Plasma Membrane Localization of Solanum tuberosum
Remorin from Group 1, Homolog 3 Is Mediated by
Conformational Changes in a Novel C-Terminal Anchor
and Required for the Restriction of Potato Virus
X Movement1[C][W]
Artemis Perraki, Jean-Luc Cacas, Jean-Marc Crowet, Laurence Lins, Michel Castroviejo,
Sylvie German-Retana, Sébastien Mongrand 2*, and Sylvain Raffaele 2
Centre National de la Recherche Scientifique, Université de Bordeaux, 33076 Bordeaux cedex, France (A.P.,
J.-L.C., M.C., S.M., S.R.); The Sainsbury Laboratory, John Innes Centre, Norwich NR4 7UH, United Kingdom
(S.R.); Centre de Biophysique Moléculaire Numérique, Université de Liège, B-5030 Gembloux, Belgium
(J.-M.C., L.L.); and Equipe de Virologie, Institut Scientifique de Recherche Agronomique and Université de
Bordeaux, BP81, 33883 Villenave d’Ornon cedex, France (S.G.-R.)
The formation of plasma membrane (PM) microdomains plays a crucial role in the regulation of membrane signaling and
trafficking. Remorins are a plant-specific family of proteins organized in six phylogenetic groups, and Remorins of group
1 are among the few plant proteins known to specifically associate with membrane rafts. As such, they are valuable to
understand the molecular bases for PM lateral organization in plants. However, little is known about the structural
determinants underlying the specific association of group 1 Remorins with membrane rafts. We used a structure-function
approach to identify a short C-terminal anchor (RemCA) indispensable and sufficient for tight direct binding of potato
(Solanum tuberosum) REMORIN 1.3 (StREM1.3) to the PM. RemCA switches from unordered to a-helical structure in a
nonpolar environment. Protein structure modeling indicates that RemCA folds into a tight hairpin of amphipathic helices.
Consistently, mutations reducing RemCA amphipathy abolished StREM1.3 PM localization. Furthermore, RemCA directly
binds to biological membranes in vitro, shows higher affinity for Detergent-Insoluble Membranes lipids, and targets yellow
fluorescent protein to Detergent-Insoluble Membranes in vivo. Mutations in RemCA resulting in cytoplasmic StREM1.3
localization abolish StREM1.3 function in restricting potato virus X movement. The mechanisms described here provide new
insights on the control and function of lateral segregation of plant PM.
Protein-lipid interactions are increasingly recognized
as key regulatory processes for signal perception and
cellular signaling cascades (Cho and Stahelin, 2005).
During signal transduction and trafficking, a number
of soluble proteins dynamically associate with plasma
membranes (PMs) to deliver their cargo and to recruit
1
This work was supported by a Marie Curie Intra-European
Fellowship (contract no. 255104 to S.R.) and the French Agence
Nationale pour la Recherche (contract no. NT09_517917 PANACEA
to S.M. and J.-L.C.). L.L. is senior research assistant at the Belgian
National Funds for Research. L.L. and J.-M.C. received financial
support from the Interuniversity Attraction Poles project.
2
These authors contributed equally to the article.
* Co rr e s po n d in g au t ho r; e - ma i l s e b as t i e n .mo ng r an d @
u-bordeaux2.fr.
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is:
Sébastien Mongrand ([email protected]).
[C]
Some figures in this article are displayed in color online but in
black and white in the print edition.
[W]
The online version of this article contains Web-only data.
www.plantphysiol.org/cgi/doi/10.1104/pp.112.200519
624
pathway components to the sites of action (Seong et al.,
2011). For such proteins, membrane association can be
critical for function (Porter and Koelle, 2010).
PM targeting of peripheral proteins is achieved
through (1) binding to integral membrane proteins, (2)
posttranslational modifications, or (3) directly by intrinsic membrane anchor domains. Posttranslational
modifications function as auxiliary modifications for
transient or weak association of soluble proteins to
the intracellular face of the PM. In plants, these include
N-myristoylation, S-palmitoylation, prenylation by farnesyl or geranylgeranyl moieties, or attachment of glycosylphosphatidylinositol (GPI) anchors (Thompson and
Okuyama, 2000). GPI anchors, for example, tightly
associate proteins to the extracellular face of PMs by
interaction of the inositol head group of the membrane
lipid phosphatidylinositol with a glucosamine residue
linked to the C-terminal amino acid of the protein
(Paulick and Bertozzi, 2008). As an alternative mechanism, globular structures either recognize phospholipids in a stereospecific manner or associate with
membranes by their biophysical properties (for review,
see Lemmon, 2008). Other proteins expose unstructured
clusters of basic and hydrophobic residues to mediate
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Membrane Anchor of StREM1.3 Remorin
PM binding (McLaughlin et al., 2002; McLaughlin and
Murray, 2005).
Selective recognition of membrane compartments or
domains by protein anchors can be critical in triggering
the appropriate downstream trafficking and signaling
events (for review, see Gruenberg, 2003; De Matteis and
Godi, 2004). Membrane domain selectivity can be specified by the anchoring posttranslational modification or
by a protein anchor domain. For instance, proteins carrying GPI anchors are overrepresented in membrane
rafts, indicating that addition of this lipid anchor directs
proteins to these microdomains (Cordy et al., 2003;
Kierszniowska et al., 2009). Membrane rafts are enriched
in highly saturated long-chain sphingolipids, sterols,
and saturated phospholipids, creating tightly packed
domains, designated as “liquid ordered.” These lipids
display a stronger affinity to saturated acyl chains as
found in GPI-anchored and acylated proteins (Brown,
2006). The composition of membrane rafts also prevents
solubilization by detergent at low temperature with
nonionic detergent and allows the partial purification of
rafts in so-called Detergent-Insoluble Membrane (DIM)
fractions, which are supposedly biochemical counterparts of membrane rafts. Many signaling proteins are
found in membrane rafts, supporting the hypothesis
that they serve as key platforms for cellular signal
transduction and cell-to-cell communication (Lingwood
and Simons, 2010; Simon-Plas et al., 2011). For example,
in human (Homo sapiens) cells, key soluble signaling
components such as the Ser/Thr kinase Akt (protein
kinase B) are recruited to membrane rafts where they
activate signal transduction cascades (Lasserre et al.,
2008). Nevertheless, few protein motifs were described
to contribute to raft targeting (Rossin et al., 2010), although the 6-amino-acid-long raft target signal from
human Tyr phosphatase Src homology 2-containing
phosphatase1 is the only known motif sufficient to anchor soluble proteins specifically to domains of the intracellular face of the PM (Sankarshanan et al., 2007).
However, the anchoring mechanism itself remains to
be unraveled in plants even though DIMs also exist
(Mongrand et al., 2010) and functional PM domains
have been reported (Bhat et al., 2005). The molecular
basis for specific targeting and binding of proteins to
membrane rafts has never been described.
Remorins form a diverse family of plant-specific
proteins organized in six distinct phylogenetic groups
(Raffaele et al., 2007). Remorins from group 1 have been
reported to localize to the PM despite their overall hydrophilic nature (Reymond et al., 1996; Raffaele et al.,
2007). Moreover, group 1 Remorins almost exclusively
associate to DIMs and localize to membrane microdomains in a sterol-dependent manner (Lefebvre et al.,
2007; Kierszniowska et al., 2009; Raffaele et al., 2009).
