Download Phosphatidylinositol 4-Phosphate Formation at ER Exit Sites

Developmental Cell 11, 671–682, November, 2006 ª2006 Elsevier Inc.
DOI 10.1016/j.devcel.2006.09.001
Phosphatidylinositol 4-Phosphate Formation
at ER Exit Sites Regulates ER Export
Anna Blumental-Perry,1 Charles J. Haney,1
Kelly M. Weixel,1,2 Simon C. Watkins,1
Ora A. Weisz,1,2 and Meir Aridor1,*
1
Departments of Cell Biology and Physiology
2
Renal-Electrolyte Division
University of Pittsburgh School of Medicine
3500 Terrace Street
Pittsburgh, Pennsylvania 15261
The mechanisms that regulate endoplasmic reticulum
(ER) exit-site (ERES) assembly and COPII-mediated
ER export are currently unknown. We analyzed the
role of phosphatidylinositols (PtdIns) in regulating
ER export. Utilizing pleckstrin homology domains
and a PtdIns phosphatase to specifically sequester
or reduce phosphorylated PtdIns levels, we found
that PtdIns 4-phosphate (PtsIns4P) is required to promote COPII-mediated ER export. Biochemical and
morphological in vitro analysis revealed dynamic
and localized PtsIns4P formation at ERES. PtdIns4P
was utilized to support Sar1-induced proliferation
and constriction of ERES membranes. PtdIns4P also
assisted in Sar1-induced COPII nucleation at ERES.
Therefore, localized dynamic remodeling of PtdIns
marks ERES membranes to regulate COPII-mediated
ER export.
capture by the COPII coat (Aridor et al., 1998, 2001; Bannykh et al., 1998; Barlowe, 2003; Kuehn et al., 1998).
Nevertheless, the affinities of coat-cargo interactions
are relatively low and may not support cargo recognition
by themselves (Miller et al., 2003; Mossessova et al.,
2003). Regulated enhancement of the binding avidity
between cargo and the coat at ERES may provide a plausible mechanism to control ER export. Thus targeted
assembly of COPII at defined ERES where cargo is
concentrated provides a mechanism to regulate COPIImediated cargo export from the ER.
How are ERES maintained as unique domains within
the fluid ER membranes? Lipid composition may regulate protein machinery needed to maintain ERES organization. Thus, identifying specific lipid signals that
dynamically mark ERES may provide key insight into
ERES function. Distinct phosphatidylinositols (PtdIns)
are known to function as specific regulators of protein
sorting and traffic in the late secretory and endocytic
pathways (De Matteis and Godi, 2004; Simonsen et al.,
2001; Wenk and De Camilli, 2004). Our previous studies
suggest a functional role for phospholipase D (PLD) activity in ER export (Pathre et al., 2003). Phosphorylation
of PtdIns is enhanced by PLD activity and can generate
specific lipid signals. Kinase (possibly lipid kinase) activity is required to regulate ER export (Aridor and Balch,
2000). We therefore hypothesized that spatially regulated phosphorylation of PtdIns supports ERES assembly and function.
Introduction
Results
The endoplasmic reticulum (ER) is a polarized compartment. Cotranslational insertion of newly synthesized
polypeptides takes place in the ribosome-bound rough
ER, whereas protein selection for export occurs from ribosome-free smooth membranes (Palade, 1975). Membrane protrusions are formed on smooth ER membranes
at ER exit sites (ERES) where cargo proteins destined for
export are concentrated (Bannykh et al., 1996). ERES
are distributed at the cell periphery and perinuclear
region (Aridor et al., 2004; Bannykh et al., 1996; Bevis
et al., 2002; Stephens, 2003). Vesicles and elongated
saccular/tubular carriers that protrude from ERES export cargo proteins to the Golgi complex (Bannykh
et al., 1996; Mironov et al., 2003). Cargo proteins are selected for export at ERES by the activity of the cytosolic
COPII coat, composed of the small GTPase Sar1 and the
Sec23/24 and Sec13/31 protein complexes (Antonny
and Schekman, 2001). COPII proteins are sequentially
recruited to ERES following the activation of Sar1
(Figure 1A) (Aridor et al., 1998; Matsuoka et al., 1998).
The recruited Sec23/24 complex exposes extensive
protein surfaces that present multiple peptide-binding
pockets (Miller et al., 2003; Mossessova et al., 2003).
These binding sites specifically recognize a variety of
ER export motifs on cargo proteins to mediate their
PtdIns Are Required to Support COPII-Mediated
ER Export
Specific PtdIns may serve as membrane codes to support ER export. Thus, we sequestered PtdIns formed
on ER membranes to test their role in vesicle formation
from the ER. We utilized an in vitro vesicle-budding
assay to follow the release of the biosynthetic cargo reporter tsO45 VSV-G from ER. tsO45 VSV-G (referred to
hereafter as VSV-Gts) is a temperature-sensitive protein
that displays a reversible folding defect. It is arrested
in the ER at a nonpermissive temperature (39.5 C) yet
synchronously folds trimerizes and exits the ER upon
shift to a permissive temperature (32 C) (Doms et al.,
1987).
Microsomes expressing VSV-Gts were incubated with
cytosol in the presence or absence of the polybasic antibiotic neomycin. At the end of incubation, the vesicle
fraction was separated from donor membranes by centrifugation, and the mobilization of VSV-Gts to vesicles
was determined by western blotting (Figure 1A). Neomycin, which binds and sequesters phosphorylated PtdIns,
inhibited vesicular export of VSV-Gts from the ER
(Figure 1B). However, neomycin has additional targets
that can affect ER export. Therefore, to examine the contributions of specific PtdIns to ER export, recombinant
GST-tagged pleckstrin homology (PH) domains with
high binding affinity for specific PtdIns were added to
the assay (Lemmon and Ferguson, 2001). Importantly,
Summary
*Correspondence: [email protected]
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672
Figure 1. The Fapp1-PH Domain Inhibits ER
Export
(A) Schematic representation of COPII coat
assembly and ER export. The release of
cargo proteins to the vesicular fraction was
measured by differential centrifugation. M
designates medium-speed donor-membranecontaining pellet (16,000 3 g). H designates
high-speed
vesicle
containing
pellet
(186,000 3 g).
