dynamics of AtERD2-Y–CFP (the putative
Arabidopsis H/KDEL receptor, translationally fused to
either the YFP or CFP) and another Golgi reporter fusion
protein (ST–YFP, partial rat sialyltransferase fused to YFP)
with subsequent redistribution of these markers into the ER
(Chatre et al., 2005). In addition, the trafficking of a
secreted marker (secYFP) was mainly blocked at the level
of the ER by co-expression of the two v-SNAREs (Chatre
et al., 2005). Thus, AtSec22 and AtMemb11 appear to be
critical v-SNAREs of the ER–Golgi interface in tobacco
leaf epidermal cells, and are at least involved in the
anterograde pathway. Whether these SNAREs are also
involved in the retrograde pathway as reported for Sec22p
in yeast (Burri et al., 2003) has to be determined.
AtBet11 (translationally fused to CFP), another v-
SNARE closely related to animal Bet1 (regulating ER–
Golgi transport in other eukaryotic cells; Hong, 2005), did
not affect the distribution of either AtERD2–YFP or ST–
YFP, an observation which is in agreement with its
estimated trans-Golgi location (Uemura et al., 2004; Chatre
et al., 2005). On the contrary, the trafficking of secYFP was
partly blocked at the level of the ER by the expression of
AtBet11–CFP (Chatre et al., 2005). As a consequence, it is
suggested that AtBet11 is also involved in the anterograde
pathway but probably at a later stage in the Golgi than
AtSec22 and AtMemb11. Due to a cis-Golgi distribution,
the AtBet11 isoform AtBet12 could be required in earlier
steps in the Golgi (Uemura et al., 2004).
The situation is perhaps more complex for AtSYP31
(named AtSed5 in Chatre et al., 2005). Brefeldin A
treatment was found to change the labelling of AtSYP31
to a more typical ER pattern in A. thaliana (Uemura et al.,
2004) than in tobacco leaf epidermal cells where aggre-
gated membranes (cAggr. in Table 1) were also observed.
Table 1. Plant ER/Golgi SNAREs: subcellular location, BFA effect and putative functions
The data are compiled from Uemura et al. (2004) and Chatre et al. (2005). The subcellular locations are deduced from the observations by confocal
microscopy of chimeric SNARE–XFP constructs expressed in transformed plants or protoplasts. The cis/trans localization was evaluated from the
assumption that brefeldin A selectively redistributes the cis- or far trans-Golgi-localized proteins (respectively cis redistributed in ER, and trans in
cytosolic aggregates of membranes). The role of a few SNAREs has been investigated between the ER and the Golgi (AtSYP 31, AtMemb11, and
AtSec22). AtBet11 is probably more involved later in the secretory pathway according to its subcellular location. For the other SNAREs, their role is
either estimated by analogy to their animal and yeast counterparts (AtSYP81), or still unknown. cAggr., membrane aggregations in the cytosol induced
by brefeldin A.
AtSYP31, At5g05760; AtSYP32, At3g24350; AtSYP81, At1g47920; AtMemb11, At2g36900; AtMemb12, At5g50440; AtGos11, At1g15590;
AtGos12, At2g45200; AtSYP71, At3g09740; AtSYP72, At3g45280; AtSYP73, At3g61450; AtBet11, At3g58670; AtBET12, At4g14450; AtSEC22,
At1g11890; AtVAMP714, At5g22360; AtVAMP723, At2g33110.
Plant ER/Golgi SNAREs
ER + cAggr.
ER + cAggr.
Golgi–ER or ER fission?
Golgi–ER or ER fission?
Golgi–ER or ER fission?
54 Moreau et al.
by guest on November 6, 2015
cells did not affect the dynamics of AtERD2–YFP and
ST–YFP to the same extent as did overexpression of
AtSec22 and AtMemb11, but the trafficking of secYFP
was predominantly blocked at the level of the ER (Chatre
et al., 2005). Since AtSYP31 is believed to be the t-SNARE
syntaxin in the cis-Golgi, it could be predicted that when
AtSYP31 is overexpressed, Golgi reporter proteins are less
retained into the ER but more at the ERES, which cannot be
distinguished from the Golgi bodies (daSilva et al., 2004).
It has recently been shown in yeast that AtSYP31 may,
under certain conditions, be by-passed in the activation of
SNARE pairing for the formation of fusogenic SNARE
complexes (Peng and Gallwitz, 2004). Such discrepancies
could be due to different levels or distributions of the
expressed proteins. A 3–5 times higher level of expressed
tagged SNAREs compared with the endogenous proteins
can lead to a substantial shift in the distribution of the
expressed proteins (Cosson et al., 2005), and the conse-
quence of such a partial shift on the functionality of the
tagged protein may vary according to the fusion protein
considered. In addition, these authors have shown that the
location of SNAREs is not due to retention mechanisms but
proceeds from dynamic transport equilibrium. The distur-
bance of such an equilibrium and overconcentration at
specific sites may perturb membrane dynamics and protein
function. Therefore, further investigations by electron
microscopic immunocytochemistry need to be conducted
between ER, ERES, and the different Golgi cisternae.
This discussion may question the validity of localization/
functional analyses using such overexpressed tagged pro-
teins. The use of endogenous promoters may solve this
problem, but insofar as the level of expression is sufficient
to visualize the fusion proteins. It has also to be considered
that, besides approaches using loss of function, approaches
using gain of function such as overexpression can report
No studies are yet available on the other putative ER–
Golgi SNAREs presented in Table 1. We can only speculate
that the ER-localized AtSYP81, which is a homologue of
the animal Syn18 and the yeast Ufe1 known to be active in
Golgi–ER retrograde transport, could be involved in any
retrograde pathway ending in the ER. Finally, the existence
of several syntaxin-like SNAREs such as AtSYP71, 72,
and 73 in the ER could suggest their requirement in multiple
retrograde pathways and/or fission events of the ER in
other membrane biogenesis processes. All the SNAREs so
far reported at the ER and Golgi level seem to be expressed
ubiquitously (Uemura et al., 2004), but their abundance
and tissue specificity will have to be determined through
quantitative polymerase chain reaction (QPCR).
Although the central role of SNAREs in membrane fusion
has been elucidated, only a few studies have been devoted
to investigating how theses proteins are targeted to their
It has been shown that the targeting of the mammalian
YKT6, a C-terminal isoprenylated SNARE, is dependent
on the N-terminus profiling-like longin domain and not
driven by the isoprenylation per se (Hasegawa et al., 2003).
In fact, it has been proposed that the longin domain
interacts with membrane lipids and the SNARE motif in
the cytosol so that only the longin domain is available for
interaction with the target membrane (Hasegawa et al.,
2004). It has also been reported that targeting of mamma-
lian SNAP-25, a plasma membrane palmitoylated SNARE,
is independent of both palmitoylation and its interaction
with the syntaxin 1A, and that an interaction with another
membrane factor must be responsible for its targeting
(Loranger and Linder, 2002).
It is known that type I membrane proteins can be
anchored in specific membranes according to the length
of the TMD (Brandizzi et al., 2002a), but the situation for
C-terminal anchored proteins such as SNAREs is unknown.
In yeast, Bet1p, Gos1p, Sft1p, and Sed5p have a 15 amino
Fig. 1. Effect of brefeldin A (BFA) on the subcellular location of
AtSec22 and AtMemb11. (A) Localization of AtSec22–YFP in a tobacco
leaf epidermal cell. Golgi bodies and ER membranes are labelled by the
fluorescent protein construct. (B) Effect of BFA (50 lg ml?1for 30 min)
on AtSec22–YFP distribution. The protein construct is mostly redirected
to the ER membranes. (C) Localization of AtMemb11–YFP in a tobacco
leaf epidermal cell. Only the Golgi bodies are labelled by the fluorescent
protein construct. (D) Effect of BFA (50 lg ml?1for 30 min) on
AtMemb11–YFP distribution. The protein construct is entirely redis-
tibuted to the ER membranes.
The plant ER–Golgi interface55
by guest on November 6, 2015
acidTMDand are located in the Golgi, whereas Sec22p and
Slt1p, located in the ER, have a 19–20 amino acid TMD
(Burri and Lithgow, 2004). In plants, AtSec22 and
AtSYP31 both have a 17 amino acid TMD with different
subcellular localizations (Uemura et al., 2004; Chatre et al.,
2005). Therefore, it is unlikely that the length of the TMD
of the integral SNAREs explains their location. This may
be due to the different orientation of the proteins in the ER
membranes withrespect to type I proteins. Itmay have to be
asked instead whether the N-terminal cytosolic domain(s)
and the SNARE motifs of these proteins are required in
SNARE targeting. The SNARE motif of Bet1 has been
shown to contribute its subcellular targeting in NRK cells,
and this function is independent of the heteromeric SNARE
interactions of the protein (Joglekar et al., 2003). This
highlights a very important point: that the SNARE motif
has the ability to interact with different partners for different
tasks. In the case of the animal and yeast Sec22, it has been
shown that both the longin and the SNARE domains are
required for an efficient packaging of Sec22p into COPII
vesicles (Liu et al., 2004). In addition, the longin domain is
not a regulatory factor for SNARE complex assembly, and
SNARE pairing is not required for COPII-dependent
Sec22p export from the ER (Liu et al., 2004). It appears
that a sequence of 10 amino acids in the SNARE motif of
Sec22p is critical for recognition by COPII, and Liu et al.