The function of Remorins is mostly unknown, but we
showed in a previous study that StREM1.3 (for potato
(Solanum tuberosum) Remorin from group 1, homolog 3;
initially described in Reymond et al., 1996) regulates
cell-to-cell propagation of the potato virus X (PVX),
likely by directly interacting with the viral movement
protein Triple Gene Block protein1 (TGBp1; Raffaele
et al., 2009). StREM1.3 localizes to the inner leaflet of
PMs and along plasmodesmata, bridges connecting
neighbor cells essential for cell-to-cell communication
in plants (Maule, 2008). Other members of the Remorin
family group 1 are likely involved in innate immune
responses (Liu et al., 2009; Widjaja et al., 2009; Keinath
et al., 2010). Remorins from group 2 are involved in
the control of infection by symbiotic bacteria at nodular infection threads and the peribacteroid membrane
(Lefebvre et al., 2010) These data suggest general roles
for Remorins in regulating signaling in plant-microbe
interactions (Jarsch and Ott, 2011).
Elucidating the mechanisms driving StREM1.3 association with PM microdomains therefore provides a
unique opportunity for understanding the regulation
and function of membrane lateral segregation in plants.
StREM1.3 does not contain predictable transmembrane
or membrane-associated domains. The bases for its association to PMs and selective targeting to DIMs are
unknown. Here we identified a novel membrane anchor domain required for StREM1.3 tight and direct
association with the detergent-insoluble fraction of the
PM. We combined biophysics, in silico analysis, and
directed mutagenesis to unravel the molecular bases of
StREM1.3 membrane binding and its biological significance in the control of PVX propagation.
RESULTS
StREM1.3 Is a Strongly Associated Peripheral
Membrane Protein
To identify structural domains potentially involved
in PM localization of StREM1.3, we performed in silico
domain analysis and secondary structure prediction.
Proteins of the Remorin family typically contain a
variable N-terminal region (Marín and Ott, 2012) and
a conserved C-terminal domain, termed Remorin_C
(PF03763), of approximately 120 amino acid residues
that encompasses a predicted coiled-coil domain. In
StREM1.3, the Remorin_C domain extends from amino
acid 85 to 195 with a coiled-coil domain predicted
between amino acid 116 and 152 (Fig. 1A). StREM1.3
C-terminal region is predicted to be mostly a-helical
composed of an 84-amino-acid-long coiled a-helix
ranging from position 85 to 169. This helix is predicted
to be interrupted at Gly residue 171. A short (12-amino
acid) a-helix is predicted between residues 172 and
187. Finally, the most C-terminal residues are predicted to form a random coil, except for residues 192 to
195 where a short a-helix is predicted. The N-terminal
region of StREM1.3 contains the Prorich Remorin_N
domain (PF03766), extending from amino acid 27 to 84,
and is mostly unordered (Marín and Ott, 2012). No
recognizable transmembrane domains or any other
membrane-targeting signals could be detected, indicating that StREM1.3 is a peripheral membrane protein with an atypical membrane-binding domain.
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Perraki et al.
Figure 1. A C-terminal anchor peptide (RemCA)
from StREM1.3 Remorin is required and sufficient
for strong PM anchoring in vivo. A, Predicted
secondary structure and protein domains in
StREM1.3. Numbers indicate amino acid position. B, Stripping of peripheral PM proteins is not
sufficient to wash out StREM1.3. Purified PMs
from tobacco leaves were washed according to
Santoni et al., 2003 (Stripped) or treated with TBS
(Mock). On average, 50% (n = 3) of the PMassociated proteins such as Actin were stripped,
whereas the quantity of Remorin proteins associated to the PM was unaffected, as for the PIP2
aquaporin intrinsic PM protein. C, Subcellular
localization of YFP-tagged StREM1.3, truncated
StREM1.3 comprising amino acids 1 to 170,
and truncated StREM1.3 comprising amino acids
171 to 198 (RemCA peptide only) transiently
expressed in tobacco epidermal cells. Scale bars
indicate 20 mm. D, Cell fractionation of the
constructs shown in C. Crude protein extracts (CE)
from tobacco leaf cells expressing each construct
were fractionated into soluble (sol.) and microsomal (m) compartments by centrifugation (15 mg
of proteins per lane). In spite of various attempts
to solve this problem, we were not able to prevent
partial cleavage of the YFP:RemCA construct,
resulting in a free YFP signal as shown. [See
online article for color version of this figure.]
To test the nature of Remorin association with biological membranes, we purified native microsomes and
PMs from tobacco (Nicotiana tabacum) leaves and tested
its affinity by removal of extrinsic membrane proteins.
In accordance with published data, PM stripping resulted in an approximately 50% loss of total protein,
representing peripheral proteins with weak affinity to
the PM (Santoni et al., 2003). Typical of peripheral
membrane proteins is Actin, which was almost completely removed from the PM after treatment (Fig. 1B).
However, endogenous StREM1.3 remained almost
entirely associated with the PM, similar to true integral
proteins such as the aquaporin Plasma Membrane
Intrinsic Protein2 (PIP2) that was used as an integral
membrane protein control. Similar results were obtained when purified PMs were washed under high
ionic strength conditions, increasing EDTA concentrations, and increasing pH for 30 min (Supplemental
Fig. S1A). These results suggest that the association of
StREM1.3 with the membrane is not limited to electrostatic interactions.
Remorin C-Terminal Anchor Is Required and Sufficient
for PM Targeting in Vivo
A series of full-length and truncated protein variants
was generated and fused to GFP/YFP (for yellow fluorescent protein) to analyze the structural requirements
for StREM1.3 PM localization. The full-length protein
localized to the PM (Fig. 1C). This localization is consistent with immunolocalization studies reported earlier (Raffaele et al., 2009; Lefebvre et al., 2010)
suggesting that the fluorescent tag did not affect localization of the fusion proteins. Deletion of the
C-terminal residues (StREM1.31–170) resulted in an entire loss of PM association indicating the presence of a
PM anchor motif within these residues (Fig. 1C). We
will refer to this 28-amino acid region as the “Remorin
C-terminal Anchor” (RemCA). This hypothesis was
confirmed by reciprocal experiments where expression
of soluble YFP tagged with RemCA led to clear YFP
association with the PM (Fig. 1C). We designed a variety of different constructs to express the fluorescently
tagged RemCA, but we were not able to prevent a
partial cleavage of the tag, resulting in a residual
nucleocytoplasmic fluorescence in vivo. To confirm
that RemCA is sufficient to anchor soluble proteins to
the PM, we performed biochemical fractionation experiments on cells expressing StREM1.3 GFP/YFP fusion constructs (Fig. 1D). These experiments showed
that the GFP:StREM1.3 is fully microsomal whereas
YFP:StREM1.31–170 is entirely soluble. The YFP:RemCA
fusion was recovered exclusively in the microsomal
fraction, with only cleaved YFP in the soluble fraction.
These results demonstrate that RemCA is required for
StREM1.3 PM localization and sufficient to fully associate soluble proteins with the PM.
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Membrane Anchor of StREM1.3 Remorin
Nonpolar Environments Trigger Conformational
Changes in RemCA
To analyze the structural properties of RemCA,
we obtained chemically synthesized tag-free ultrapure
peptides that were assessed using circular dichroism
(CD) spectroscopy. CD spectra were acquired in 100%
2,2,2-trifluoroethanol (TFE), a solvent mimicking the
hydrophobicity of biological membranes (causing partial desolvation of the peptide, a situation that occurs
in membranes and during protein-protein and proteinlipid interactions), or in semihydrophobic solvent (50%
TFE, 50% Tris-HCl). CD spectra showed distinct minima at 208 and 222 nm, with a marked maximum at 195
nm, consistent with RemCA forming an a-helical structure (Fig. 2A). By contrast, the spectrum of RemCA
in aqueous solution (100% Tris-HCl) showed a reduced
minimum at 208 nm and reduced concavity around 195
nm, indicating that a-helical contributions to the overall
RemCA structure are reduced under these conditions.