(B–D) VSV-Gts-containing microsome membranes were incubated with cytosol for
30 min at 32 C in the presence or absence
of neomycin (1.5 mM) (B) or the indicated
GST-tagged PH domains (C and D), and the
release of VSV-Gts (B and C) or Bet1 (D) to
the vesicle fraction was analyzed.
(E–G) VSV-Gts-expressing microsomes were
incubated with cytosol (E) or purified COPII
components (Sar1 [1 mg], Sec23/24 [0.4 mg],
and Sec13/31 [6 mg], 40 ml final volume) (F
and G) in the presence of PLCd1-PH (E and
F) or Fapp1-PH (G) as indicated for 30 min
at 32 C. The release of VSV-Gts to the vesicular fraction was analyzed. The results are
representative of at least two independent
experiments.
GST-PH domains were tested at concentrations within
the range of their binding affinities to PtdIns containing
liposomes. GST-Fapp1-PH, which selectively interacts
with PtdIns4P (Dowler et al., 2000), inhibited VSV-Gts export at concentrations of 2–3 mM (Figures 1C and 1G and
titration in Figure 2D). GST-Fapp1-PH also inhibited the
mobilization of COPII vesicle SNARE protein Bet1 (Figure 1D), suggesting that sequestration of PtdIns4P led
to general inhibition of ER export. In contrast, addition
of GST, or GST fused to the PH domains of dynamin 2,
which has weak affinities for PtdIns(4,5)P2 and
PtdIns(3,4,5)P3 (Lee and Lemmon, 2001), or Grp1, which
has high affinity for PtdIns(3,4,5)P3, did not modulate
VSV-Gts export from the ER (Figure 1C). The PH domain
of PLCd1, which binds PtdIns(4,5)P2 (Lemmon and Ferguson, 2001), a possible product of PtdIns4P phosphorylation, inhibited ER export to a variable degree at higher
concentrations (7–15 mM) (Figure 1E). Direct analysis of
PtdIns phosphorylation suggested that kinase activities
in cytosol promoted PtdInsP2 synthesis (see below,
Figure 3F), yet ER export was effectively reconstituted
in the absence of cytosol with purified mammalian COPII
components (Aridor et al., 1998). Indeed, PLCd1-PH was
ineffective in inhibiting the budding reaction reconstituted without cytosol in the presence of purified COPII
proteins (Figure 1F). In contrast, full inhibition of vesicle
formation by GST-Fapp1-PH domain was observed at
concentrations as low as 2 mM, when budding was
reconstituted with purified COPII (Figure 1G). These results suggest that PtdIns4P may function as a possible
regulator of ER export.
The Inhibition of ER Export by Fapp1-PH Is Mediated
by PtdIns4P Binding
Fapp1 proteins localize to the TGN in NRK cells. TGN
localization is mediated by the Fapp1-PH domain and
is directed by specific interactions of the domain with
PtdIns4P lipids and ARF1 (Godi et al., 2004). We have
previously demonstrated that under the budding assay
conditions utilized, inhibition of ARF1 function by the
addition of high concentrations of dominant-negative
myrARF1(T31N) or by depletion of cytosolic COPI does
not affect the reconstituted COPII-dependent vesicle
formation step (Rowe et al., 1996). Indeed, ARF1 is not
present in the vesicle formation assay reconstituted
with purified COPII coat components under conditions
where full inhibition of ER export by Fapp1-PH is observed (Figure 2A). We did not observe specific interactions between Sar1 and any of the PH domains tested
(not shown). We thus analyzed whether the inhibitory
effect of Fapp1-PH on ER export is due to PtdIns4P
binding. Fapp1-PH added together with PtdIns4Pcontaining liposomes (but not PtdIns-free liposomes)
displayed little inhibition of ER export (Figure 2B). The
added liposomes may have augmented membrane
PtdIns4P content or simply sequestered Fapp1-PH; in
either case PtdIns4P binding was required for Fapp1PH function. We further mutated the PtdIns4P binding
site of Fapp1-PH, by replacing a conserved tryptophan
and arginine residues with asparagine and lysine
(W15N, R18K, mutated protein termed Fapp1-PHNK)
akin to the mutations that inhibit the recognition of
PtdIns(4,5)P2 by PLCd1-PH (Figure S1A; see the Supplemental Data available with this article online). Fapp1PHNK displayed significantly reduced affinity for
PtdIns4P as analyzed in protein-lipid overlay and liposome binding assays (Figures S1B–S1E). Consistent
with its reduced affinity for PtdIns4P, Fapp1-PHNK failed
to affect vesicle formation at concentrations of up to
5 mM (Figures 2C–2D). To further explore a possible
role for PtdIns4P in ER export, the synchronized mobilization of VSV-Gts from the ER was reconstituted in vitro
in semi intact cells and measured by indirect immunofluorescence (IF) (Plutner et al., 1992). VSVts-infected NRK
cells were permeabilized and incubated in the presence
of WT or mutated Fapp1-PH domains. Addition of rat
liver cytosol (RLC) quantitatively mobilized VSV-Gts
from the reticular ER to punctate vesicular tubular
Regulation of ER Export by Phosphoinositides
673
Figure 2. PtdIns4P Binding Mediates Fapp1-PH Inhibition of ER Export
(A) VSV-Gts-infected microsome membranes were incubated with purified COPII components (as in Figure 1) in the presence of GST-Fapp1-PH
domain as indicated for 30 min at 32 C. The mobilization of VSV-Gts to the vesicular fraction and the presence of ARF1 in the analyzed fractions
were determined by western blot. Recombinant ARF1 was loaded separately as a control.