(2004) have proposed that the longin and the SNARE
motifs interact with the Sec23/Sec24 complex at two
Although AtSec22 has a KXKXX sequence at the C-
terminus which could be an ER targeting motif, since
AtSec22 is a C-terminus tail-anchored protein, the C-
terminus is in the luminal side of the membrane and thus
unlikely to be an ER targeting determinant. Mutagenesis
approaches are required to unravel the motifs or domains
involved in its targeting. For example, it would be in-
teresting to investigate if its longin domain is critical since
the vacuolar targeting of VAMP7 SNAREs appears to be
dependent on their entire N-terminus (Uemura et al., 2005).
It is clear that SNARE–coat protein interactions can be
critical for SNARE targeting and dynamics, but also for
COP protein recruitment. This has recently been high-
lighted for the COPII and COPI machineries, and the ER–
Golgi SNAREs. Specific interactions were first observed
between Sed5p and the COPII component Sec24p in yeast
(Peng et al., 1999). More recently, several binding sites on
the Sec23/Sec24 complex and especially on Sec24p have
been identified which govern the selection of several yeast
SNAREs (Bet1, Sec22, and Sed5) and other cargo proteins
(Miller et al., 2003; Mossessova et al., 2003). In addition,
the selective Sec23/Sec24-dependent packaging of the
SNAREs was determined to be specific for the fusogenic
monomeric or complexed forms of the SNAREs, suggest-
ing a tight control of fusion starting at the budding level
(Mossessova et al., 2003). Finally, additional sites must
exist in the COPII proteins to explain the efficiency and the
selectivity of the ER–Golgi step for different cargo proteins
including the SNAREs (Miller et al., 2005). Interestingly,
it has also been suggested that v-SNAREs may function
as ARF receptors on membranes (Rein et al., 2002; Honda
et al., 2005). Therefore, different sets of regulators (GTP-
binding proteins such as ARF, Sar1, Rab, etc., SNAREs,
COPI and COPII coat proteins, and tethering factors) may
provide multiple specific interactions between several
partners, and drive the specificities of the targeting and
fusion events. In addition, it has recently been observed that
different SNAREs can have various affinities for specific
membrane domains (Salau ¨n et al., 2005a), and that the
degree of their association with these domains may regulate
the efficiency of exocytosis (Salau ¨n et al., 2005b). As
a consequence, domain-forming lipids and lipid-modifying
enzymes will also have to be considered in the driving and
regulation of these events.
SNAREs in fusion and beyond?
The physical action of SNAREs in membrane fusion has
been investigated in several animal and yeast models, and
all studies conclude that fusion can proceed through
a hemifusion intermediate (Xu et al., 2005; Reese et al.,
2005; Ungermann and Langosh, 2005). In addition, hemi-
fusion may not be just an intermediate step to complete
fusion but may also participate in transient fusion reactions
during reversiblekiss-and-run events(Giraudoet al., 2005).
In all fusion events, v-SNAREs may control the successive
steps involved: the N-terminus regulates vesicle priming,
the SNARE motif drives initiation of fusion, and the TMD
may control fusion pore formation (Borisovska et al., 2005;
Ungermann and Langosh, 2005). The SNARE complexes
comprise four helices with a Qa/Qb/Qc/R stoichiometry,
and replacing one Q by an R in the central zero layer can be
lethalorinduces a growthdefect in yeastthatcanberescued
by replacing the R of the R-SNARE with a Q (Graf et al.,
2005). However, all zero level positions do not seem to be
critical to the same extent as the fact that the functionality of
some of these proteins can be retained after residue
substitution. In addition, it has been observed that only
some SNARE combinations are functional. For example,
Sec22p can be replaced by YKT6p in yeast (Liu and
Barlowe, 2002) but the reverse is not true. The extent to
which some redundancy exists in the SNAREs of Arabi-
dopsis encoding 54 of these proteins is a relevant question
waiting to be addressed. As a support observation, Niihama
et al. (2005) have shown that a single mutation in the
SNARE VTI12 can rescue the zig-1 mutant of Arabidopsis
which lacks the SNARE VTI11, indicating some flexibility
in SNARE functions through gene duplication.
In addition, it has recently been suggested that SNAREs
themselves and SNARE-like proteins (amysin, tomosyn)
may compete to participate in non-functional SNARE
56 Moreau et al.
by guest on November 6, 2015
complexes that will inhibit and regulate fusion (Scales
et al., 2002; Varlamov et al., 2004; Constable et al., 2005).
The specificities of SNARE complexes in the different
membrane trafficking pathways are, to some extent, re-
capitulated in cell-free fusion assays with isolated
SNAREs. It has been suggested that non-functional com-
plexes may also have a physiological relevance, and the so-
called ‘i-SNAREs’ (inhibitory SNAREs) may contribute to
control thespecificity of membrane fusion (Varlamov et al.,
2004). For example, according to SNARE concentration
gradients in the Golgi, SNAREs may have a normal
SNARE function or an i-SNARE function in different
cisternae, and this would confer a sort of buffering effect on
the fusion capacity in the cisternae (Varlamov et al., 2004).
As proposed for Bet1 in animal cells (Varlamov et al.,
2004), Bet11 in the trans-Golgi of plant cells could also
have such an i-SNARE role. Finally, animal proteins
(Sec22a and Sec22c) are expressed without any genuine
SNARE motif and may participate in the regulation of
SNARE complex formation (Gonzales et al., 2001).
Lastly, another function which could be managed by
SNAREs, especially by syntaxins, is a more structural
action on membrane architecture. It has recently been found
that mutations affecting the phosphorylation of Sed5p
cannot only disturb ER–Golgi traffic in yeast but can also
affect the morphology of the Golgi (Weinberger et al.,
2005). AtSYP31 seems to have a wide distribution across
plant Golgi (Uemura et al., 2004; Chatre et al., 2005). Its
cis-Golgi location is compatible with a function as a syn-
taxin in ER–Golgi transport, but a distribution from the
cis- to the trans-cisternae could indicate that this syntaxin
is also participating in structural aspects of plant Golgi.
homologous to Sec22b
Lipids, lipid-modifying enzymes, and
As stated by Engelman (2005), ‘membranes are more
mosaic than fluid’. The random state of membrane organi-
zation of the Singer–Nicholson model is now replaced by
one reflecting a more patchy organization with different
domains (composition, thickness, turnover, and, therefore,
homeostasis). In such a view, membrane properties are
highly dependent on protein–lipid interactions where lipids
organize proteins and vice versa. As a consequence,
membrane structure and membrane trafficking are con-
trolled by close relationships between protein-based and
lipid-based machineries (De Matteis and Godi, 2004).
Beside the roles of coat proteins and SNAREs in membrane
organization and dynamics (structure, deformation, fusion),
lipids, lipid-modifying enzymes (phospholipases, acyltrans-
ferases), and lipid-modified proteins (acylated or isopreny-
lated GTP-binding proteins for example) also have key
roles to play in membrane dynamics, for instance through
the control of membrane curvature (McMahon and Gallop,
2005). In this last section, lipids are considered not as
membrane bricks but more as molecular architects (by
themselves or through protein modifications) of membrane
dynamics regulating trafficking.
Sterols, oxysterols, and related proteins
The structure of the sterol molecules can affect and regulate
the curvature of the membranes which will help to prepare
a membrane for budding or fusion (Bacia et al., 2005). In
animal cells, cholesterol has been shown to be required for
the formation of both constitutive and regulated post-Golgi
secretory vesicles (Wang et al., 2000),and in the biogenesis
of synaptic vesicles (Thiele et al., 2000). It has been shown
that cholesterol may facilitate vesicle fusion in exocytosis
by virtue of its intrinsic negative curvature, and contributes
to fast Ca2+-triggered membrane fusion (Churchward et al.,
2005). In yeast, ergosterol has also been found to be a key
regulator of endocytosis, and several mutants of this
pathway, the erg mutants, are affected in the ergosterol
metabolic pathway (Heese-Peck et al., 2002). On the other
hand, the cholesterol levels in mammalian Golgi mem-
branes must be tightly regulated because an excess of
cholesterol can lead to a vesiculation of the Golgi complex
itself, and this process is dynamin and phospholipase A2
(PLA2)-dependent (Grimmer et al., 2005).