We estimated RemCA protein secondary structure composition from CD spectra (Fig. 2B) and found that it undergoes transition from 15% a-helical content in aqueous
solution to 50% in hydrophobic nonpolar solvents. Such
estimations remain challenging and should thus be taken
with caution. Nevertheless, these data imply that the
RemCA peptide is largely unordered in aqueous solution and folds into a mostly a-helical structure in hydrophobic environments.
To gain further insights into the structural mechanisms
underlying RemCA binding to PMs, we performed homology three-dimensional (3-D) modeling of this peptide
using the I-TASSER server. The predicted structure for
RemCA corresponds to two a-helices arranged as a tight
hairpin (Fig. 2C; Supplemental File S1). This structure is
reminiscent of a recently published structure for influenza
hemagglutinin fusion peptide (HAfp; Lorieau et al., 2010;
Supplemental Fig. S2). The predicted structure has a high
helical content, therefore probably corresponding to the
conformation adopted by RemCA in hydrophobic environments. This hypothesis is consistent with the structure
of the HAfp, which forms a tight hairpin of a-helices in
the presence of lipids (Lorieau et al., 2010). The distribution of the relative hydrophobicity of amino acids reveal
an accumulation of hydrophobic amino acids at the
center of the hairpin, with the side chains of hydrophiliccharged amino acids pointing outward (Fig. 2C). The
turn connecting the two a-helices is composed of the
palindromic neutral sequence Ala-Thr-Gly-Thr-Ala.
Amphipathy of RemCA a-Helices Is Required
for Remorin Anchoring to the PM in Vivo
The predicted hydrophobic core at the center of the
RemCA hairpin and solvent-exposed charged surfaces
(Fig. 2C) implies an asymmetry in the distribution of
hydrophobic and charged residues around the a-helices,
a property known as amphipathy. Amphipathic helices
are crucial for transient anchoring of some peripheral
membrane proteins, notably viral fusion proteins (e.g.
Figure 2. Nonpolar environments trigger conformational changes in
the RemCA peptide. A, CD spectra of synthetic ultrapure RemCA
peptide in various environments. The spectral signatures of a-helices
(at 195, 208, and 222 nm) are indicated by arrows. Spectra were
recorded in aqueous (100% Tris-HCl), semihydrophobic (50% TFE)
and hydrophobic (100% TFE) environments. B, Estimated distribution
of secondary folds in RemCA peptide in the different environments
assayed. C, Sequence and predicted structure of the RemCA peptide.
Homology modeling predicts a tight hairpin of a-helices for RemCA,
consistent with the CD spectra collected in hydrophobic environments. The hydrophobicity of residues (Kyte-Doolittle scale) is indicated by a color code.
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Perraki et al.
Lins and Brasseur, 2008). We used the heliquest server
to calculate the difference, named hydrophobic moment (mH), in hydrophobicity between the two faces
of each a-helix forming RemCA. The E1722A186 helix
had a mH of 0.153, similar to values obtained for viral
fusion peptides (e.g. 0.171 for the Simian influenza
virus [SIV] fusion peptide), whereas helical fragment
P1912F198 had a mH of 0.661, with each helix organized with clearly distinct charged and hydrophobic
faces (Fig. 3A).
To test the importance of a-helices amphipathy in
the function of RemCA, we designed a StREM1.3*
variant harboring a mutated RemCA (RemCA*) with
reduced amphipathy (Fig. 3B; Supplemental File S2).
The transient expression of a YFP:StREM1.3* in tobacco leaves showed that the mutations completely
abolished membrane localization, leading to a fully
cytoplasmic protein (Fig. 3C). Loss of PM association
was confirmed biochemically by protein fractionation
where the YFP:StREM1.3* protein was exclusively detected in the soluble fraction, whereas endogenous Remorin remained in the PM fraction (Fig. 3D).
RemCA Directly Binds to Lipid Bilayers in Vitro
To test whether RemCA anchors StREM1.3 to the
PM independently of posttranslational modifications
or association with other proteins, recombinant 6His:
StREM1.3 (+), 6His:StREM1.31–170 (–), where “+” indicates wild-type StREM1.3 with native RemCA, and “–”
indicates deletion of RemCA, and 6His:StREM1.3* were
purified from Escherichia coli and tested for binding to
liposomes mimicking plant PM lipid composition in
vitro. Liposomes were separated at the top of a Suc
gradient by centrifugation. The presence of StREM1.3 in
the liposome-bound or unbound fraction was assessed
Figure 3. Amphipathy of RemCA a-helices plays a critical role in anchoring StREM1.3 Remorin to the PM. A, A helical wheel
projection of RemCA as a hairpin of a-helices showing amphipathy of each helix. Overall helix hydrophobicity (H) and mH are
indicated in the middle of the helices, with an arrow showing the direction and intensity of the mHs. B, Ribbon and surface
models of the RemCA and mutated RemCA* peptides with hydrophobicity (Kyte-Doolittle scale) shown by a color code. The
surface model of RemCA shows the predicted hydrophobic core in red and charged surfaces in blue. Substitutions introduced in
the RemCA* mutant aimed at reducing the amphipathy of the a-helices by altering the distribution of hydrophobic residues
(Y184V, A190S, and L195S) and positively charged residues (A186S, G188V, and G196V). C, Maximum projection of confocal
scans through tobacco epidermal cells expressing YFP:StREM1.3* mutant, with plastids shown in red. Scale bar indicates 20
mm. D, Cell fractionation analysis of cells expressing GFP-tagged StREM1.3 with native (+) and mutated (*) RemCA sequence,
confirming that mutations introduced in StREM1.3* abolish PM localization.
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Membrane Anchor of StREM1.3 Remorin
by western blot, and lipids were analyzed by thin-layer
chromatography (Fig. 4A; Supplemental Fig. S3).
StREM1.3 bound to liposomes in the absence of any
other proteins (Fig. 4A). Although similar amounts of
liposomes were isolated for each sample (Supplemental
Fig. S3B), binding was completely abolished in
StREM1.31–170 and StREM1.3* variants (Figure 4A).
Furthermore, and in agreement with our data obtained
using purified PMs, treatment with denaturing agents
did not significantly reduce the amount of 6His:
StREM1.3 bound to liposomes (Supplemental Fig. S1B).
These results demonstrate that RemCA confers direct
membrane binding to StREM1.3. Loss of membrane
binding in StREM1.3* indicates that amphipathy plays a
crucial role in this process.
Insertion of a peptide into membranes results in a
partial destabilization of artificial bilayers and can thus
be taken as a measure to discriminate between physical insertion and outer association (Longo et al., 1997).
To test whether RemCA induces membrane destabilization, we performed lipid-mixing assays in which
application of purified peptides to a mixture of liposomes, labeled or not by the lipophilic fluorescent
dye octadecyl rhodamine chloride (R18), leads to a
dequenching of fluorescence. We confirmed the functionality of this assay by the application of the well-known
SIV fusion peptide (Lorin et al., 2008) to liposomes (see
“Materials and Methods”). By contrast, liposome
treatment with TFE, used as a solvent for the peptides,
did not result in lipid mixing. Application of synthetic
ultrapure RemCA peptides resulted in an approximately 40% dequenching of fluorescence relative to the
values obtained with the SIV fusion peptide (Fig. 4B),
indicating destabilization of the membrane and insertion of the peptide in the bilayer.