(B) VSV-Gts-containing microsomes were incubated with cytosol in the presence of the indicated concentrations of GST-Fapp1-PH domain with
or without addition of liposomes (Lpsm) composed of PC or PC/PtdIns4P (420/50 mM) for 30 min at 32 C as indicated. The release of VSV-Gts to
the vesicular fraction is quantified in the graph below. Data are averaged from three independent experiments (means 6 SD).
(C) VSV-Gts-expressing microsomes were incubated with cytosol in the presence or absence of GST-Fapp1-PHWT or GST-Fapp1-PHNK for
30 min at 32 C, and the budding of VSV-Gts was determined.
(D) Quantitation of VSV-Gts released into vesicular fractions in the presence of varying concentrations of GST-Fapp1-PHWT and Fapp1-PHNK
domains. Data are means 6 SD derived from five independent experiments.
(E) VSVts-infected cells were permeabilized and incubated in the presence or absence of rat liver cytosol (RLC) GST-Fapp1-PHWT (3 mM) or GSTFapp1-PHNK (5 mM) as indicated for 30 min at 32 C. The mobilization of VSV-Gts from reticular ER (upper left image) to VTCs (upper right image)
was determined by IF. The images are a projection of five consecutive 0.1 mm optical sections. Scale bar is 5 mm.
(F) Liposomes (40 ml) composed of phosphatidylserine (500 mM) and PtdIns4P (100 mM) were incubated at 32 C for 3 min with indicated amounts
of the cytosolic domain (amino acid 1–507) of Sac or SacC392S mutant. Reactions were terminated by the addition of NEM (50 mM). Phosphate
release was quantified by a malachite green assay. Data are means 6 SD of triplicate samples.
(G) VSV-Gts microsomes (40 ml final volume) were incubated with cytosol in the presence or absence of of Sac1 (0.9 mg) or Sac1C392S (3.6 mg) as
indicated for 30 min at 32 C and the mobilization of VSV-Gts to the vesicle fraction was determined. The results are representative of at least three
independent experiments.
(H) VSVts infected cells were permeabilized and incubated (220 ml final volume) in the presence or absence of RLC and Sac1 (1 mg) as indicated for
30 min at 32 C. The movement of VSV-Gts from reticular ER (upper image) to VTCs (middle image) and the effects of Sac1 on VSV-Gts mobilization
(lower image) were determined by IF. Quantitation of VSV-Gts transport to VTCs at 20 and 30 min incubations in the presence or absence of the
indicated amounts of Sac1 is shown on the right panel. The number of VSV-Gts containing VTCs was derived from analysis of random fields of
cells by automatically counting 100 cells or more for each data point.
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674
Figure 3. Sar1 Recruitment and Activation on ER Membranes Stimulates PtdInsP and PtdInsP2 Formation
(A) ER enriched membrane fractions were incubated with buffer (control) or with Sar1-GTP (H79G) (2 mg, 60 ml final volume) for 15 min at 32 C.
Membranes were washed and incubated in the presence of g-32P-ATP and cytosol for the indicated times at 32 C. At the end of incubations,
lipids were extracted and separated by TLC. Phospholipids were quantified on a phosphor imager, and lipid markers were identified by iodine
vapors.
(B) Quantitation of (A) (means 6 SEM derived from four independent experiments).
(C) ER enriched membrane fractions, were incubated with buffer (control), Sar1-GTP, or Sar1-GDP (T39N, 2 mg of each, 60 ml final volume)
for 15 min at 32 C. Membranes were washed and further incubated for 3 min as described in (A). Data are means 6 SD of three independent
experiments.
(D) Membranes were incubated and lipids extracted as described in (C) except that orthovanadate (200 mM) was included in the second incubation to inhibit phosphatase activities.
(E) Quantitation of (D). Data represent means 6 SD of four independent experiments.
(F) Membranes were incubated in the absence or presence of Sar1-GTP (6 mg, 40 ml final volume) with g-32P-ATP in the presence or absence of
wortmannin (5 mM) or adenosine (500 mM) for 9 min at 32 C as indicated. Lipids were extracted and processed as described above.
(G) The effect of indicated inhibitors (analyzed as in [F]) was calculated by dividing the fold stimulation of PtdInsP formation by Sar1 (over control)
in the presence or absence of inhibitors, at time points at which peak PtdInsP formation was observed. Data are mean 6 SD of three to five
independent experiments.
clusters (VTCs) (Figure 2E, upper right panel). VSV-Gts
mobilization was markedly inhibited by neomycin (not
shown) and by the addition of GST-Fapp1-PH domain
(Figure 2E, bottom left panel). In contrast, GST-Fapp1PHNK did not inhibit the mobilization of VSV-Gts to
VTCs (Figure 2E, bottom right panel). Together, the
structure-function analysis and liposome rescue experiments suggest that the inhibitory effect of Fapp1-PH on
ER export is probably due to sequestration of PtdIns4P.
We tested whether reduction of PtdIns4P levels by the
yeast phosphatase Sac1 can modulate ER export. Sac1
is an ER and Golgi-localized enzyme that dephosphorylates the 3 and 4 positions of PtdIns3P or PtdIns4P, exerts low phosphatase activity toward PtdIns(3,5)P2,
does not act on PtdIns (4,5)P2, and exhibits in vivo preference for PtdIns4P (Hughes et al., 2000). We produced
the cytosolic domain of Sac1 (amino acid 1–507) and
tested its phosphatase activity by using a colorimetric
assay (Maehama et al., 2000). Sac1 efficiently dephosphorylated PtdIns4P present in PtdIns4P-containing liposomes, while a catalytic inactive mutant (Sac1C392S)
did not (Figure 2F). Sac1 inhibited vesicle budding
Regulation of ER Export by Phosphoinositides
675
from the ER, whereas the catalytically inactive mutant
(Sac1C392S) had no effect (Figure 2G). The inhibitory effect was reproduced in morphological ER export assays. Sac1 quantitatively delayed the cytosol-dependent mobilization of VSV-Gts to VTCs as analyzed by IF
in permeabilized VSV-Gts expressing cells (Figure 2H).