Oxysterols are naturally occurring hydroxylated sterol
derivatives of interest in the secretory pathway of eukary-
otic cells. Oxysterol-binding proteins (OSBP) are effec-
tively lipid receptors involved in sterol homeostasis, being
able to regulate vesicular transport in both animal and yeast
cells (Xu et al., 2001; Li et al., 2002). In yeast, the related
OSBP (Kes1p) has been shown to be targeted to the Golgi
complex through binding to phosphoinositides formed by
a phosphatidylinositol kinase (Pik1p), and may regulate
Golgi secretory function through interactions with the
ARF(s) and Sec14p pathways (Li et al., 2002). In animal
cells, one OSBP and several OSBP-related proteins were
found to interact with a syntaxin-like VAMP-associated
protein-A, and have been shown to participate in the
organization of the COPII-dependent ER–Golgi pathway
(Wyles et al., 2002; Wyles and Ridgway, 2004).
In plant cells, the sterol biosynthetic pathways lead not
only to mature membrane sterols such as sitosterol and
stigmasterol, but also to cholesterol or the plant steroid
hormones (brassinosteroids). The study of dwarf mutants
disturbed in brassinosteroid synthesis has clearly demon-
strated the role of these steroid hormones in plant de-
velopment (Fujioka and Yokota, 2003; Schaller, 2003).
Interestingly, it has then been found that membrane sterols
are also critical for controlling cell growth, cell polarity,
and embryonic development (Schaeffer et al., 2001;
Schrick et al., 2002; Willemsen et al., 2003; Schaller,
The plant ER–Golgi interface57
by guest on November 6, 2015
2004). On the contrary, little is known about the role of
sterols in the plant secretory pathway and Golgi dynamics.
Preliminary approaches to answer these questions, based
upon pharmacological experiments, are yielding some
promising results. For instance, blocking cyclopropylsterol
maturation at the level of the cycloeucalenol–obtusifoliol
isomerase by fenpropimorph disturbs the secretory pathway
and induces a fenestration of the Golgi bodies (Hartmann
et al., 2002). Interestingly, a strong perturbation of the
morphology of the Golgi by brefeldin A leads to a reduced
synthesisof phytosterols (Me ´rigout et al., 2002). Therefore,
some relationships may exist between sterol metabolism
and Golgi morphology in plant cells. Recently, lipid rafts
have been isolated from the plasma membrane of plant
cells, and as expected, the phytosterols are enriched in these
domains (Mongrand et al., 2004; Borner et al., 2005). We
have recently found evidence of lipid rafts in the Golgi
membranes of plant cells, and observed that a perturbation
of sterol metabolism by fenpropimorph can block sterol
precursors and lipid rafts in the Golgi (Laloi et al.,
unpublished results). As a consequence, mature sterols
are also probably critical for membrane trafficking as in
Sphingolipid metabolism and organelle structure
Although the metabolism of cerebrosides and inositol-
sphingolipids (the two families of sphingolipids in plants)
have been extensively studied (Dunn et al., 2004; Sperling
et al., 2004), nothing is known about their roles in the
secretory pathway of plants. These lipids are also enriched
in plasma membrane lipid rafts (Mongrand et al., 2004;
Borner etal., 2005),buttheir actionin membrane traffic has
yet to be established in plants.
Several studies have, however, investigated such a role
in other eukaryotic cells. For example, natural long chain
ceramides, either incorporated or formed upon sphingo-
myelinase treatment in animal cells, enhance the disassem-
bly of the Golgi induced by brefeldin A, suggesting that the
levels of ceramides may be critical for Golgi stability
(Fukunaga et al., 2000). Pharmacological approaches using
inhibitors of glucosylceramide synthase have confirmed a
role for sphingolipids in the maintenance of Golgi architec-
ture, and in anterograde membrane trafficking (Nakamura
et al., 2001). It has been shown that short chain ceramides
can decrease the binding of ARF to Golgi membranes and
therefore affect the formation of COPI vesicles (Abousal-
ham et al., 2002). Finally, recent investigations in animal
cells have suggested that high concentrations of sphingo-
sine formed from the hydrolysis of ceramides can induce
Golgi fragmentation (Hu et al., 2005b). Overall, these
studies highlight a likely role for sphingolipids in Golgi
morphodynamics as found for cholesterol. However, the
situation is less clear in yeast where the various post-Golgi
pathways appear not to be systematically dependent on
sphingolipid biosynthesis (Lisman et al., 2004).
Phospholipases, acyltransferases, and lipid-binding
More and more studies reveal the critical impact of
phospholipid modifications (acylations or de-acylations)
on membrane dynamics and particularly in the case of the
Golgi membranes in animal cells. Evidence has been ob-
tained for the regulation of Golgi structure and membrane
trafficking by PLA2which cleaves the fatty acids of phos-
pholipids at the sn-2 position of the glycerol (Choukroun
et al., 2000), and the phospholipase activity induces mem-
brane tubulation (De Figueiredo et al., 1998). Such proper-
ties of PLA2to promote deformation of membranes have
been confirmed on giant liposomes where PLA2-induced
budding and fission events have been observed (Staneva
et al., 2004), and these events are caused by the formation
of positive membrane curvatures induced by the lysophos-
pholipids (Brown et al., 2003). Interestingly, the reverse
reaction of phospholipid synthesis by acyltransferases sta-
bilizes membrane bilayers, and inhibition of such enzymes
also leads to Golgi membrane tubulation (Drecktrah et al.,
2003). PLA1-related enzymes (which cleave the fatty acids
of phospholipids at the sn-1 position of the glycerol) have
been found to induce dispersion of the Golgi complex
and/or aggregation of the ER membranes (Nakajima et al.,
2002), and they can interact with proteins of the COPII
machinery to participate in the organization of the ER exit
sites (Shimoi et al., 2005). No evidence for the involvement
of such enzymes in Golgi dynamics and the early secretory
pathway of plant cells has yet been shown.
Another critical phospholipase in Golgi function is
phospholipase D (PLD) that cleaves the polar head to
produce phosphatidic acid (PA). Inhibition of its activity
(stimulated by ARF and/or Sar1p) decreases the level of PA
but also of phosphoinositides that are required for structural
integrity of the Golgi complex in animal cells (Siddhanta
et al., 2000). Interestingly, PA regulates the synthesis of
phosphoinositides but the latter can also regulate the PLD
activity. It has been found that PLD activation participates
in the formation of COPII complexes in ER export (Pathre
et al., 2003). An isoform of PLD has been localized to the
rims of the Golgi complex and may be involved in Golgi
morphodynamics playing a role in vesicular transport from
the Golgi (Freyberg et al., 2002). The critical role of PLD
has also been determined in yeast (Routt et al., 2005 and
references therein). Beside the role of PA in stimulating
phosphoinositide synthesis, a role for PA and its derivatives
lyso-PA and diacylglycerol in regulating membrane curva-
ture has also been proposed (Kooijman et al., 2003).
In plant cells, it is clear from the literature that PLD and
the formation of PA are required for normal plant growth
and polarized cell expansion (Gardiner et al., 2003; Potocky
et al., 2003). It has recently been shown that inhibition
of PLD in the pollen tube considerably reduces the number
of secretory vesicles, and that PA and phosphoinositides
58 Moreau et al.
by guest on November 6, 2015
regulate pollen tube growth (Monteiro et al., 2005). Despite
numerous studies on PLD and PA signalling in plant
physiology, no studies have yet addressed the putative role
of PLD in the early secretory pathway of plants.
Another family of proteins relevant to Golgi function are
the phospholipid-binding proteins. It has been shown that
Sec14p and Nir2p are both involved in the control of
diacylglycerol levels in Golgi membranes in yeast and
mammal cells, respectively (Kearns et al., 1997; Litvak
et al., 2005). Maintenance of diacylglycerol concentrations
in Golgi membranes is certainly related to the property of
diacylglycerol in membrane fission. In plant cells, a Sec14
candidate has been cloned from Arabidopsis which can
complement the yeast mutant (Jouannic et al., 1998), but no
role at the Golgi level has been established. Recently, a
Sec14p-nodulin domain PITP from Arabidopsis (a Sec14p-
like protein) has been demonstrated to be key regulator of
polarized membrane growth in root hairs (Vincent et al.,
Lipid domains and selective transport at the ER–Golgi
Phosphatidylserine accumulates in the plasma membrane of
leek cells and originates from both the ER after intracellular
trafficking and a local synthesis by the serine exchange
enzyme (Bessoule and Moreau, 2004). Both phosphatidyl-
serine synthase and the serine exchange enzyme can
synthesize phosphatidylserine in the ER (Bessoule and
Moreau, 2004). As a first example of phospholipid sorting
at the ER–Golgi interface, it has been determined that only
the serine exchange enzyme synthesizes phosphatidylserine
with very long chain fatty acids that are targeted to the sec-
retory pathway (Sturbois-Balcerzak et al., 1999; Vincent
et al., 2001). Phosphatidylserine was also found to be
targeted to ER-derived vesicles in rat liver membranes
(Moreau et al., 1992, 1993). This suggests the existence of
specific ER domains where such lipids are concentrated
before transport, and this poses the question as to whether
these domains correspond to the ER export sites revealed
by the COPII machinery (da Silva et al., 2004; Stefano
et al., 2006). Interestingly, C-tail-anchored proteins with
a moderate hydrophobic TMD can reside in the ER,
whereas addition of non-polar amino acids in the TMD
can translocate the protein to the plasma membrane. This
protein sorting toward the secretory pathway may be linked
to lipid-based sorting mechanisms, and specific interactions
with saturated fatty acid-bearing acidic phospholipids such
as phosphatidylserine may be relevant (Ceppi et al., 2005).