To verify that the increase in fluorescence was due
to liposome destabilization rather than liposome aggregation, we performed liposome leakage assays,
in which liposomes entrapping the fluorescent dye
1-hydroxypyrene-3,6,8-trisulfonate (HPTS) and the corresponding cationic quencher a,a9-dipyridinium p-xylene
dibromide (DPX) were used. Application of fusion
peptides leads to a dequenching of fluorescence due to
physical separation of both components (Ellens et al.,
1985). We used liposomes composed of either negatively charged lipids or neutral lipids. As expected, the
use of the detergent Triton X-100 (TX100) entirely
disrupted the liposomes, leading to maximal fluorescence (Fig. 4C). Application of the SIV fusion peptide
induced approximately 90% leakage of negatively
charged liposomes, and only approximately 9% leakage of neutral liposomes, well above leakage induced
by solvents used for the peptides (leakage , 2%
by either dimethyl sulfoxide or TFE for negatively
charged and neutral liposomes, respectively). Application of synthetic RemCA peptide yielded approximately 5% of leakage of negatively charged liposomes,
but did not cause any significant leakage of neutral
liposomes (Fig. 4C). These experiments demonstrate
that RemCA physically inserts into membranes independently of other proteins, and predominantly binds
negatively charged lipids, suggesting a differential affinity for classes of lipids.
RemCA Confers Lipid-Specific Binding to StREM1.3
Figure 4. RemCA confers intrinsic lipid binding to StREM1.3 Remorin.
A, Affinity-purified 6His:StREM1.3 (WT) binds to protein-free liposomes in vitro in a RemCA-dependent manner. Liposome binding is
abolished in 6His:StREM1.3 recombinant protein lacking RemCA
(ΔCA) or harboring a mutated RemCA (*). The soluble (Sol.) and
liposome-bound (Bound) fractions recovered after centrifugation are
shown. Lipid mixing assays (B) and liposome leakage assays (C) show
that RemCA ultrapure peptides insert into liposome membranes containing negatively charged lipids, as indicated by an increase in fluorescence dequenching. By contrast, liposome leakage assays (C)
showed that RemCA is not able to insert into neutral liposome membranes. Solvent used in (C) was dimethyl sulfoxide for negatively
charged liposomes and TFE for neutral liposomes. Error bars indicate
SD of six to eight independent measurements.
To test whether RemCA binds selectively to certain
lipids, we probed immobilized lipids with purified
recombinant 6His:StREM1.3 and 6His:StREM1–170 in
lipid overlay assays. StREM1.3 showed differential
affinity to five of the seven lipids tested (Fig. 5A).
Affinity of StREM1.3 was strong to PI(3,4)P2 [for phosphatidylinositol (3,4)-bisphosphate] and sphingosine-1P, moderate to phosphatidic acid and PI(3,5)P2, and
weak to phosphatidyl-Ser. Binding to these immobilized lipids was dependent on RemCA because it
was almost completely abolished in the StREM1.31–170
protein, with only a weak residual binding to PI(3,4)P2
remaining. Similar results were obtained with the purified recombinant 6His:StREM1.3* mutant (Supplemental
Fig. S4A). These results indicate that RemCA has affinity
for minor anionic phospholipids and negatively charged
lipids, and that RemCA confers selectivity to the range
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Finally, to test the specificity of RemCA-mediated
binding in vivo, we extracted DIMs from PM purified
from tobacco plants expressing RemCA tagged with
the soluble YFP. As expected, approximately 80% of
total proteins were excluded from the DIM fraction
(Fig. 5C; Supplemental Fig. S4C; see also Raffaele et al.,
2009) but the endogenous tobacco Remorin proteins
were almost exclusively found in DIM fractions (Fig. 5C).
Strikingly, the majority of YFP:RemCA proteins were
also detected in DIMs (Fig. 5C). These results demonstrate that RemCA is sufficient to target soluble proteins specifically to DIMs of plant PM.
StREM1.3 Forms Homotrimers in Vitro Independently
of RemCA
Figure 5. RemCA targets soluble proteins to the PM in a lipid-specific
manner. A, Lipid overlay assays performed with 6His-tagged StREM1.3
and StREM1.31–170 show a differential affinity of StREM1.3 for various
classes of lipids, dependent on RemCA. On the right, western blot of
the protein input material illustrating equal loading for StREM1.3 and
StREM1.31–170 is shown. PA, Phosphatidic acid; PIP3, PI(3,4,5)P3; PIP2,
phosphatidylinositol bisphosphate, the position of phosphate groups is
indicated below; S1P, sphingosine 1-P. B, 6His:StREM1.3 shows higher
affinity for liposomes with a composition mimicking DIM compared
with PM (described in “Materials and Methods”). Histograms show
quantifications and SE based on western-blot signals for three independent experiments. Differential affinity is significant with P , 0.05
in a Student’s t test. C, RemCA-tagged GFP transiently expressed in
tobacco leaves segregates into the DIM fractions of the PM, similar to
the endogenous Remorin. Total proteins were quantified by bicinchoninic acid assay relative to total proteins in purified PM (error bars
indicated SD from three independent replicates). m, Microsomal fraction; DSM, detergent-soluble membranes.
of lipids bound in vitro. Interestingly, these lipids are
enriched in membrane rafts from plants (Vermeer et al.,
2009; Furt et al., 2010) or animal cells (Hope and Pike,
1996; Pike et al., 2002), suggesting that RemCA lipidspecificity contributes to driving the segregation of
StREM1.3 into membrane microdomains.
To support this hypothesis, we first tested the
binding of 6His:StREM1.3 to liposomes with lipid
composition mimicking either the PM or DIMs in vitro
(see “Materials and Methods”). Although the total
amount of liposomes pulled down was similar for all
samples (Supplemental Fig. S4B), the relative quantification of 6His:StREM1.3 bound demonstrated a higher
affinity for DIM-like liposomes compared with PM-like
liposomes (Fig. 5B).
Previous work showed by direct electron microscopy
observations and chemical crosslinking with glutaraldehyde that group 1 Remorins forms homopolymeric filaments (Bariola et al., 2004). To test whether RemCA
plays a role in the assembly or stability of these higherorder polymers, we used exclusion column chromatography to estimate the native Mr of 6His:StREM1.3 and
6His:StREM1.3* recombinant proteins. Chromatography
revealed a peak of elution at 1.25 mL (Supplemental Fig.
S5A). Western-blot analysis after denaturing SDS-PAGE
of the corresponding fractions confirmed that 6His:
StREM1.3 eluted at this volume (Supplemental Fig. S5B).
Calibration of the gel filtration column with standards of
known Mr allowed to estimate 6His:StREM1.3 native Mr
to 120 kD, suggesting that 6His:StREM1.3 forms metastable trimers in vitro (Fig. 6A). Neither monomers nor
dimers were detected by gel filtration. Chromatography
performed with 6His:StREM1.3* protein showed similar
elution pattern, indicating that a nonfunctional RemCA
does not alter the assembly of StREM1.3 into trimers.
Interestingly, as discussed previously (Supplemental Fig.
S2), RemCA shows structural similarities with influenza
virus hemagglutinin (HA) fusion peptide. Influenza HA
adopts a central trimeric a-helical coiled-coil structure,
and the formation of trimers of fusion peptide hairpins is
critical for membrane fusion (Cross et al., 2001). To get
insights on the possible consequences of StREM1.3 trimer
formation on its membrane-binding properties, we used
homology modeling and geometry-based docking to
produce a 3-D model of an StREM1.3 C-terminal domain
(E87-F198) trimer (Fig. 6B). This model suggests that
RemCAs are organized circularly, forming a hydrophobic core at the center of the RemCA trimer of hairpins
and exposing patches of positively charged residues all
around the C terminus of the trimer. This conformation
would create a strong mH directed inward in the RemCA
trimer, and likely increases the strength and lipid specificity of the membrane binding.