Importantly, Sac1 mediated delay of VSV-Gts mobilization to VTCs correlated with the inhibition detected in
budding assays (Figure 2G) to further support a role
for PtdIns4P in regulating ER export.
Sar1 Recruitment to ER Membranes Enhances
the Formation of PtdInsP and PtdInsP2
Sar1 recruitment to ER membranes was unaffected by
the sequestration of PtdIns4P (not shown). Therefore,
we analyzed whether Sar1 recruitment can affect
PtdIns4P levels. Sar1 binding and activation is specific
for ER membranes (Figure 6A). We nevertheless fractionated rat liver to enrich for ERES (Figure S2) and reduce the signal derived from lipid kinase activities present in crude membrane preparations. A two-stage assay
was utilized to analyze the phosphorylation state of
PtdIns in response to Sar1 activation (Figure 3). In stage
1, membranes were incubated in the absence or presence of GTPase-deficient Sar1 (Sar1-GTPH79G). The
membranes were washed and incubated (second stage)
in the presence of g-32P-ATP and cytosol. Subsequently, lipids were extracted and separated by thinlayer chromatography (TLC), and phosphorylated
PtdIns visualized by autoradiography. Recruitment of
Sar1-GTP to membranes caused a cyclic increase in
PtdIns mono- and bis-phosphate levels, with a maximal
(2-fold) effect observed at 3 min (Figures 3A, 3B, and 3C)
and subsequent increases at 9 min. This activity is similar to the previously reported transient elevation in
phosphorylated PtdIns observed upon incubation of
Golgi membranes with activated ARF1 (Godi et al.,
1999; not shown) and may reflect limiting PtdIns substrate levels that are dynamically regenerated on the
membranes. Activated Sar1 also increased 32P-phosphatidic acid (PA) levels generated by diacyl glycerol
(DAG) kinase activity (Figure 3B). The elevation in
PtdInsP and PtdInsP2 levels required Sar1 activation
as incubations with Sar1-GDPT39N, a Sar1 mutant deficient in GTP binding that cannot support ER export,
did not enhance PtdIns phosphorylation (Figure 3C)
(Aridor et al., 1995).
Sar1 could increase PtdIns levels through enhanced
PtdIns phosphorylation or through inhibition of PtdIns
phosphatase activities. To discriminate between these
possibilities, we analyzed PtdIns phosphorylation in
the presence of the phosphatase inhibitor orthovanadate. In the presence of orthovandate, incubations
with Sar1-GTP led to a 4-fold increase in PtdInsP and
PtdInsP2 levels over control, suggesting that kinase activities were enhanced under these conditions (Figures
3D–3E).
Sar1 also stimulated PtdInsP formation when measured in one-stage incubations carried out without
cytosol in the presence g-32P-ATP (up to 11-fold stimulation observed). Under these conditions, formation of
PtdInsP2 was largely diminished (Figure 3F). Therefore,
in agreement with our analysis of ER export (Figure 1G),
Sar1-regulated kinase activity that generates PtdInsP is
membrane associated. Pharmacological analysis indicated that unlike the PtdIns kinases (PI Kinases) activated by ARF1 (Godi et al., 1999), Sar1-induced PtdInsP
formation was insensitive to wortmannin, a drug that inhibits PI 3-kinases and type III PI 4-kinases (Figure 3F),
suggesting that these proteins are not involved in the
regulating ER export. In agreement, Sar1-induced phosphorylation of PtdIns was inhibited by adenosine, added
at equimolar concentration to ATP (Figures 3F and 3G).
This inhibition is characteristic of type II PI 4-kinases,
which display comparable affinities to ATP and adenosine (Balla et al., 2002). Therefore Sar1 activates a membrane-associated type II PI 4-kinase. The stimulation of
PtdInsP formation by Sar1 supports the role we uncovered for PtdIns4P on ER membranes (Figures 1 and 2).
Sar1 activation may provide coupling between COPII
assembly and PtdIns4P formation during ER export.
Dynamic and Localized Formation of PtdIns4P
at ERES
To provide a unique environment within fluid ER membranes, lipid remodeling should be dynamic. Because
Sar1 activation stimulated only limited PtdInsP,
PtdInsP2, or PA production (Pathre et al., 2003), lipid remodeling should also be localized to ERES in order to be
effective. To test these predictions, we reconstituted the
synchronized export of VSV-Gts from the ER in permeabilized VSVts-infected cells and visualized the dynamic
localization of VSV-Gts and PtdIns4P in real time by
time-lapse confocal video microscopy (Figure 4). VSVGts was labeled with a tagged Fab fragment of antiVSV-G antibody, and PtdIns4P formation was followed
by using recombinant GFP-Fapp1-PH (GST-GFPFapp1-PH) added at noninhibitory concentrations (0.2
mM). At the beginning of the experiment, VSV-Gts resided
in reticular ER membranes, and little apparent colocalization with the PtdIns4P reporter was observed. GFPFapp1-PH efficiently labeled the Golgi complex (Weixel
et al., 2005), which remained intact throughout the experiments (Figure 4). ER export was initiated with the
typical lag period of 5–7 min (Figures 4B and 4C)
required for VSV-Gts trimerization (Doms et al., 1987).