Another interesting finding was the requirement for phos-
phatidylserine in the formation of ER-derived COPII
vesicles in vitro (Matsuoka et al., 1998). Finally, the
well-established association of GTP-binding proteins
(such as Sar1, ARFs, and Rabs) with membranes, governed
by lipid modifications and lipid interactions, indicates that
lipids are crucial at several levels in membrane trafficking.
Challenges for the future
We can wonder whether Golgi bodies mature from the ER
export sites in a similar manner to that occurring in Pichia
pastoris, S. cerevisiae, or Drosophila oocytes (Bevis et al.,
2002; Glick, 2002; Herpers and Rabouille, 2004), and
whether the cis-Golgi cisternae may progressively mature
into trans-Golgi cisternae as in P. pastoris (Mogelsvang
et al., 2003). Figure 2 proposes several questions about the
organization of the ER–GA interface, and the interactions
between the different families of proteins (GTP-binding
proteins, coat proteins, SNAREs, etc.) that may drive the
specificic exchange of material between the ER and the
Another challenge for the future will be to unravel the
physical role of lipids and lipid/membrane-interacting
proteins in membrane domain formation and dynamics
related to membrane trafficking in plants. As highlighted by
McMahon and Gallop (2005), membrane deformation and
regulation of membrane curvature are fundamental events
that must be considered. Such physical deformations can be
managed by changes in lipid composition and asymmetry,
Fig. 2. Hypotheses and questions about the ER–Golgi interface. The
locations of the plant SNAREs Sec22, Memb11, Bet11, and SYP31 at the
ER–GA level are summarized. The retrograde COPI and anterograde
COPII pathways are also indicated at this interface. Beside the ERES (ER
export sites) that have been demonstrated to be associated with Golgi
bodies in plants (daSilva et al., 2004; Yang et al., 2005), it could be
speculated whether ERES are connected to specific cis-Golgi import sites
(cGIS) in the anterograde pathway. By analogy, cis-Golgi export sites
(cGES) in the cis-Golgi may act as export domains towards import sites in
the ER (ERIS) for the retrograde pathway. Do such domains for cargo
selection and membrane specific contacts exist? If yes, how are the
different families of proteins (GTP-binding proteins, SNAREs, COPI and
COPII coat proteins, etc) and lipids involved in the structural
organization of the ER–Golgi interface? Those are challenging questions
for the future.
The plant ER–Golgi interface59
by guest on November 6, 2015
integral membrane proteins, cytoskeletal proteins, scaffold-
ing by peripheral membrane proteins, and finally by active
helix insertion of membrane-associated proteins (Lee et al.,
2005; McMahon and Gallop, 2005). In addition, membrane
curvature may regulate the strength of association of per-
ipheral proteins, and membrane curvature-sensing pro-
teins may contribute to the architecture and dynamics of
membrane domain formation (De Matteis and Godi,
2004). In the case of lipid asymmetry, its loss in cis-Golgi
membranes may affect Golgi retrograde transport to the
ER by perturbing protein recruitment (Hua and Graham,
2003). Promising investigations in plants on interactions
between lipids and proteins of the traffic machineries have
begun to emerge from the literature (Jensen et al., 2000;
Lam et al., 2002).
The complexity and diversity of protein and lipid
machineries involved at the ER–GA level will undoubtedly
keep researchers busy for a long while.
AbousalhamA, HobmanTC, Dewald J,Garbutt M, Brindley DN.
2002. Cell-permeable ceramides preferentially inhibit coated
vesicle formation and exocytosis in Chinese hamster ovary
compared with Madin–Darby canine kidney cells by preventing
the membrane association of ADP-ribosylation factor. Biochemi-
cal Journal 361, 653–661.
AndreevaAV, Zheng H, Saint-Jore CM, KutuzovMA, Evans DE,
Hawes CR. 2000. Organization of transport from endoplasmic
reticulum to Golgi in higher plants. Biochemical Society Trans-
actions 28, 505–512.
Aoe T, Cukierman E, Lee A, Cassel D, Peters PJ, Hsu VW. 1997.
The KDEL receptor, ERD2, regulates intracellular traffic by
recruiting a GTPase-activating protein for ARF1. The EMBO
Journal 16, 7305–7316.
Bacia K, Schwille P, Kurzchalia T. 2005. Sterol structure
determines the separation of phases and the curvature of the
liquid-ordered phase in model membranes. Proceedings of the
National Academy of Sciences, USA 102, 3272–3277.
Barlowe C, Orci L, Yeung T, Hosobuchi M, Hamamoto S,
Salama N, Rexach MF, Ravazzola M, Amherdt M, Schekman
R. 1994. COPII: a membrane coat formed by Sec proteins that
drive vesicle budding from the endoplasmic reticulum. Cell 77,
Bar-Peled M, Raikhel NV. 1997. Characterization of AtSEC12 and
AtSAR1. Proteins likely involved in endoplasmic reticulum and
Golgi transport. Plant Physiology 114, 315–324.
Batoko H, Zheng HQ, Hawes C, Moore I. 2000. A rab1 GTPase
is required for transport between the endoplasmic reticulum and
Golgi apparatus and for normal Golgi movement in plants. The
Plant Cell 12, 2201–2218.
Belles-Boix E, Babiychuk E, Montagu MV, Inze ´ D, Kushnir S.
2000. CEF, a sec24 homologue of Arabidopsis thaliana, enhances
the survival of yeast under oxidative stress conditions. Journal of
Experimental Botany 51, 1761–1762.
Bessoule JJ, Moreau P. 2004. Phospholipid synthesis and dynamics
in plant cells. Topics in Current Genetics 6, 89–124.
Bevis BJ, Hammond AT, Reinke CA, Glick BS. 2002. De novo
formation of transitional ER sites and Golgi structures in Pichia
pastoris. Nature Cell Biology 4, 750–756.
Bielli A, Haney CJ, Gabreski G, Watkins SC, Bannykh SI,
Aridor M. 2005. Regulation of Sar1 NH2 terminus by GTP
binding and hydrolysis promotes membrane deformation to control
COPII vesicle fission. Journal of Cell Biology 171, 919–924.
Boevink P, Oparka K, Santa Cruz S, Martin B, Betteridge A,
Hawes C. 1998. Stacks on tracks: the plant Golgi apparatus traffics
on an actin/ER network. The Plant Journal 15, 441–447.
Borisovska M, Zhao Y, Tsytsyura Y, Glyvuk N, Takamori S,
Matti U, Rettig J, Su ¨dhof T, Bruns D. 2005. v-SNAREs control
exocytosis of vesicles from priming to fusion. The EMBO Journal
BornerGH, Sherrier DJ,
Hawkins ND, Macaskill A, Napier JA, Beale MH, Lilley KS,
Dupree P. 2005. Analysis of detergent-resistant membranes in
Arabidopsis. Evidence for plasma membrane lipid rafts. Plant
Physiology 137, 104–116.
Boutte ´Y, Crosnier M-T, Carraro N, Traas J,
Jeunemaitre B. 2006. Immunocytochemistry of the plasma mem-
brane recycling pathway and cell polarity in plants: studies on
PIN proteins. Journal of Cell Science 119, 1255–1265.
Brandizzi F, Frangne N, Marc-Martin S, Hawes C, Neuhaus JM,
Paris N. 2002a. The destination for single-pass membrane
proteins is influenced markedly by the length of the hydrophobic
domain. The Plant Cell 14, 1077–1092.
Brandizzi F, Fricker M, Hawes CR. 2002b. A greener world: the
revolution in plant bioimaging. Nature Reviews Molecular and
Cell Biology 3, 520–530.
Brandizzi F, Irons SL, Johansen J, Kotzer A, Neumann U. 2004.
GFP is the way to glow: bioimaging of the plant endomembrane
system. Journal of Microscopy 214, 138–158.
Brandizzi F, Snapp EL, Roberts AG, Lippincott-Schwartz J,
Hawes C. 2002c. Membrane protein transport between the
endoplasmic reticulum and the Golgi in tobacco leaves is energy
dependent but cytoskeleton independent: evidence from selective
photobleaching. The Plant Cell 14, 1293–1309.