Mutations in RemCA* Abolish StREM1.3 Restriction
of PVX Cell-to-Cell Movement
To test for the biological significance of StREM1.3
PM localization, we exploited the previously reported
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Membrane Anchor of StREM1.3 Remorin
role of StREM1.3 in restricting the cell-to-cell propagation of the PVX in tomato (Solanum lycopersicum) leaves
(Raffaele et al., 2009). Here, we transiently coexpressed
in tobacco leaves 35S:StREM1.3 or 35S:StREM1.3* with
a highly diluted suspension of Agrobacterium tumefaciens harboring an infectious PVX:GFP clone, so that
only isolated individual cells were infected with the
PVX:GFP. The accumulation of PVX:GFP in distinct
infection foci was visible 5 d post infection (dpi).
StREM1.3 or StREM1.3* expression was verified by
western blot (Supplemental Fig. S6). The effect of the
expression of the StREM1.3 proteins was assessed by
visualizing the virus infection foci and counting the
number of fluorescent cells of at least 30 infection foci in
comparison with plants infiltrated with A. tumefaciens
carrying the PVX:GFP clone only. At 5 dpi, plants
infiltrated with A. tumefaciens carrying PVX:GFP alone
showed infection foci covering 17 cells in average
(Figure 7, A and B). As expected, the coexpression of
StREM1.3 lead to a significant decrease in PVX:GFP
spreading, with foci covering 12 cells on average (Fig.
7, A and B). In contrast, coexpression of StREM1.3* did
not significantly alter the size of the PVX:GFP infection
foci (Fig. 7, A and B). StREM1.3 was shown to directly
interact with TGBp1, the viral movement protein,
during PVX infection (Raffaele et al., 2009). To determine if the loss of activity in StREM1.3* was due to a
loss of interaction with TGBp1, we tested the interaction of the StREM1.3* and StREM1.31–170 mutants with
TGBp1 in a split ubiquitin assay. In both cases, a clear
interaction was observed, similar to the wild-type
StREM1.3 (Fig. 7C). These results clearly demonstrate
that RemCA-mediated PM anchoring of StREM1.3 is
required for the function of this protein in the control
of virus propagation, and indicate that this activity
takes place at the PM.
DISCUSSION
Figure 6. StREM1.3 Remorin forms homotrimers independently of
RemCA in vitro. A, Exclusion column chromatography of 6His:
StREM1.3, 6His:StREM1.3*, and elution standards allow for an estimate of the native Mr of 6His:StREM1.3 to 120 kD, suggesting that it
assembles into trimers, and shows that mutations introduced in 6His:
StREM1.3* do not alter the ability to form trimers. Standards used: 1,
Blue dextran (2 megadaltons); 2, thyroglobuline (670 kD); 3, catalase
(232 kD); 4, ovalbumine (43 kD); 5, chymotrypsinogene (25 kD); 6,
soybean (Glycine max) trypsin inhibitor (21.5 kD); 7, RNase (13.7 kD);
8, cytochrome C (12.3 kD). B, A trimer model of the C-terminal domain of StREM1.3. Views from the C-terminal end of the trimer, with
residues colored according to hydrophobicity (Kyte-Doolittle scale),
reveal the formation of a hydrophobic core (in red) at the center of the
trimer, with positive surface patches surrounding the RemCA region (in
blue).
In this paper, we showed that StREM1.3 directly
binds to the PM via a short C-terminal anchor domain,
RemCA. This domain is unordered in aqueous solution
and spontaneously folds, probably into a tight hairpin
of amphipathic a-helices, in nonpolar environments.
The amphipathy of RemCA helices is required for
StREM1.3 PM localization. RemCA selectively binds to
lipids enriched in DIMs and targets the soluble YFP to
DIMs in vivo. StREM1.3 forms trimers in vitro. This
does not require a functional RemCA and may increase
the stability of PM anchoring in vivo. PM anchoring is
crucial for StREM1.3 function in restricting PVX movement, because mutations in RemCA that abolish PM
binding also abolish StREM1.3 function toward PVX.
Molecular mechanisms controlled by raft-mediated signaling and invasion of host cells via membrane microdomains may be altered by directly targeting proteins
to these sites. Using RemCA as an anchor may thus
provide the opportunity to modulate raft-mediated signaling processes or alter host resistance toward pathogenic microbes.
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Perraki et al.
creating an environment favorable to RemCA folding
(Fig. 8, B and C). Association of group 1 Remorins into
higher-order complexes has been reported in vitro
(Bariola et al., 2004) and in vivo (Raffaele et al., 2009;
Lefebvre et al., 2010), but whether this is triggered by
external signals remains unknown. We propose that
RemCA anchoring occurs as described for other lipidbinding proteins (Stahelin et al., 2003; Peter et al.,
2004) in a two-step manner. First, positively charged
residues E172-A186 increase the affinity of RemCA
for negatively charged lipids of the inner leaflet of
membrane rafts (Fig. 8, A and D). This destabilizes
the internal monolayer of the PM and allows RemCA
to insert deeper in the membrane (Fig. 8, B and E).
The complete folding of RemCA results in the formation of a tight hairpin of a-helices defining a
hydrophobic pocket at its core. As described for influenza HAfp (Han et al., 2001), formation of this
hairpin could result in a deeper insertion of RemCA
explaining the intrinsic-like behavior of Remorins (see
Fig. 1B). Considering that influenza HAfp is nonfusogenic if not properly kinked (Lai and Tamm,
2007), the tight closure of the RemCA hairpin of helices might be required for stable anchoring in the PM.
Finally, the possibility that the formation of StREM1.3
multimers occurs at the PM, eventually under control of
external signals (Fig. 8, B and E), would allow the aggregation of membrane rafts into higher-order clusters, as suggested during the response to pathogens
(Raffaele et al., 2009).
Figure 7. RemCA-mediated PM binding is required for StREM1.3 restriction of PVX cell-to-cell propagation. A, Expression of PVX:GFP
alone and in combination with StREM1.3 or the mutated StREM1.3*
variant unable to bind to the PM. Images of infection foci were taken
at 5 dpi of tobacco leaves. Scale bars indicate 150 mm. B, Quantification of PVX cell-to-cell spreading showing the average number of
cells in the infection foci at 5 dpi. For each treatment, 90 pictures
were analyzed, with error bars indicating SD. Significance was
assessed by Dunn’s multiple comparison test (***, P , 0.001; n = 3).
C, Split ubiquitin assays showing that StREM1.31–170 and StREM1.3*
interact with PVX TGBp1 movement protein similar to the wild-type
StREM1.3. [See online article for color version of this figure.]
Molecular Bases for the Assembly of Remorin
Microdomains at the PM: a Model
Our results indicate that the folding of RemCA into
amphipathic a-helices, probably organized as a tight
hairpin, is required for StREM1.3 PM anchoring and
function. Our CD data showed that RemCA switches
from unordered to a-helical folding in nonpolar environments. RemCA folding could be triggered by proximity with the hydrophobic vicinity of the PM (Fig. 8A).
Similar observations were made for Fis1 Tail-anchor
protein (Suzuki et al., 2003), the ArfGAP1 lipid sensor
motif (Bigay et al., 2005), and for the amphipathic
membrane-binding helix of the epsin N-terminal homology family of clathrin adaptors (Ford et al., 2002).