Export was detected by the elevation of VSV-Gts fluorescence intensity on ER membranes, reflecting the initial
concentration of VSV-Gts at ERES (Figure 4, and Movie
S1). Despite sequential pair-image acquisition, limited
sampling and single plane analysis utilized to ensure
the measurement of colocalization, PtdIns4P transiently
colocalized with VSV-Gts at ERES. At early time points,
colocalization of PtdIns4P and VSV-Gts was clearly
observed at peripheral ERES that reside on distinct reticular membranes remote from the juxtanuclear Golgi
complex (Figure 4C). At later time points, VSV-Gts was
depleted from the ER and efficiently mobilized to punctate VTCs, which for the most part did not colocalize
with Fapp1-PH (Figures 4D and 4E). Transient localization of PtdIns4P at reticular ER sites where VSV-Gts concentrates was also observed in subsequent rounds of
VSV-Gts export from the ER (Movie S1). The ability to visualize transient association of VSV-Gts with PtdIns4Pcontaining ER membranes during the synchronized
export of VSV-Gts from the ER provides direct evidence
to suggest that dynamic and localized lipid remodeling
at ERES supports vesicular traffic from the ER.
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676
Figure 4. Dynamics of PtdIns4P Formation at
ER Export Sites
Infected cells that accumulated VSV-Gts at
40 C in the ER were permeabilzed and incubated on ice for 20 min in the presence of
an Alexa 594-labeled Fab fragment of a
monoclonal antibody to VSV-Gts. The cells
were washed and shifted to 32 C to initiate
ER export in the presence of cytosol and
GFP-Fapp1-PH-domain (0.2 mM). Fluorescence pair images were acquired every
2 min by time-lapse spinning disc confocal
video microscopy. Sample times (min) are indicated for each panel on the right. Arrows
indicate colocalization between VSV-Gts and
PtdIns4P at ERES. The area labeled with the
large arrowhead in (B), (C), and (D) is enlarged
in the inserts. Note colocalization of VSV-Gts
and PtdIns4P at sites of initial VSV-Gts
concentration (B and C) during the mobilization of VSV-Gts from the reticular ER (A) to
mature VTCs (E). Transient colocalization of
PtdIns4P reporter with VSV-Gts is mainly observed in the initial stages of VSV-Gts trimerization and ER export (B and C). Scale bar is
5 mm.
Coat Assembly Is Regulated by PtdIns4P
VSV-Gts is mobilized through ERES by the activity of the
COPII coat. We therefore analyzed the colocalization of
Fapp1-PH with ERES stably coated with COPII. COPII
assembly on ERES was stabilized by the addition of
Sar1-GTP in semi-intact cells in reactions supplemented
with cytosol and GFP-Fapp1-PH. The localization of
PtdIns4P (using GFP-Fapp1-PH internal fluorescence)
with respect to ERES (Sec23) and the Golgi complex
(mannosidase II) was determined by confocal microscopy (Figure 5A). GFP-Fapp1-PH strongly labeled the
juxtanuclear Golgi complex (Weixel et al., 2005). COPII
coated ERES were localized within the Golgi region
and throughout the cell periphery. Substantial colocalization between ERES and PtdIns4P was observed. To
quantify the colocalization of PtdIns4P at ERES, permeabilized cells were incubated with GST-Fapp1-PH,
fixed, and probed for Sec23 and GST-Fapp1-PH
localization (Supplemental Experimental Procedures).
35% 6 1.5% (SEM, n = 55 cells) of peripheral Sec23-positive sites were colabeled with the Fapp1-PH domain.
Thus, in agreement with the dynamic localization of
VSV-Gts and Fapp1-PH on reticular ER during synchronized ER exit of VSV-Gts (Figure 4), PtdIns4P is localized
to ERES.
Can PtdIns4P be utilized to support COPII assembly?
We analyzed the effect of PtdIns4P sequestration on
Sar1-induced Sec23/24 recruitment to microsome
membranes. Under conditions in which ER export is inhibited, GST-Fapp1-PH or neomycin did not affect Sar1induced Sec23 membrane binding (not shown). To focus
on the role of membrane associated PI-kinases in regulating COPII assembly, we analyzed the recruitment of
purified Sec23/24 subunits by Sar1-GTP, in reactions
in which we restricted the levels of ATP (1 mM ATP instead of an ATP-regeneration system) to reduce the
activity of lipid kinases (Figure 5B). These limiting conditions significantly reduce Sar1-GTP dependent COPII
Regulation of ER Export by Phosphoinositides
677
assembly, but under these conditions, GST-Fapp1-PH
inhibited Sec23 recruitment (K1/2 of 12–15 mM). Furthermore, inhibition of Sar1-dependent coat recruitment by
GST-Fapp1-PH (K1/2 of 8 mM) was also observed in assays reconstituted with cytosol and restricted levels of
ATP, in which the membranes were subjected to high
salt wash (2.5 M urea) (Figure 5C). Therefore Sec23/24
recruitment to membranes is assisted by PtdIns4P.
However, the sensitivity of Sar1-dependent Sec23/24
recruitment to PtdIns4P sequestration by Fapp1-PH is
lower than that observed for ER export (Figure 1).
Can PI 4-kinase activity explain the dependency
of COPII assembly on ATP? We analyzed the ability of
PtdIns4P to support ATP-independent recruitment of
Sec23. Membranes were incubated with PtdIns4P micelles or with ATP or an ATP-regenerating system as
controls. Sar1-dependent Sec23 recruitment required
ATP and was markedly enhanced when an ATP-regenerating system was added (Figure 5D). Addition of
PtdIns4P led to ATP-independent Sar1-induced recruitment of Sec23/24. The added PtdIns4 micelles associated with or delivered PtdIns4P to ER membranes. In either case, added PtdIns4P provided binding sites for the
mammalian COPII, akin to the role of PtdIns4P in supporting the assembly of yeast COPII on synthetic liposomes (Matsuoka et al., 1998). PtdIns4P micelles did
not support robust Sec23 recruitment. Therefore dynamic PtdIns de- and rephosphorylation reactions may
regulate COPII assembly. Alternatively, additional ATPdependent steps that are independent of PtdIns phosphorylation may operate to facilitate COPII assembly.
The nucleation of COPII at ERES, rather than COPIImembrane binding, may be affected by PtdIns4P sequestration. We therefore used IF to examine the effect
of PtdIns4P sequestration on Sar1-induced nucleation
of COPII at ERES. COPII assembly was reconstituted
in semi-intact cells by the addition of cytosol and
Sar1-GTP in the presence of ATP-regeneration system.