Brown WJ, Chambers K, Doody A. 2003. Phospholipase A2
(PLA2) enzymes in membrane trafficking: mediators of membrane
shape and function. Traffic 4, 214–221.
Burri L, Lithgow T. 2004. A complete set of SNAREs in yeast.
Traffic 5, 45–52.
Burri L, Varlamov O, Doege CA, Hofmann K, Beilharz T,
Rothman JE, So ¨llner T, Lithgow T. 2003. A SNARE required
for retrograde transport to the endoplasmic reticulum. Proceedings
of the National Academy of Sciences, USA 100, 9873–9877.
Ceppi P, Colombo S, Francolini M, Raimondo F, Borgese N,
Masserini M. 2005. Two tail-anchored protein variants, differing
in transmembrane domain length and intracellular sorting, interact
differently with lipids. Proceedings of the National Academy of
Sciences, USA 102, 16269–16274.
Chatre L, Brandizzi F, Hocquellet A, Hawes C, Moreau P. 2005.
Sec22 and Memb11 are v-SNAREs of the anterograde endoplas-
mic reticulum–Golgi pathway in tobacco leaf epidermal cells.
Plant Physiology 139, 1244–1254.
Chen Y, Shin YK, Bassham DC. 2005. YKT6 is a core constituent
of membrane fusion machineries at the Arabidopsis trans-Golgi
network. Journal of Molecular Biology 350, 92–101.
Choukroun GJ, Marshansky V, Gustafson CE, McKee M,
Hajjar RJ, Rosenzweig A, Brown D, Bonventre JV. 2000.
Cytosolic phospholipase A2regulates Golgi structure and mod-
ulates intracellular trafficking of membrane proteins. Journal of
Clinical Investigation 106, 983–993.
Churchward MA, Rogasevskaia T, Ho ¨fgen J, Bau J, Coorssen JR.
Ca2+-triggered membrane fusion. Journal of Cell Science 118,
60 Moreau et al.
by guest on November 6, 2015
Constable JRL, Graham ME, Morgan A, Burgoyne RD. 2005.
Amisyn regulates exocytosis and fusion pore stability by both
syntaxin-dependent and syntaxin-independent mechanisms. Jour-
nal of Biological Chemistry 280, 31615–31623.
Contreras I, Ortiz-Zapater E, Aniento F. 2004a. Sorting signals in
the cytosolic tail of membrane proteins involved in the interaction
with plant ARF1 and coatomer. The Plant Journal 38, 685–698.
Contreras I, Yang Y, Robinson DG, Aniento F. 2004b. Sorting
signals in the cytosolic tail of plant p24 proteins involved in the
interaction with the COPII coat. Plant and Cell Physiology 45,
Cosson P, Ravazzola M, Varlamov O, So ¨llner TH, Di Liberto M,
Volchuk A, Rothman JE, Orci L. 2005. Dynamic transport of
SNARE proteins in the Golgi apparatus. Proceedings of the
National Academy of Sciences, USA 102, 14647–14652.
daSilva LL, Snapp EL, Denecke J, Lippincott-Schwartz J,
Hawes C, Brandizzi F. 2004. Endoplasmic reticulum export sites
and Golgi bodies behave as single mobile secretory units in plant
cells. The Plant Cell 16, 1753–1771.
De Figueiredo P, Drecktrah D, Katzenellenbogen JA, Strang M,
Brown WJ. 1998. Evidence that phospholipase A2activity is
required for Golgi complex and trans Golgi network membrane
tubulation. Proceedings of the National Academy of Sciences,
USA 95, 8642–8647.
De Matteis MA, Godi A. 2004. Protein–lipid interactions in mem-
brane trafficking at the Golgi complex. Biochimica et Biophysica
Acta 1666, 264–274.
Denecke J, Botterman J, Deblaere R. 1990. Protein secretion
in plant cells can occur via a default pathway. The Plant Cell
Denecke J, De Rycke R, Botterman J. 1992. Plant and mammalian
sorting signals for protein retention in the endoplasmic reticulum
contain a conserved epitope. The EMBO Journal 11, 2345–2355.
d’Enfert C, Gensse M, Gaillardin C. 1992. Fission yeast and
a plant have functional homologues of the Sar1 and Sec12 proteins
involved in ER to Golgi traffic in budding yeast. The EMBO
Journal 11, 4205–4211.
Drecktrah D, Chambers K, Racoosin EL, Cluett EB, Gucwa A,
Jackson B, Brown WJ. 2003. Inhibition of a Golgi complex
lysophospholipid acyltransferase induces membrane tubule forma-
tion and retrograde trafficking. Molecular Biology of the Cell 14,
Duden R, Presley J, Storrie B. 2005. The Golgi complex.
Biochimica et Biophysica Acta 1744, 257–258.
Dunn TM, Lynch DV, Michaelson LV, Napier JA. 2004. A post-
genomic approach to understanding sphingolipid metabolism in
Arabidopsis thaliana. Annals of Botany 93, 483–497.
Engelman DM. 2005. Membranes are more mosaic than fluid.
Nature 438, 578–580.
Fasshauer D, Sutton RB, Brunger AT, Jahn R. 1998. Conserved
structural features of the synaptic fusion complex: SNARE
proteins reclassified as Q- and R-SNAREs. Proceedings of the
National Academy of Sciences, USA 95, 15781–15786.
Freyberg Z, Bourgoin S, Shields D. 2002. Phospholipase D2 is
localized to the rims of the Golgi apparatus in mammalian cells.
Molecular Biology of the Cell 13, 3930–3942.
Fujioka S, Yokota T. 2003. Biosynthesis and metabolism of
brassinosteroids. Annual Review of Plant Biology 54, 137–164.
Fukunaga T, Nagahama M, Hatsuzawa K, Tani K, Yamamoto A,
Tagaya M. 2000. Implication of sphingolipid metabolism in the
stability of the Golgi apparatus. Journal of Cell Science 113,
Gardiner J, Collings DA, Harper JDI, Marc J. 2003. The effects
of the phospholipaseD-antagonist 1-butanol onseedling
development and microtubule organisation in Arabidopsis. Plant
and Cell Physiology 44, 687–696.
Geldner N, Anders N, Wolters H, Keicher J, Kornberger W,
Muller P, Delbarre A, Ueda T, Nakano A, JurgensG. 2003. The
Arabidopsis GNOM ARF-GEF mediates endosomal recycling,
auxin transport, and auxin-dependent plant growth. Cell 112,
Giraudo CG, Hu C, You D, Slovic AM, Mosharov EV, Sulzer D,
Melia TJ, Rothman JE. 2005. SNAREs can promote complete
fusion and hemifusion as alternative outcomes. Journal of Cell
Biology 170, 249–260.
Glick BS. 2002. Can the Golgi form de novo. Nature Reviews
Molecular Cell Biology 3, 615–619.
Gonzales Jr LC, Weis WI, Scheller RH. 2001. A novel SNARE
N-terminal domain revealed by the crystal structure of Sec22b.
Journal of Biological Chemistry 276, 24203–24211.
Graf CT, Riedel D, Schmitt HD, Jahn R. 2005. Identification of
functionally interacting SNAREs by using complementary sub-
stitutions in the conserved ‘0’ layer. Molecular Biology of the Cell
Grimmer S, Ying M, Wa ¨lchli S, van Deurs B, Sandvig K. 2005.
Golgi vesiculation induced by cholesterol occurs by a dynamin-
and cPLA2-dependent mechanism. Traffic 6, 144–156.
Hanton SL, Renna L, Bortolotti LE, Chatre L, Stefano G,
Brandizzi F. 2005. Diacidic motifs influence the export of trans-
membrane proteins from the endoplasmic reticulum in plant cells.
The Plant Cell 17, 3081–3093.
Hartmann MA, Perret AM, Carde JP, Cassagne C, Moreau P.
2002. Inhibition of the sterol pathway in leek seedlings impairs
phosphatidylserine and glucosylceramide synthesis but triggers an
accumulation of triacylglycerols. Biochimica et Biophysica Acta
Hasegawa H, Zinser S, Rhee Y, Vik-Mo EO, Davanger S, Hay
JC. 2003. Mammalian YKT6 is a neuronal SNARE targeted to
a specialized compartment by its profilin-like amino terminal
domain. Molecular Biology of the Cell 14, 698–720.
Hasegawa H, Yang Z, Oltedal L, Davanger S, Hay JC. 2004.
Intramolecular protein–protein and protein–lipid interactions con-
trol the conformation and subcellular targeting of neuronal Ykt6.
Journal of Cell Science 117, 4495–4508.
Hawes C. 2005. Cell biology of the plant Golgi apparatus. New
Phytologist 165, 29–44.
Hawes C, Saint-Jore C, Martin B, Zheng HQ.2001. ER confirmed
as the location of mystery organelles in Arabidopsis plants
expressing GFP! Trends in Plant Science 6, 245–246.