Alternatively, the formation of StREM1.3 trimers would
result in the formation of a hydrophobic core, thereby
Biophysical Properties Defining RemCA Lipid-Specificity
In this work, we have shown that RemCA alone is
able to anchor soluble hydrophilic proteins to the PM,
such as GFP/YFP (Fig. 1). We cannot exclude that
additional structural features of StREM1.3 protein
modulate this activity, such as posttranslational modifications (e.g. phosphorylation), and the formation of
StREM1.3 multimers that would exacerbate local gradients of hydrophobicity and charge. It is remarkable
that RemCA preferentially binds to negatively charged
lipids abundant in membrane rafts, and targets the
soluble YFP to DIMs. A number of globular domains
are known to bind to the surface of the PM by
specifically recognizing certain lipids, particularly
discriminating between different phospholipid types
(Lemmon, 2008; Kutateladze, 2010). A combination of
steric recognition of specific lipid polar head groups,
electrostatic attraction to negatively charged lipids and
membrane destabilization-mediated penetration govern the function of these domains. Some of those such
as the Annexin, epsin N-terminal homology, AP180
N-terminal homology, BAR, and Phox homology domains adopt an a-helical fold. Overall, the hydrophobicity, the mH, the net charge, and helix-breaking amino
acids contribute to the membrane-binding properties
of amphipathic helices (Drin and Antonny, 2010).
In small GTPases, polybasic clusters provide PM
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Membrane Anchor of StREM1.3 Remorin
Figure 8. Model for the molecular mechanisms
leading to the assembly of StREM1.3 microdomains at the PM. Conformational changes in
RemCA, leading to folding into amphipathic
a-helices, are required to anchor StREM1.3 to the
PM. These conformational changes are triggered
by nonpolar environments, such as the vicinity of
the PM (A). Helix E172-A186 folds exposing
clusters of positively charged residues (+). These
residues establish electrostatic interactions with
the negatively charged lipids of the inner leaflet of
membrane rafts, destabilizing the PM and allowing insertion of RemCA in the bilayer (B). Hydrophobicity triggers RemCA hairpin formation,
creating a hydrophobic pocket (small red star)
that deepens and strengthens insertion in the
membrane. Alternatively, an external signal may
trigger the formation of StREM1.3 trimers (C). The
hydrophobic core of the trimer (large yellow star)
favors folding of helix E172-A186 of RemCA (D),
leading to PM anchoring as previously described.
The formation of trimers, either prior to or after
PM anchoring, is likely to increase the strength
of the lipid binding, may promote membrane
domain aggregation, lead to membrane deformation, or mediate the assembly of protein complexes
at the PM via Remorin protein-protein interaction
domains (E).
specificity by binding to negatively charged PI(4,5)P2
and PI(3,4,5)-P3 (for trisphosphate) of the PM (Heo
et al., 2006). The charge of the polar face of ArfGAP1
amphipathic helices has been associated with the
ability to perceive membrane curvature, with the introduction of Lys residues at the polar-nonpolar interface reducing curvature sensitivity and allowing
binding to large liposomes (Drin et al., 2007). It is
tempting, therefore, to speculate that RemCA membrane binding could sense membrane curvature as a
means to specifically target certain domains in the
membrane. In addition, large hydrophobic residues on
both sides of the kink are required for setting the angle
of the hairpin structure and for function in influenza
HAfp (Lai and Tamm, 2007). The L174, L180, I194, and
F198 could play this role in RemCA. The overall conformation of RemCA, as well as individual residue
side chains, is likely to contribute to its membranebinding ability, and further investigations will be required to dissect how this activity is mediated.
Toward Understanding the Molecular Bases of Specific
Association with Membrane Rafts
RemCA drives the soluble YFP to DIM PM fractions,
and lipids bound by RemCA are enriched in DIMs.
Indeed, polyphosphoinositides and sphingolipids are
enriched in membrane rafts from plants (Vermeer
et al., 2009; Furt et al., 2010) and animal cells (Hope
and Pike, 1996), the latter being also enriched in
phosphatidylserine (PS) (Pike et al., 2002). The ability
to specifically target DIMs, therefore, might be related
to the differential affinity of RemCA to certain negative
lipids, as known for SNARE proteins (Mima and
Wickner, 2009). However, It should be noted that the
presence of a protein in DIMs is not sufficient to associate it with membrane rafts (Munro, 2003; Simons
and Gerl, 2010; Tanner et al., 2011). In the case of
StREM1.3, we previously reported that enrichment in
DIMs correlates with clustering into microdomains of
70 nm in the PM detected by immunogold labeling
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Perraki et al.
(Raffaele et al., 2009). More important, both DIM enrichment and clustering at the PM were phytosterol
dependent (Kierszniowska et al., 2009; Raffaele et al.,
2009; Tanner et al., 2011). Because lipid-lipid interactions
are sufficient to drive the segregation of liquid-ordered
and liquid-disordered phases in model membranes
(Dietrich et al., 2001; Baumgart et al., 2003), it can be
hypothesized that RemCA-selective lipid-binding properties could be sufficient to restrict the localization of
StREM1.3 to membrane rafts.
Nevertheless, it is reasonable to assume that in addition to RemCA, other domains or motifs may determine
the organization of StREM1.3 into PM microdomains.
Indeed, the association of Remorins with DIMs is regulated quantitatively by signals such as cold (Minami
et al., 2009) and pathogen-associated molecular patterns
(Keinath et al., 2010). The molecular bases of this regulation are not fully understood but may likely involve
protein-protein interactions or posttranslational modifications. It is also not known whether the high-ordered
organization of Remorins into multimers is regulated by
external signals. In any case, protein-protein interactions
mediated by the coiled-coil or the N-terminal domain of
Remorins (Lefebvre et al., 2010; Tóth et al., 2012) likely
modulate the localization of Remorins in vivo.
2010) sheds new light on the involvement of host
membrane proteins in virus movement. Arabidopsis
synaptotagmin1 resides in the cortical endoplasmic
reticulum, endosomes, DIMs at the PM, and plasmodesmata (PDs; Schapire et al., 2008; Minami et al.,
2009; Lewis and Lazarowitz, 2010; Yamazaki et al.,
2010), and regulates endocytosis and PM repair processes (Schapire et al., 2008; Yamazaki et al., 2008;
Lewis and Lazarowitz, 2010). These studies support a
link between DIMs and PDs, and provide evidence for
a role of DIM proteins in cell-to-cell communication
via PDs.
Considering the diversity of proteins in the Remorin
family (Raffaele et al., 2007), there is probably a corresponding diversity of subcellular localizations and
functions to be discovered. Analyzing the variability of
the RemCA region in relation to Remorins subcellular
locations, the variability of the membrane lipid composition according to the nature of membrane compartments and plant species, and specific functions
devoted to each protein in the Remorin family will
provide valuable insights in the mechanisms regulating lateral heterogeneity of plant PMs.
MATERIALS AND METHODS
Implications for the Mechanisms Underlying PVX
Cell-to-Cell Movement
In a previous study, we reported that StREM1.3 restricts PVX cell-to-cell movement independently of
viral replication and that StREM1.3 physically associates in vivo and in vitro with the TGBp1 of the virus
(Raffaele et al., 2009). Here we show that restriction of
PVX movement is abolished in StREM1.3* cytoplasmic
mutant. We also show that deletion or mutation of
RemCA does not alter StREM1.3 interaction with
PVX TGBp1 in a split ubiquitin assay, which is consistent with evidence that RemCA would insert into
the PM. This indicates that RemCA is not directly involved in the interaction with TGBp1 but rather that
StREM1.3-TGBp1 interaction takes place at the PM.