Sar1-GTP induced robust recruitment of both Sec23
(COPII inner layer) and Sec13 (COPII outer layer) to
defined punctate sites (Figures 5E and 5F). Fapp1-PH
domain (Figure 5E) (5 mM) or neomycin (Figure 5F)
(1 mM), which did not affect COPII recruitment to microsome membranes (see above), markedly reduced Sar1induced nucleation of both COPII layers at ERES. It
could be that methodological differences between the
two assays resulted in the observed enhanced sensitivity of COPII recruitment to PtdIns4P sequestration as
detected by IF. Alternatively, the inhibition of COPII recruitment in the morphological assay may reflect inhibition of COPII nucleation at ERES, whereas the overall
levels of recruited COPII as measured on microsome
membranes were mildly affected.
PtdIns4P Marks ERES and Is Required to Support
Carrier-Membrane Fission
Interfering with the dynamics of ERES assembly may result in the formation of stabilized ERES intermediates
that precede coat assembly and accumulate PtdIns4P.
We therefore generated enlarged ERES for further morphological analysis. Permeabilized VSV-Gts-expressing
cells were incubated with Sar1-GTP in the absence of
cytosol. Under these conditions, highly elongated and
constricted Sar1-induced ERES that selectively concen-
trate cargo are formed (see VSV-Gts, Figures 6B and 6D)
(Aridor et al., 2001). These tubular transport intermediates are formed independently and prior to COPII coat
assembly. Sar1 is targeted exclusively to these elongated tubules that are well segregated from the Golgi
complex (Figure 6A). Sar1-GTP-dependent tubules
were generated in the presence of low concentrations
of either GST-Fapp1-PH or anti-PtdIns4P antibodies,
which provide an alternative independent probe for
PtdIns4P localization. PtdIns4P visualized by both
PtdIns4P reporters decorated VSV-Gts containing elongated tubular ERES (Figures 6B and 6D) as well as Golgi
structures (arrowheads in Figure 6B and not shown). The
distinct colocalization between PtdIns4P and cargo
in tubular export domains suggests that the lipid composition of ERES differ from that of the bulk of ER
membranes. To analyze the possible contribution of
PtdIns4P to the formation of Sar1-dependent tubular
ERES, we increased the concentration of the GSTFapp1-PH domain from 0.6 mM to 5 mM (Figure 6C). Strikingly, while export sites marked by the enrichment of
VSV-Gts (Figure 6C) and Sar1 (not shown) were still observed, the elongation of these domains into tubular
structures was prevented. The colocalization between
VSV-Gts and the PtdIns4P reporter was preserved. Similar results were obtained with neomycin (see Sar1-decorated tubules, Figure 6E). We have recently provided
evidence to suggest that Sar1-mediated membrane
constriction and tubulation at ERES controls vesicle fission (Bielli et al., 2005). Thus, PtdIns4P in ERES tubules
may support Sar1-induced tubule constriction and elongation, a step that precedes vesicle fission.
Discussion
The concentration and export of cargo via organized
ERES provides a facet of selectivity to the ER-sorting
machinery of higher eukaryotic cells (Aridor et al., 2004;
Bevis et al., 2002). The molecular basis for ERES formation and function is unknown. We hypothesized that domains of distinct lipid composition are established in
fluid ER membranes to define ERES. We now demonstrate that PtdIns4P-containing domains are dynamically generated to regulate ERES membrane deformation and coat nucleation to support ER export.
Several independent lines of evidence support our
conclusions. A specific PtdIns-binding domain that sequesters PtdIns4P inhibited ER export (Figure 1). Structure-function analysis and lipid-binding competition
correlated the inhibitory effects of the domain with specific lipid recognition (Figure 2 and Figure S1). The cytosolic domain of the ER-localized PtdIns4P phosphatase
Sac1 delayed ER export (Figure 2). Direct analysis of
PtdIns phosphorylation on ER membranes demonstrated that recruitment and activation of Sar1, which
initiates ER export, transiently elevated PtdInsP and
PtdInsP2 levels (Figure 3). These biochemical results
correlated well with in vitro real-time imaging, which
demonstrated that PtdIns4P accumulated transiently
and locally during the concentration of cargo at ERES
at sites of COPII assembly and membrane constriction
(Figures 4, 5, and 6).
The recognition that PtdIns are regulators of endocytic
traffic led to the identification of PtdIns(4,5)P2-binding
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678
Figure 5. The Recruitment of Sec23 to ERES Is Assisted by PtdIns4P
(A) Permeabilized cells were incubated in the presence of Sar1-GTP (6 mg, 220 ml final volume), RLC, and GFP-Fapp1-PH (3 mM) for 30 min at 32 C.
The cells were fixed and stained for Sec23 and mannosidase II (Man II) as indicated. The localization of GFP-Fapp1-PH (by GFP fluorescence)
ERES (Sec23, red) and the Golgi complex (Man II, blue) was determined. Merge images for Man II and GFP-Fapp1-PH, Sec23 and Man II, and
a merged image for all three markers are shown. Arrows indicate colocalization of Fapp1-PH with peripheral Sec23-containing ERES. Scale bar is
5 mm.
(B) Microsomes were incubated with purified Sec23/24 complex (0.7 mg, 60 ml final volume) and 1 mM ATP at 32 C for 15 min in the presence or
absence of Sar1-GTP (0.25 mg) and GST-Fapp1-PH domain as indicated. The membrane recruitment of Sec23 was determined by western blot
with affinity-purified anti-Sec23 antibody.
(C) Microsomes were washed in buffer containing 2.5 M urea for 30 min on ice. The membranes were reisolated by centrifugation and incubated
with RLC and 1 mM ATP as in (B), in the presence or absence of Sar1-GTP (0.1 mg) and GST-Fapp1-PH as indicated, and membrane recruitment
of Sec23 was determined.