Hawes C, Satiat-Jeunemaitre B. 2005. The plant Golgi apparatus—
going with the flow. Biochimica et Biophysica Acta 1744, 93–107.
Heese-Peck A, Pichler H, Zanolari B, Watanabe R, Daum G,
Riezman H. 2002. Multiple functions of sterols in yeast endocy-
tosis. Molecular Biology of the Cell 13, 2664–2680.
Herpers B, Rabouille C. 2004. mRNA localization and ER-based
protein sorting mechanisms dictate the use of tER–Golgi units
involved in gurken transport in Drosophila oocytes. Molecular
Biology of the Cell 15, 5306–5317.
Honda A, Al-Awar O, Hay JC, Donaldson JG. 2005. Targeting of
ARF-1 to the early Golgi membrin, an ER–Golgi SNARE. Journal
of Cell Biology 168, 1039–1051.
Hong W. 2005. SNAREs and traffic. Biochimica et Biophysica Acta
Hu C, Ahmed M, Melia TJ, Sollner TH, Mayer T, Rothman JE.
2005a. Fusion of cells by flipped SNAREs. Science 300,
Hu W, Xu R, Zhang G, Jin J,Szulc ZM, Bielawski J, Hannun YA,
Obeid LM, Mao C. 2005b. Golgi fragmentation is associated
The plant ER–Golgi interface 61
by guest on November 6, 2015
with ceramide-induced cellular effects. Molecular Biology of the
Cell 16, 1555–1567.
Hua Z, Graham TR. 2003. Requirement for Neo1p in retrograde
transport from the Golgi complex to the endoplasmic reticulum.
Molecular Biology of the Cell 14, 4971–4983.
Jensen RB, Lykke-Andersen K, Frandsen GI, Nielsen HB,
Haseloff J, Jespersen HM, Mundy J, Skriver K. 2000. Pro-
miscuous and specific phospholipids binding by domains in
ZAC, a membrane-associated Arabidopsis protein with an ARF
GAP zinc finger and a C2 domain. Plant Molecular Biology 44,
Joglekar AP, Xu D, Rigotti DJ, Fairman R, Hay JC. 2003. The
SNARE motif contributes to rbet1 intracellular targeting and
dynamics independently of SNARE interactions. Journal of
Biological Chemistry 278, 14121–14133.
Jouannic N, Lepetit M, Vergnolle C, Cantrel C, Gardies AM,
Kader JC, Arondel V. 1998. Isolation of a cDNA from
Arabidopsis thaliana that complements the sec14 mutant of yeast.
European Journal of Biochemistry 258, 402–410.
Ju ¨rgens G. 2005. Cytokinesis in higher plants. Annual Review of
Plant Biology 56, 281–299.
Kearns BG, McGee TP, Mayinger P, Gedvilaite A, Phillips SE,
Kagiwada S, Bankaitis VA. 1997. Essential role for diacylglycer-
ol in protein transport from the yeast Golgi complex. Nature 387,
Ke ´pe `s F, Rambourg A, Satiat-Jeunemaitre B. 2005. Morphody-
namics of the secretory pathway. International Review of Cytology
Kooijman EE, Chupin V, de Kruijff B, Burger KNJ. 2003.
Modulation of membrane curvature by phosphatidic acid and
lysophosphatidic acid. Traffic 4, 162–174.
Lam BCH, Sage TL, Bianchi F, Blumwald E. 2002. Regulation of
ADL6 activity by its associated molecular network. The Plant
Journal 31, 565–576.
Latijnhouwers M, Hawes C, Carvalho C. 2005. Holding it all
together? Candidate proteins for the plant Golgi matrix. Current
Opinion in Plant Biology 8, 1–8.
Lee MCS, Orci L, Hamamoto S, Futai E, Ravazzola M,
Schekman R. 2005. Sar1p N-terminal helix initiates membrane
curvature and completes the fission of a COPII vesicle. Cell
Lee MH, Min MK, Lee YJ, Jin JB, Shin DH, Kim DH, Lee KH,
Hwang I. 2002. ADP-ribosylation factor 1 of Arabidopsis plays
a critical role in intracellular trafficking and maintenance of
endoplasmic reticulum morphology in Arabidopsis. Plant Physi-
ology 129, 1507–1520.
Lewis MJ, Pelham HR. 1992. Ligand-induced redistribution of
a human KDEL receptor from the Golgi complex to the
endoplasmic reticulum. Cell 68, 353–364.
Lewis MJ, Sweet DJ, Pelham HR. 1990. The ERD2 gene
determines the specificity of the luminal ER protein retention
system. Cell 61, 1359–1363.
LiX,RivasMP,Fang M,Marchena J,MehrotraB,ChaudharyA,
Feng L, Prestwich GD, Bankaitis VA. 2002. Analysis of
oxysterol binding protein homologue Kes1p function in regulation
of Sec14p-dependent protein transport from the yeast Golgi
complex. Journal of Cell Biology 157, 63–78.
Lisman Q, Pomorski T, Vogelzangs C, Urli-Stam D, de Cocq van
Delwijneen W, Holthuis JCM. 2004. Protein sorting in the late
Golgi of Saccharomyces cerevisiae does not require mannosylated
sphingolipids. Journal of Biological Chemistry 279, 1020–1029.
Litvak V, Dahan N, Ramachandran S, Sabanay H, Lev S. 2005.
Maintenance of the diacylglycerol level in the Golgi apparatus by
the Nir2 protein is critical for Golgi secretory function. Nature Cell
Biology 7, 225–234.
Liu Y, Barlowe C. 2002. Analysis of sec22p in endoplasmic
reticulum/Golgi transport reveals cellular redundancy in SNARE
protein function. Molecular Biology of the Cell 13, 3314–3324.
Liu Y, Flanagan JJ, Barlowe C. 2004. Sec22p export from the
endoplasmic reticulum is independent of SNARE pairing. Journal
of Biological Chemistry 279, 27225–27232.
Loranger SS, Linder ME. 2002. SNAP25 traffics to the plasma
membrane by a syntaxin-independent mechanism. Journal of
Biological Chemistry 277, 34303–34309.
Matsuoka K, Orci L, Amherdt M, Bednarek SY, Hamamoto S,
Schekman R, Yeung T. 1998. COPII-coated vesicle formation
reconstituted with purified coat proteins and chemically defined
liposomes. Cell 93, 263–275.
McMahon HT, Gallop JL. 2005. Membrane curvature and mech-
anisms of dynamic cell membrane remodelling. Nature 438,
Me ´rigout P, Ke ´pe `s F, Perret AM, Satiat-Jeunemaitre B,
Moreau P. 2002. Effects of brefeldin A and nordihydroguaiaretic
acid on endomembrane dynamics and lipid synthesis in plant
cells. FEBS Letters 518, 88–92.
Miller EA, Beilharz TH, Malkus PN, Lee MCS, Hamamoto S,
Orci L, Schekman R. 2003. Multiple cargo binding sites on the
COPII subunit Sec24p ensure capture of diverse membrane
proteins into transport vesicles. Cell 114, 497–509.
Miller EA, Liu Y, Barlowe C, Schekman R. 2005. ER–Golgi
transport defects are associated with mutations in the Sed5p-
binding domain of the COPII coat subunit, Sec24p. Molecular
Biology of the Cell 16, 3719–3726.
Mogelsvang S, Gomez-Ospina N, Soderholm J, Glick BS,
Staehelin LA. 2003. Tomographic evidence for continuous
turnover of Golgi cisternae in Pichia pastoris. Molecular Biology
of the Cell 14, 2277–2291.
Mongrand S, Morel J, Laroche J, Claverol S, Carde JP,
Hartmann MA, Bonneu M, Simon-Plas F, Lessire R,
Bessoule JJ. 2004. Lipid rafts in higher plant cells: purification
and characterization of Triton X-100-insoluble microdomains
from tobacco plasma membrane. Journal of Biological Chemistry
Monteiro D, Liu Q, Lisboa S, Scherer GEF, Quader H, Malho R.
2005. Phosphoinositides and phosphatidic acid regulate pollen
tube growth and reorientation through modulation of [Ca2+]c
and membrane secretion. Journal of Experimental Botany 416,
Moreau P, Bessoule JJ, Mongrand S, Testet E, Vincent P,
Cassagne C. 1998. Lipid trafficking in plant cells. Progress in
Lipid Research 37, 371–391.
Moreau P, Cassagne C, Keenan TW, Morre ´ DJ. 1993. Ceramide
excluded from cell-free vesicular lipid transfer from endoplasmic
reticulum to Golgi apparatus. Evidence for lipid sorting. Biochi-
mica et Biophysica Acta 1146, 9–16.
Moreau P, Juguelin H, Cassagne C, Morre ´ DJ. 1992. Molecular
basis for low temperature compartment formation by transitional
endoplasmic reticulum of rat liver. FEBS Letters 310, 223–228.