The subcellular localization of TGBp1 is complex and
depends on the other TGB proteins, viral RNA, and the
host plant species (Verchot-Lubicz et al., 2010). When
expressed alone in tobacco, TGBp1 is mostly nuclear
and cytoplasmic. However in the presence of viral
RNA, TGBp1 could traffic along the endoplasmic reticulum with TGBp2 and TGBp3 granules, and reach
plasmodesmata. Given that StREM1.3 is anchored on
the cytosolic face on the PM, it could associate with
cytoplasmic TGBp1 at the periphery of the cell, explaining why we could detect association when TGBp1
was expressed alone. Future work will investigate the
specificity of StREM1.3 association with TGBp1 and
the impact of viral RNA on this interaction. The recent
report of the role of Arabidopsis (Arabidopsis thaliana)
synaptotagmin1 in the control of Begomovirus and Tobamovirus cell-to-cell movement (Lewis and Lazarowitz,
Molecular Cloning and Sequence Analyses
StREM1.3 was cloned from potato (Solanum tuberosum ‘Désirée’) complementary DNAs by standard Gateway techniques using the pDONR221 entry
vector and primers containing the recombination site sequences and the
gene-specific sequences 59-ATGGCAGAATTGGAAGCT-39 (forward) and
59-TCAAAATATTCCAAGGAT-39 (reverse for N-terminal fusion), 59-AAATATTCCAAGGAT-39 (reverse for C-terminal fusion), or 59-TCAACGTTTAGCTTCAATCAT-39 (reverse for YFP:StREM1–170). A forward primer
containing the gene-specific sequence 59-GGAGAAGATCTTCTC-39 was used to
clone the RemCA constructs. StREM1.3* was generated by PCR amplification
using a reverse primer containing the sequence 59-TCAAAATATTACAGAGATTTTCTTTGGAGAAGTTACAGTGGAACGGACTTTTGC-39. Mutated residues were selected at the C terminus of RemCA to facilitate PCR-based
mutations. The position of Glys and the aromatic residue were shown to be
important in HA fusion peptide function (Cross et al., 2001); therefore, G188V
and G196V were introduced to preserve small size while introducing hydrophobic amino acids and Y184V to remove the aromatic residue. A186S and
A190S were introduced to preserve the tiny size of amino acids while removing hydrophobic residues, and L195S allowed further reduction of the
local hydrophobicity of RemCA peptide. These mutations allowed reduction
of the mH from 0.182 to 0.112 for the whole RemCA peptide. These entry
clones were recombined into the pDEST17 (6His fusions), pH7YWG2,
pH7WGY2, or pk7WGF2 (YFP/GFP fusions) vectors. Secondary structure
predictions were ran on SSPro (http://download.igb.uci.edu/sspro4.html)
and JPred 3 (http://www.compbio.dundee.ac.uk/www-jpred/) servers. Coiledcoil domains were predicted using the Marcoil1.0 server (http://bioinf.wehi.edu.
au/folders/mauro/Marcoil/index.html). Amphipathic helix analyses were performed with the heliquest server (http://heliquest.ipmc.cnrs.fr/) and 3-D modeling via the I-TASSER server (http://zhanglab.ccmb.med.umich.edu/I-TASSER/;
Roy et al., 2010). StREM1.3 C-terminal domain models were assembled into a
trimer using the SymmDock server (Schneidman-Duhovny et al., 2005). Models
were rendered using University of California, San Francisco Chimera (Pettersen
et al., 2004).
Plant Transformation and Fluorescence Microscopy
Four-week-old tobacco plants (Nicotiana tabacum ‘Xanthi’) were used
for Agrobacterium tumefaciens (strain GV3101)-mediated transient expression
(Batoko et al., 2000). Confocal imaging was performed 2 d after agroinfiltration
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Membrane Anchor of StREM1.3 Remorin
using Leica TCS SP2/5 confocal microscopes with 203 to 633 oil/water immersion objectives. Laser and image settings were as described by (Raffaele
et al., 2009).
Preparation of Pure PM and DIM
Microsomal fractions and PM from tobacco leaves were purified described
in (Mongrand et al., 2004). For DIM preparation, TX100 (10% [w/v]) was
added to a detergent-to-PM proteins ratio of 10 (at 1% [w/v] final concentration), and the PM preparations were solubilized at 4°C for 30 min. Treated
PMs were brought to a final concentration of 48% Suc (w/w), overlaid successively with layers of 40%, 35%, and 30% Suc in Tris-buffered saline (TBS)
buffer (w/w), and then centrifuged for 16 h at 200,000g in a TST41 rotor
(SORVALL). DIMs could be recovered above the 30% to 35% layers as an
opaque band, and this fraction was washed in 4 mL of TBS buffer to remove
residual Suc. The protein concentration was determined with a BCA protein
assay to avoid TX100 interference, using bovine serum albumin as a protein
standard.
Protein Analysis, Western Blots, and Protein Purification
6His-tagged constructs were expressed in Escherichia coli BL21 DE3 cells.
Cells were lysed by ultrasonication, and the crude extract was centrifuged
at 10,000g for 15 min. 6His-tagged proteins were mostly recovered in the
soluble fraction. Soluble proteins were then purified using fast flow chelating
sepharose resin (Amersham) according to manufacturer’s instructions. Ultrapure RemCA peptides were obtained by de novo peptide synthesis (GL
Biochem Shanghai Ltd; www.glschina.com). Western-blot analyses were performed using either anti-REM (Raffaele et al., 2009), anti-Plasma Membrane
proton pump ATPase (Lefebvre et al., 2004), anti-PIP (Santoni et al., 2003), antiactin, or commercial anti-GFP (Millipore/Roche) antibodies.
in the wavelength range from 190 to 250 nm. Peptide secondary structures
were determined using CDpro software package, which involved CDSSTR,
SELCON3, and CONTINLL methods (Adam et al., 2004; Zakharov et al.,
2004). Percentages were calculated by averaging the percentages provided by
the three methods. The peptides stock solutions used for the measurements
were diluted in 1 mM Tris buffer, pH 7.4, or in TFE.
Liposome Leakage Assays
Membrane perturbation and vesicle release can be measured by the assay of
(Ellens et al., 1985) based on the quenching of HPTS by DPX. HPTS and DPX
were coencapsulated in the aqueous phase of the same liposomes. When
leakage of vesicle content occurs, the quenching by DPX stops, and subsequently the fluorescence of HPTS increases. Small unilamellar vesicles were
used in our experiments. These vesicles were prepared from a solution of
multilamellar vesicles obtained after hydration for 1 h at 37°C of dry lipid
films. These films were mixtures by weight of 26.6% PC, 26.6% sphingomyelin
(SM), 26.6% PE, and 20.2% cholesterol (CHOL) for neutral liposomes; and 30%
PC, 30% PE, 2.5% PI, 10% PS, 5% SM, and 22.5% CHOL for negatively charged
liposomes (% expressed as w/v). Large unilamellar vesicles were prepared by
the extrusion technique of Mayer et al., 1986. The multilamellar vesicle suspension was submitted to five successive cycles of freezing and thawing and
thereafter extruded 10 times through stacked polycarbonate filters (pore size:
0.08 mm), under a nitrogen pressure of 20 bars using an extruder (Lipex Biomembranes, Vancouver, Canada). The concentration of the liposome suspensions was determined by phosphorus analysis. The peptides, dissolved
in TFE or hexafluoropropene/TFE, were added to a mixture of liposomes
with encapsulated HPTS and DPX. HPTS fluorescence was measured on a
PerkinElmer LS-50B fluorometer (excitation: 360 nm; emission: 520 nm). Liposomes were prepared as described above but were rehydrated with 1 mL of
12.5 mM HPTS (45 mM NaCl), 45 mM DPX (20 mM NaCl), and 10 mM Tris-HCl,
pH 7.4, and passed through a Sephadex G-75 column to remove unencapsulated
material.