(D) Microsomes were incubated with RLC at 32 C for 15 min in the presence or absence of Sar1-GTP (0.3 mg, 60 ml final volume), ATP-regeneration
system (indicated as reg.), 1 mM ATP, or 50 mM PtdIns4P micelles as indicated, and the recruitment of Sec23 was determined. Data are representative of three independent experiments.
Regulation of ER Export by Phosphoinositides
679
Figure 6. PtsIns4P Is Formed and Accumulated on Sar1-Induced Tubules
(A) Permeabilized cells (220 ml final volume) were incubated in the presence of Sar1-GTP (6 mg) for 30 min at 32 C. The cells were fixed and stained
for Sar1 and giantin (merged images are shown as indicated).
(B and C) VSVts-infected cells were permeabilized cells and incubated as in (A) in the presence of 0.6 mM (B) or 5 mM (C) of GST-Fapp1-PH for
30 min at 32 C to form VSV-Gts-containing tubular ERES. The cells were fixed and stained with VSV-Gts and GST-Fapp1-PH as indicated.
(D) Permeabilized cells were incubated with anti-PtdIns4P antibodies as in (A). The cells were fixed and further stained for VSV-Gts. The localization of VSV-Gts and the anti-PtdIns4P antibody was determined by IF. A projection of five consecutive optical sections is presented. Bars are
10 mm. Note the colocalization of VSV-Gts and PtdIns4P reporters on Sar1 tubular domains (arrow in [B]). Arrowheads in (B) point to juxtanuclear
Golgi elements positive for Fapp1-PH.
(E) Permeabilized cells were incubated as in (A) in the presence or absence of neomycin (1 mM). The cells were fixed stained for Sar1. Note
inhibition of Sar1 tubule elongation with neomycin or Fapp1-PH.
accessory proteins that support clathrin-mediated endocytosis (Wenk and De Camilli, 2004). PtdIns(4,5)P2
may function in place of small GTPases to support the
membrane binding of AP-2/clathrin. However, PtdIns4P
is required for ARF1 regulated AP-1/clathrin membrane
binding and both PtdIns4P and PtdIns(4,5)P2 are required to support VSV-G exit from the TGN (Wang
et al., 2003). Our studies suggest that utilization of PtdIns
signals may be a general mechanism to define sites for
regulated cargo sorting and vesicle formation.
Both PtdInsP and PtdInsP2 levels were elevated by
Sar1 activation on ER membranes (Figure 3). Membrane-associated type II PI 4-kinase is activated by
Sar1 in the absence of cytosol under conditions that
do not support efficient formation of PtdInsP2. Accord-
ingly, the PtdIns(4,5)P2-binding domain of PLCd1-PH effectively inhibited ER export only in the presence of cytosol under conditions that promote efficient PtdInsP2
formation (Figure 1 and 3). Thus, PtdIns4P itself is a signal that regulates ER export. However, subsequent
formation of PtdIns (4,5)P2 may be utilized in a positive
feedback loop activating PLD (Pathre et al., 2003) and
enhancing COPII vesicle formation.
Active populations of type II PI 4-kinase a and b are localized to the ER (Waugh et al., 2003; Wei et al., 2002).
Future studies will address the possible role of these
enzymes in regulating ER export. Interestingly, type II
PI 4-kinases require detergent-induced perturbation of
membranes for activation. Sar1 utilizes its NH2 terminal
amphipathic domain to perturb and deform membranes
(E–F) Permeabilized cells (220 ml final volume) were incubated with RLC and ATP-regeneration system in the presence or absence of Sar1-GTP
(6 mg) and GST-Fapp1-PH domain (5 mM) (E) or neomycin (1 mM) (F) as indicated for 30 min at 32 C. The cells were fixed and stained for Sec23 (E)
or Sec23 and Sec13 (F). Images for each experiment were acquired under identical conditions. Scale bar is 5 mm.
Developmental Cell
680
(Bielli et al., 2005; Lee et al., 2005). An intriguing possibility is that the membrane-deforming activity of Sar1 provides a mechanism to couple Sar1-induced membrane
curvature with the activation of PI 4-kinase signaling
required to regulate ER export.
While the major PI 4-kinase activities in budding yeast
(PiK1 and Stt4) are not localized to the ER and do not
regulate ER export (Audhya et al., 2000), recent studies
have demonstrated that deletion of the phosphatase
Sac1, which is mainly localized in the ER, leads to elevated levels of PtdIns4P in yeast ER (Tahirovic et al.,
2005). Elevation of PtdIns4P due to rapid inactivation
of Sac1p does not inhibit ER export, yet prolonged imbalance of PtdIns4P hydrolysis in Sac1-deleted strains
culminate in defects in ER-to-Golgi transport (Foti
et al., 2001). Sac1 mutations display negative genetic interactions with genes encoding for COPII inner and
outer layers (Sec23 and Sec13) (Cleves et al., 1989).
Therefore, PtdIns4P may regulate some aspects of the
yeast ER. However, the early secretory pathway in budding yeast markedly differs from that of mammalian cells
as it lacks organized ERES and Golgi structures. Organized mammalian ERES may have evolved to further utilize PtdIns signals. The transient nature of PtdIns4P formation observed in our morphological and biochemical
analysis suggests that tight spatial and temporal control
of PtdIns4P levels by both PI kinases and phosphatases
is required to regulate the nucleation of COPII at ERES
on the fluid ER membrane.
Lipid signals have previously been implicated in ERES
function (Fabbri et al., 1994; Lee et al., 2004; Nagaya
et al., 2002; Pathre et al., 2003; Runz et al., 2006; Shimoi
et al., 2005). In addition to PtdIns (current work), PA, diacylglycerol (DAG), and sterols regulate ER export. DAG
kinase d may regulate ER export, and PLCd4 that generates DAG is localized to the ER. p125, a PA-preferring
phospholipase A1 homolog that binds Sec23, is recruited by Sar1 to regulate the cellular localization of
ERES. Therefore, PtdIns signals can be viewed as part
of an extensive lipid remodeling cascade that regulates
ER export.