Mossessova E, Bickford LC, Goldberg J. 2003. SNARE selectivity
of the COPII coat. Cell 114, 483–495.
Movafeghi A, Happel N, Pimpl P, Tai GH, Robinson DG. 1999.
Arabidopsis Sec21p and Sec23p homologs. Probable coat proteins
of plant COP-coated vesicles. Plant Physiology 119, 1437–1446.
Munro S. 2005. The Golgi apparatus: defining the identity of Golgi
membranes. Current Opinion in Cell Biology 17, 395–401.
Murshid A, Presley JF. 2004. ER-to-Golgi transport and cytoskel-
etal interactions in animal cells. Cellular and Molecular Life
Sciences 61, 133–145.
Nakajima KI, Sonoda H, Mizoguchi T, Aoki J, Arai H,
Nagahama M, Tagaya M, Tani K. 2002. A novel phospholipase
62 Moreau et al.
by guest on November 6, 2015
A1with sequence homology to a mammalian Sec23p-interacting
protein, p125. Journal of Biological Chemistry 277, 11329–11335.
Nakamura M, Kuroiwa N, Kono Y, Takatsuki A. 2001. Gluco-
sylceramide synthesis inhibitors block pharmacologically induced
dispersal of the Golgi and anterograde membrane flow from the
endoplasmic reticulum: implication of sphingolipid metabolism in
maintenance of the Golgi architecture and anterograde membrane
flow. Biosciences Biotechnology Biochemistry 65, 1369–1378.
Nebenfu ¨hr A, Gallagher LA, Dunahay TG, Frohlick JA,
Mazurkiewicz AM, Meehl JB, Staehelin LA. 1999. Stop-and-
go movements of plant Golgi stacks are mediated by the acto-
myosin system. Plant Physiology 121, 1127–1142.
Neumann U, Brandizzi F, Hawes CR. 2003. Protein transport in
plant cells: in and out of the Golgi. Annals of Botany 92, 167–180.
Nicchitta CV. 2002. A platform for compartmentalized protein
synthesis: protein translation and translocation in the ER. Current
Opinion in Cell Biology 14, 412–416.
Niihama M, Uemura T, Saito C, Nakano A, Sato MH, Tasaka M,
Morita MT. 2005. Conversion of functional specificity in Qb-
SNARE VTI1 homologues of Arabidopsis. Current Biology 15,
Palmer DJ, Helms JB, Beckers CJ, Orci L, Rothman JE.
1993. Binding of coatomer to Golgi membranes requires ADP-
ribosylation factor. Journal of Biological Chemistry 268,
Pathre P, Shome K, Blumental-Perry A, Bielli A, Haney CJ,
Alber S, Watkins SC, Romero G, Aridor M. 2003. Activation
of phospholipase D by the small GTPase Sar1p is required to
support COPII assembly and ER export. The EMBO Journal 22,
Peng R, Gallwitz D. 2004. Multiple SNARE interactions of an SM
protein: Sed5p/Sly1p binding is dispensable for transport. The
EMBO Journal 23, 3939–3949.
Peng R, Grabowski R, De Antoni A, Gallwitz D. 1999. Specific
interaction of the yeast cis-Golgi syntaxin Sed5p and the coat
protein complex II component Sec24p of endoplasmic reticulum-
derived transport vesicles. Proceedings of the National Academy
of Sciences, USA 96, 3751–3756.
Phillipson BA, Pimpl P, daSilva LL, Crofts AJ, Taylor JP,
Movafeghi A, Robinson DG, Denecke J. 2001. Secretory bulk
flow of soluble proteins is efficient and COPII dependent. The
Plant Cell 13, 2005–2020.
Pimpl P, Hanton SL, Taylor JP, Pinto-DaSilva LL, Denecke J.
2003. The GTPase ARF1p controls the sequence-specific vacuolar
sorting route to the lytic vacuole. The Plant Cell 15, 1242–1256.
Pimpl P, Movafeghi A, Coughlan S, Denecke J, Hillmer S,
Robinson DG. 2000. In situ localization and in vitro induction of
plant COPI-coated vesicles. The Plant Cell 12, 2219–2236.
Potocky M, Elias M, Profotova B, Novotna Z, Valentova O,
Zarsky V. 2003. Phosphatidic acid produced by phospholipase D
is required for tobacco pollen tube growth. Planta 217, 122–130.
Pratelli R, Sutter JU, Blatt MR. 2004. A new catch in the SNARE.
Trends in Plant Science 9, 187–195.
Randazzo PA. 1997. Resolution of two ADP-ribosylation factor 1
GTPase-activating proteins from rat liver. Biochemical Journal
Reese C, Heise F, Mayer A. 2005. Trans-SNARE pairing can
precede a hemifusion intermediate in intracellular membrane
fusion. Nature 436, 410–414.
Rein U, Andag U, Duden R, Schmitt HD, Spang A. 2002. ARF-
GAP-mediated interaction between the ER–Golgi v-SNAREs and
the COPI coat. Journal of Cell Biology 157, 395–404.
Ritzenthaler C, Nebenfu ¨hr A, Movafeghi A, Stussi-Garaud C,
Behnia L, Pimpl P, Staehelin LA, Robinson DG. 2002.
Reevaluation of the effects of brefeldin A on plant cells using
tobacco Bright Yellow 2 cells expressing Golgi-targeted green
fluorescent protein and COPI antisera. The Plant Cell 14, 237–261.
Rossi V, Banfield DK, Vacca M, Dietrich LEP, Ungermann C,
D’Esposito M, Galli T, Filippini F. 2004b. Longins and their
longin domains: regulated SNAREs and multifunctional SNARE
regulators. Trends in Biochemical Sciences 29, 682–688.
Rossi V, Picco R, Vacca M, D’Esposito M, D’Urso M, Galli T,
Filippini F. 2004a. VAMP subfamilies identified by specific
R-SNARE motifs. Biology of the Cell 96, 251–256.
Routt SM, Ryan MM, Tyeryar K, Rizzieri KE, Mousley C,
Roumanie O, Brennwald PJ, Bankaitis VA. 2005. Nonclassical
PITPs activate PLD via the Stt4p PtdIns-4-kinase and modulate
function of late stages of exocytosis in vegetative yeast. Traffic
Runions J, Brach T, Kuhner S, Hawes C. 2006. Photoactivation of
GFP reveals protein dynamics within the endoplasmic reticulum
membrane. Journal of Experimental Botany 57, 43–50.
Saint-Jore CM, Evins J, Batoko H, Brandizzi F, Moore I,
Hawes C. 2002. Redistribution of membrane proteins between
the Golgi apparatus and endoplasmic reticulum in plants is re-
versible and not dependent on cytoskeletal networks. The Plant
Journal 29, 661–678.
Salau ¨n C, Gould GW, Chamberlain LH. 2005a. The SNARE
proteins SNAP-25 and SNAP-23 display different affinities for
lipid rafts in PC12 cells. Journal of Biological Chemistry 280,
Salau ¨n C, Gould GW, Chamberlain LH. 2005b. Lipid raft
association of SNARE proteins regulates exocytosis in PC12 cells.
Journal of Biological Chemistry 280, 19449–19453.
Sanderfoot AA, Assaad FF, Raikhel NV. 2000. The Arabidopsis
genome. An abundance of soluble N-ethylmaleimide-sensitive
factor adaptor protein receptors. Plant Physiology 124, 1558–1569.
Sanderfoot AA, Kovaleva V, Bassham DC, Raikhel NV. 2001.
Interactions between syntaxins identify at least five SNARE
complexes within the Golgi/prevacuolar system of the Arabidopsis
cell. Molecular Biology of the Cell 12, 3733–3743.
Satiat-Jeunemaitre B, Boevink P, Hawes C. 1999. Membrane
trafficking in higher plant cells: GFP and antibodies, partners for
probing the secretory pathway. Biochimie 81, 597–605.
Satiat-Jeunemaitre B, Cole L, Bourett T, Howard R, Hawes C.
1996. Brefeldin A effects in plant and fungal cells: something new
about vesicle trafficking? Journal of Microscopy 181, 162–177.
Scales SJ, Hesser BA, Masuda ES, Scheller RH. 2002. Amysin, a
novel syntaxin-binding protein that may regulate SNARE complex
assembly. Journal of Biological Chemistry 277, 28271–28279.
Schaeffer A, Bronner R, Benveniste P, Schaller H. 2001. The ratio
of campesterol to sitosterol which modulates growth in Arabidop-
sis is controlled by STEROL METHYLTRANSFERASE 2-1. The
Plant Journal 25, 605–615.
Schaller H. 2003. The role of sterols in plant growth and
development. Progress in Lipid Research 42, 163–175.