Extrinsic PM Proteins Stripping
Stripping on PM preparations and microsomal fractions was performed as
described in (Santoni et al., 2003) with minor modifications. Membranes (200
mg) were incubated in 40 mL/1 mL of 5 mM EDTA, 5 mM EGTA, 4 M urea, and
5 mM Tris-HCl, pH 9.5, for 5 min on ice before being centrifuged for 20 min at
100,000g. The subsequent pellet was suspended in 20 mM NaOH and centrifuged at 100,000g for 20 min. The membranes were then washed in 2 mM
EDTA, 2 mM EGTA, 100 mM NaCl, and 5 mM Tris-HCl, pH 8; centrifuged at
100,000g for 20 min; and finally resuspended in loading buffer for SDS-PAGE
and western blot where samples were loaded at equal volumes. The mean
protein-purification yield was approximately 50%. A negative control treated
with TBS was added (mock), yielding 95% recovery. Total protein amounts
were quantified by Bradford or by image quantification of the Coomassiestained gels.
Liposome and Lipid Binding Assays
Liposomes mimicking the lipid content of tobacco PM were freshly prepared in
TBS buffer according to Doms et al., 1985. For PM-like liposomes, the lipid content
was a phosphatidylethanolamine (PE)/ phosphatidylcholine (PC)/ phosphatidic
acid/glucosylcerebrosides/tobacco mix of free sterols/acylsterylglucosides/
sterylglucosides at molar ratio 1/0.6/0.2/0.6/0.8/0.5/0.5 and for DIM-like liposomes, the same lipids at molar ratio 1/0.6/0.2/1.2/2.5/3/1 (Mongrand et al.,
2004; Furt et al., 2010). Signals were quantified by ImageJ software. Standard plant
lipids were purchased from SIGMA and MATREYA. Five micrograms of purified
6His:StREM1.3, 6His:StREM1.31–170, or 6His:StREM1.3* was incubated for 30 min
at 30°C with 5 mM liposomes. The samples were further processed in 62% Suc and
placed at the bottom of a discontinuous Suc gradient (35/30/15/5 [w/w]). After
centrifugation, liposomes floating at the 15% to 5% interface (top of the gradient)
and fractions at the bottom of the gradient were collected, proteins were precipitated with 15% TCA and analyzed by western blot. To control for liposome
recovery, lipids were extracted and analyzed by thin-layer chromatography as
described previously (Mongrand et al., 2004).
CD Measurements
CD spectra were recorded on a Jasco J-815 CD spectrometer with 10-mmpath-length quartz cuvettes. Ten scans were taken and automatically averaged
Lipid Mixing Assays
The induction of vesicular lipid mixing by the different peptides was tested
with liposomes (small unilamellar vesicles) made of PC/PE/SM/CHOL at a
molar ratio 26.6/26.6/26.6/20.2 and PC/PE/PI/PS/SM/CHOL at a molar
ratio 30/30/2.1/10/5/22.5. Labeled liposomes were obtained by incorporating R18 in the dry lipid film at a concentration of 6.3% of the total lipid weight.
Labeled and unlabeled liposomes were mixed at a weight ratio of 1:4 and a
final concentration of 50 mM in 10 mM Tris-HCl, 150 mM NaCl, 0.01% EDTA,
and 1 mM NaN3 (pH 8). Mixing of liposome membranes was followed by
measuring the fluorescence increase of R18, a lipid soluble probe, occurring
after the fusion of labeled and unlabeled liposomes, as described in (Lins et al.,
2002). Incubation of labeled and unlabeled vesicles in buffer alone did not
modify the fluorescence intensity. Fluorescence was recorded at room temperature (excitation: 560 nm, emission: 590 nm) on an LS-50B PerkinElmer
fluorometer.
Size-Exclusion Chromatography
Size-exclusion chromatography was performed on an AKTApurifier
apparatus (GE Healthcare). 6His:StREM1.3 and 6His:StREM1.3* purified from
E. coli cultures (1.2 g/L) were centrifuged for 5 min at 20,000g before being
processed. The molecular size of the proteins was analyzed by chromatography on a Superdex 75 10/30 column (GE Healthcare) equilibrated with 20 mM
HEPES, pH 7.4, 300 mM NaCl. Proteins (200 mL) were eluted with a flow rate
of 0.2 mL/min and recorded by continuously monitoring the absorption at
280 nm. The column was calibrated with the standard proteins described in
Figure 6.
Viral Spreading and Split Ubiquitin Assays
To assess spreading of PVX:GFP in tobacco leaves, A. tumefaciens strain
GV3101 carrying the respective StREM1.3 constructs were infiltrated at a final
optical density at 600 nm = 0.2 together with the same strain carrying the
plasmid pGr208, which expresses the PVX:GFP complementary DNA, as
well as the helper plasmid pSoup (Peart et al., 2002) at final optical density at
600 nm = 0.001. Spreading of PVX:GFP was visualized by confocal laser-
Plant Physiol. Vol. 160, 2012
635
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Copyright © 2012 American Society of Plant Biologists. All rights reserved.
Perraki et al.
scanning microscopy at 5 dpi, and the number of fluorescent cells of at least
30 PVX:GFP infection foci were counted. Protein extracts from the infiltrated tobacco leaves were subjected to cell fractionation and western blot.
The experiment was repeated three times with the same results. Split
ubiquitin assays were performed as described in (Raffaele et al., 2009) with
yeasts (Saccharomyces cerevisiae) grown on synthetic dextrose medium supplement with uracil (50 mg L21), Leu (380 mg L21), Lys (76 mg L21), and
59fluoro-orotic acid (1 g L21).
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure S1. Membrane washing experiments show that
StREM1.3 strongly binds to purified PM and liposomes.
Supplemental Figure S2. Sequence and structure comparison between
StREM1.3 RemCA and influenza HAfp.
Supplemental Figure S3. Controls for the production of purified recombinant proteins and liposome-binding assays.
Supplemental Figure S4. Controls for lipid-specific binding of StREM1.3.
Supplemental Figure S5. StREM1.3 Remorin forms trimers in vitro.
Supplemental Figure S6. Western-blot controls for the expression of
StREM1.3 and StREM1.3* in PVX:GFP propagation experiments.
Supplemental File S1. Protein Databank file containing atom coordinates
for the predicted StREM1.3 structure.
Supplemental File S2. Protein Databank file containing atom coordinates
for the predicted StREM1.3* structure.
ACKNOWLEDGMENTS
We would like to thank Mohammed-Amine Belka (Laboratoire de Biogenese
Membranaire [LBM]), Christelle Flore (Centre de Biophysique Moléculaire
Numérique, Université de Liège), and Jeanny Laroche-Traineau (LBM) for their
valuable contributions. We are very grateful to Claudia Popp and Thomas Ott
(Ludwig-Maximilians-University) for inspiring discussions, shared data, contagious enthusiasm, and a lot of help. We thank all other members of the labs of
Sébastien Mongrand (LBM) and Thomas Ott (Ludwig-Maximilians-University)
for fruitful discussions. We acknowledge the platform Métabolome-FluxomeLipidome of Bordeaux (http://www.biomemb.cnrs.fr/INDEX.html https://
www.bordeaux.inra.fr/umr619/RMN_index.htm) for contribution to lipid analysis and the unit of plant imaging of Bordeaux Imaging Center for confocal
imaging. A.P., L.L., S.M., and S.R. designed research. A.P., J.-L.C., J.-M.C.,
S.M., and S.R. performed research. J.-M.C., L.L., and S.G-R. contributed new reagents and analytic tools. A.P., J.-M.C, S.M., and S.R. analyzed data. S.M. and S.R.
wrote the paper. All authors read, edited, and approved the manuscript.
Received May 16, 2012; accepted July 31, 2012; published August 1, 2012.
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