How is the PtdIns4P signal decoded at ERES? COPII
assembly is assisted by PA, and a preference for
PtdIns4P was revealed for Sar1-dependent COPII binding to synthetic liposomes (Matsuoka et al., 1998; Pathre
et al., 2003). Thus, COPII subunits may participate in decoding the PtdIns4P signal. However, our analysis suggests that the information encoded by PtdIns4P is also
utilized in steps that precede coat binding perhaps by
accessory proteins that assist in ERES assembly. Our
results suggest that the lipid composition of ERES may
resemble that of the Golgi complex rather than that of
the bulk ER and thus may utilize functionally similar
PtdIns regulated proteins for organization. One possible
accessory protein that regulates the Golgi structure in
a PtdIns-dependent manner is b III spectrin (Siddhanta
et al., 2003). Fragments of spectrin inhibit ER export in
vivo and in vitro (Devarajan et al., 1997; Godi et al.,
1998). Lipid transfer proteins play a key role in regulating
lipid distribution at the Golgi complex. Some, including
FFAT-motif-tagged proteins like the PH-domain containing OSBP-related protein 9, localize to ERES (Wyles
and Ridgway, 2004), and their ER localized receptors
VAP-B and VAP-A participate in regulating the organiza-
tion of the ER and ER export (Amarilio et al., 2005).
Therefore, a possible role for such receptors and members of their PtdIns-interacting protein ligands in regulating the micro-organization of ER membranes and
ER export may be envisioned.
Our results point to a functional role for PtdIns4P in
the formation and elongation of Sar1-induced tubules
that proliferate from ERES and serve as intermediates
in vesicle fission (Aridor et al., 2001; Bielli et al., 2005).
Fission-intermediate tubular domains have also been
observed during export from the TGN (Liljedahl et al.,
2001). Fapp proteins (Fapp1 and Fapp2) localize to the
TGN through PtdIns4P binding and propagate the formation of TGN tubules to control TGN export (Godi
et al., 2004). PtdIns4P may regulate functionally homologous PtdIns4P-binding Fapp-like proteins to control
tubule elongation at ERES. Such proteins may utilize
lipid binding/transfer activities to achieve local lipid
composition favorable for coat nucleation and vesicle
fission (Vieira et al., 2005).
The targets that decode PtdIns4P at ERES remain to
be defined. The demonstration of localized formation
and function of PtdIns4P at ERES provides a basis to
study how these unique domains are established within
the ER membrane to support and regulate ER export.
Experimental Procedures
Materials
Recombinant proteins and antibodies utilized in the study are described in Supplemental Data. Phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), and brain PtdIns
were from Sigma. Dilauryl-PA, PtdIns, PtsIns4P, and PtdIns(4,5)P2
were from Avanti polar lipids. Lipid arrays and PtdIns4P utilized for
micelle preparations were from Echelon, g-32P-ATP (3000 Ci/mmol)
from ICN, TLC plates were from Voigt Global Distribution, LLC, Digitonin from Wako, and Malachite Green Phosphate assay kit from
BioAssay Systems. Tissue culture reagents were from Gibco. Other
reagents were from Sigma.
Infection of NRK Cells
NRK cells were infected with tsO45 VSV as described (Plutner et al.,
1992).
Preparation of Membrane Microsomes
Microsomes from VSVts-infected or -uninfected NRK cells were prepared as described (Rowe et al., 1996).
Coat Recruitment Assay
Sec23 recruitment assays were performed as described (Aridor and
Balch, 2000; Aridor et al., 1998). Salt wash details are described in
the Supplemental Data.
ER-Vesicle Budding Assays
COPII budding assays were performed as described (Aridor et al.,
1998; Rowe et al., 1996).
Semi-Intact Cell Morphological Analysis and Microscopy
Morphological analysis of ER export was performed as described
(Aridor et al., 2001). Images were acquired on an Olympus Fluoview
500 confocal microscope. Real-time imaging conditions are described in the Supplemental Data. Quantitation of colocalization between GST-Fapp-PH with Sec23 and the mobilization of VSV-Gts to
VTCs is described in the Supplemental Data. Alexa 594-tagged Fab
fragment of anti-VSV-G antibody (P5D4) was added during incubations to label VSV-G in ER export assays, which quantify the effects
of Sac1 on mobilization of VSV-Gts to VTCs.
Regulation of ER Export by Phosphoinositides
681
Measurement of Phosphoinositide Formation on Isolated ER
Membranes
Measurement of phosphoinositide formation on isolated ER membranes is described in the Supplemental Data.
Measurement of Phosphatase Activity of Soluble Sac1
Phosphatase activity of Sac1 was determined as described
(Maehama et al., 2000).
Supplemental Data
The Supplemental Data include analysis of lipid-binding properties
of wt and mutated Fapp1-PH domain (Figure S1), properties of fractionated rat liver membranes (Figure S2), and the dynamics of
PtdIns4P formation at ER exit sites depicted in a movie. The supplemental text includes additional details on experimental procedures
utilized in the study. Supplemental Data are available at http://
www.developmentalcell.com/cgi/content/full/11/5/671/DC1/.
Acknowledgments
We thank Drs. T. Balla (National Institutes of Health), W.E. Balch (The
Scripps Research Institute), M.A. Lemmon (University of Pennsylvania), and G.S. Taylor (University of Nebraska) for valuable reagents.
The study was supported by grants from the Edward Mallinckrodt Jr.
Foundation (M.A.), NIH DK062318 (M.A.), NIH DK064613 (O.A.W.),
American Heart Association (A.B.P.), and NIH T32-DK61296
(K.M.W.).
Received: June 28, 2005
Revised: June 25, 2006
Accepted: September 4, 2006
Published: November 6, 2006
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