Schaller H. 2004. New aspects of sterol biosynthesis in growth and
development of higher plants. Plant Physiology and Biochemistry
Schrick K, Mayer U, Martin G, Bellini C, Kuhnt C, Schmidt J,
Ju ¨rgens G. 2002. Interactions between sterol biosynthesis genes
in embryonic development of Arabidopsis. The Plant Journal 31,
Semenza JC, Hardwick KG, Dean N, Pelham HR. 1990. ERD2,
a yeast gene required for the receptor-mediated retrieval of luminal
ER proteins from the secretory pathway. Cell 61, 1349–1357.
Shimoi W, Ezawa I, Nakamoto K, Uesaki S, Gabreski G, Aridor
M, YamamotoA,NagahamaM, Agaya M,TaniK.2005. p125is
localized in endoplasmic reticulum exit sites and involved in their
organization. Journal of Biological Chemistry 280, 10141–10148.
The plant ER–Golgi interface 63
by guest on November 6, 2015
Siddhanta A, Backer JM, Shields D. 2000. Inhibition of phos- Download full-text
phatidic acid synthesis alters the structure of the Golgi apparatus
and inhibits secretion in endocrine cells. Journal of Biological
Chemistry 275, 12023–12031.
Sperling P, Warnecke D, Heinz E. 2004. Plant sphingolipids.
Topics in Current Genetics 6, 337–381.
Staneva G, Angelova MI, Koumanov K. 2004. Phospholipase A2
promotes raft budding and fission from giant liposomes. Chemistry
and Physics of Lipids 129, 53–62.
Stefano G, Renna L, Chatre L, Hanton SL, Moreau P, Hawes C,
Brandizzi F. 2006. In tobacco leaf epidermal cells, the integrity of
ER export sites and of protein export from the endoplasmic
reticulum in plant cells depends on active COPI machinery. The
Plant Journal, 46, 95–1110.
Sturbois-Balcerzak B, Vincent P, Maneta-Peyret L, Duvert M,
Satiat-Jeunemaitre B, Cassagne C, Moreau P. 1999. ATP-
dependent formation of phosphatidylserine-rich vesicles from
the endoplasmic reticulum of leek cells. Plant Physiology 120,
Surpin M, Raikhel N. 2004. Traffic jams affect plant development
and signal transduction. Nature Reviews Molecular Cell Biology
Szafer E, Pick E, Rotman M, Zuck S, Huber I, Cassel D. 2000.
Role of coatomer and phospholipids in GTPase-activating protein-
dependent hydrolysis of GTP by ADP-ribosylation factor-1.
Journal of Biological Chemistry 275, 23615–23619.
Szafer E, Rotman M, Cassel D. 2001. Regulation of GTP
hydrolysis on ADP-ribosylation factor-1 at the Golgi membrane.
Journal of Biological Chemistry 276, 47834–47839.
Tai WCS, Banfield DK. 2001. AtBS14a and AtBS14b, two Bet1/
Sft1-like SNAREs from Arabidopsis thaliana that complement
mutations in the yeast SFT1 gene. FEBS Letters 500, 177–182.
Takeuchi M, Tada M, Saito C, Yashiroda H, Nakano A.
1998. Isolation of a tobacco cDNA encoding Sar1 GTPase and
analysis of its dominant mutations in vesicular traffic using a
yeast complementation system. Plant and Cell Physiology 39,
Takeuchi M, Ueda T, Sato K, Abe H, Nagata T, Nakano A. 2000.
A dominant negative mutant of sar1 GTPase inhibits protein
transport from the endoplasmic reticulum to the Golgi apparatus in
tobacco and Arabidopsis cultured cells. The Plant Journal 23,
Takeuchi M, Ueda T, Yahara N, Nakano A. 2002. Arf1 GTPase
plays roles in the protein traffic between the endoplasmic reticulum
and the Golgi apparatus in tobacco and Arabidopsis cultured cells.
The Plant Journal 31, 499–515.
Thiele C, Hannah MJ, Fahrenholz F, Huttner WB. 2000.
Cholesterol binds to synaptophysin and is required for biogenesis
of synaptic vesicles. Nature Cell Biology 2, 42–49.
Trombetta ES, ParodiAJ. 2003. Quality control and protein folding
in the secretory pathway. Annual Review of Cell and Develop-
mental Biology 19, 649–676.
Uemura T, Sato MH, Takeyasu K. 2005. The longin domain
regulates subcellular targeting of VAMP7 in Arabidopsis thaliana.
FEBS Letters 579, 2842–2846.
Uemura T, Ueda T, Ohniwa RL, Nakano A, Takeyasu K,
Sato MH. 2004. Systematic analysis of SNARE molecules in
Arabidopsis: dissection of the post-Golgi network in plant cells.
Cell Structure and Function 29, 49–65.
Ungermann C, Langosh D. 2005. Functions of SNAREs in
intracellular membrane fusion and lipid bilayer mixing. Journal
of Cell Science 118, 3819–3828.
Varlamov O, Volchuk A, Rahimian V, et al. 2004. i-SNAREs:
inhibitory SNAREs that fine-tune the specificity of membrane
fusion. Journal of Cell Biology 164, 79–88.
Villarejo A, Buren S, Larsson S, et al. 2005. Evidence for a protein
transported through the secretory pathway en route to the higher
plant chloroplast. Nature Cell Biology 7, 1124–1131.
Vincent P, Chua M, Nogue F, Fairbrother A, Mekeel H, Xu Y,
Allen N, Bibikova TN, Gilroy S, Bankaitis VA. 2005. A sec14p-
nodulin domain phosphatidylinositol transfer protein polarizes
membrane growth of Arabidopsis thaliana root hairs. Journal of
Cell Biology 168, 801–812.
Vincent P, Maneta-Peyret L, Cassagne C, Moreau P. 2001.
Phosphatidylserine delivery to endoplasmic reticulum-derived
vesicles of plant cells depends on two biosynthetic pathways.
FEBS Letters 498, 32–36.
Vitale A, Denecke J. 1999.The endoplasmic reticulum—gateway of
the secretory pathway. The Plant Cell 11, 615–628.
Wang Y, Thiele C, Huttner WB. 2000. Cholesterol is required for
the formation of regulated and constitutive secretory vesicles from
the trans-Golgi network. Traffic 1, 952–962.
protein complex containing subunits of non-clathrin-coated Golgi
transport vesicles. Nature 349, 248–251.
Weber T, Zemelman BV, McNew JA, Westermann B, Gmachl M,
Parlati F, Sollner TH, Rothman JE. 1998. SNAREpins: minimal
machinery for membrane fusion. Cell 92, 759–772.
Weinberger A, Kamena F, Kama R, Spang A, Gerst JE. 2005.
Control of Golgi morphology and function by sed5 t-SNARE
phosphorylation. Molecular Biology of the Cell 16, 4918–4930.
Willemsen V, Friml J, Grebe M, van den Toorn A, Palme K,
Scheres B. 2003. Cell polarity and PIN protein positioning in
Arabidopsis require STEROL METHYLTRANSFERASE 1 func-
tion. The Plant Cell 15, 612–625.
Wyles JP, McMaster CR, Ridgway ND. 2002. VAMP-associated
protein-A (VAP-A) interacts with the oxysterol binding protein
(OSBP) to modify export from the endoplasmic reticulum. Journal
of Biological Chemistry 277, 29908–29918.
Wyles JP, Ridgway ND. 2004. VAMP-associated protein-A regu-
lates partitioning of oxysterol-binding protein-related protein-9
between the endoplasmic reticulum and Golgi apparatus. Experi-
mental Cell Research 297, 533–547.
Xu J, Scheres B. 2005. Dissection of Arabidopsis ADP-RIBOSY-
LATION FACTOR 1 function in epidermal cell polarity. The
Plant Cell 17, 525–536.
Xu Y, Liu Y, Ridgway ND, McMaster CR. 2001. Novel members
of the human oxysterol-binding protein family bind phospholipids
and regulate vesicle transport. Journal of Biological Chemistry
Xu Y, Zhang F, Su Z, McNew JA, Shin YK. 2005. Hemifusion in
SNARE-mediated membrane fusion. Nature Structural and Mol-
ecular Biology 12, 417–422.
Yang YD, Elamawi R, Bubeck J, Pepperkok R, Ritzenthaler C,
Robinson DG. 2005. Dynamics of COPII vesicles and the Golgi
apparatus in cultured Nicotiana tabacum BY-2 cells provides
evidence for transient associationof Golgi stacks with endoplasmic
reticulum exit sites. The Plant Cell 17, 1513–1531.
Yoshihisa T, Barlowe C, Schekman R. 1993. Requirement for
a GTPase-activating protein in vesicle budding from the endo-
plasmic reticulum. Science 259, 1466–1468.
Yuasa K, Toyooka K, Fukuda H, Matsuoka K. 2005. Membrane-
anchored prolyl hydroxylase with an export signal from the
endoplasmic reticulum. The Plant Journal 41, 81–94.
64 Moreau et al.
by guest on November 6, 2015