1 Department of Physiology, University of Tennessee Health Science Center, Memphis, Tennessee 38163; 2 Department of Medicine, Johns Hopkins University, Baltimore, Maryland 21205; and 3 Laboratory of Cell Signaling, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892
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ABSTRACT |
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The actin-regulatory protein villin is tyrosine
phosphorylated and associates with phospholipase C-1
(PLC-
1) in the brush border of intestinal
epithelial cells. To study the mechanism of villin-associated
PLC-
1 activation, we reconstituted in vitro the tyrosine
phosphorylation of villin and its association with PLC-
1. Recombinant villin was phosphorylated in vitro by
the nonreceptor tyrosine kinase c-src or by expression in the TKX1 competent cells that carry an inducible tyrosine kinase gene. Using in
vitro binding assays, we demonstrated that tyrosine-phosphorylated villin associates with the COOH-terminal Src homology 2 (SH2) domain of
PLC-
1. The catalytic activity of PLC-
1
was inhibited by villin in a dose-dependent manner with half-maximal
inhibition at a concentration of 12.4 µM. Villin inhibited
PLC-
1 activity by sequestering the substrate
phosphatidylinositol 4,5-bisphosphate (PIP2), since
increasing concentrations of PIP2 reversed the inhibitory effects of villin on PLC activity. The inhibition of
PLC-
1 activity by villin was reversed by the tyrosine
phosphorylation of villin. Further, we demonstrated that tyrosine
phosphorylation of villin abolished villin's ability to associate with
PIP2. In conclusion, tyrosine-phosphorylated villin
associates with the COOH-terminal SH2 domain of PLC-
1
and activates PLC-
1 catalytic activity. Villin regulates
PLC-
1 activity by modifying its own ability to bind
PIP2. This study provides biochemical proof of the
functional relevance of tyrosine phosphorylation of villin and
identifies the molecular mechanisms involved in the activation of
PLC-
1 by villin.
phosphatidylinositol 4,5-bisphosphate; tyrosine phosphorylation; cytoskeleton
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INTRODUCTION |
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WE PREVIOUSLY
DEMONSTRATED for the first time that villin, an actin-regulatory
protein of epithelial cells, is tyrosine phosphorylated in vivo in
response to receptor activation (22). Since then several
other proteins of the villin superfamily, including gelsolin, CapG, and
fragmin, have been shown to be tyrosine phosphorylated in vitro
(11). Although the in vivo tyrosine phosphorylation of
these proteins remains to be elucidated, the in vitro studies and our
own observations with villin suggest that tyrosine phosphorylation may
be a feature common to all members of this family. That the tyrosine
phosphorylation of villin may be functionally significant is suggested
by the fact that tyrosine-phosphorylated villin disassociates from the
cytoskeleton (21) and forms a complex with the
brush-border membrane-bound phospholipase C-1
(PLC-
1) (22). These observations suggest a
role for villin, not only in actin organization but also in the
PLC-
1-mediated signal transduction. Because
tyrosine-phosphorylated villin associates with PLC-
1
(22), our hypothesis is that tyrosine phosphorylation of
villin is intimately linked to regulation of PLC-
1
catalytic activity. Villin can bundle, sever, cap, or nucleate actin
filaments in a Ca2+- and phospholipid-dependent manner
(18). For over a decade, it has been known that villin
binds phosphatidylinositol 4,5-bisphosphate (PIP2) and that
the association of villin with PIP2 inhibits villin's actin-severing property (19). This observation lends
further support to the idea that villin may regulate both the cortical cytoskeleton and phosphoinositide-mediated signal transduction pathways.
In this study we attempt to understand the molecular steps involved in
the tyrosine phosphorylation of villin, its association with both the
substrate and the enzyme, namely, PIP2 and
PLC-1, respectively, and thus its regulation of
phosphoinositide-mediated signal transduction pathways. Using
recombinant villin and purified PLC-
1, we reconstituted
in vitro our previous in vivo observation (22). The
advantage of reconstitution is that it allows selected and controlled
conditions to be defined to explore the molecular basis of the
phenomenon, thus making it easier to interpret the observations. The
complexity of the various actin-remodeling abilities of villin, in
addition to its association with several signaling molecules, makes
such an approach most useful to dissect the in vivo function of villin.
Our previous work in the intestinal epithelial cells and the recent
observation in the opossum kidney cell line demonstrate that tyrosine
phosphorylation of villin and its association with PLC-1
regulate changes in the microfilament structure that are crucial to the
vectorial function of ion transport in epithelial cells (22,
28). Both PLC-
1 and the microfilament network have been demonstrated to regulate ion transport functions in epithelial cells of the intestine and kidney (Refs. 22 and
28; for review see Ref. 21); however, our understanding of
the mechanisms involved remains rudimentary. This study may help us
understand the molecular and cellular mechanisms by which
PLC-
1 and the microfilament network regulate cell
morphology and function in epithelial cells.
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METHODS |
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Materials.
PLC-1 was purified from HeLa cells that had been
transfected with recombinant vaccinia virus containing a full-length
sequence of PLC-
1 (2). Native villin,
purified from chicken brush-border membranes (stored in a buffer
containing 0.5 mM
-mercaptoethanol), was a kind gift from Dr.
S. W. Craig (Johns Hopkins University, Baltimore, MD). Monoclonal
antibodies to villin, nck, and grb-2 were from Transduction
Laboratories; monoclonal antibodies to phosphotyrosine, clone PY20,
were from ICN; monoclonal antibodies to PIP2 were from
Perseptive Biosystems (Boston, MA); PIP2 and phosphatidylethanolamine (PE) were from Avanti;
[3H]PIP2 was from NEN; all other chemicals
were from Sigma. Recombinant c-src kinase expressing the kinase domain
was purchased from UBI. Glutathione Sepharose 4B fast flow was from
Amersham-Pharmacia; GelCode blue was from Pierce; BL21 and TKX1
competent cells were from Stratagene. The nonmuscle actin
polymerization kit was purchased from Cytoskeleton (Denver, CO).
Preparation of GST fusion proteins.
Recombinant proteins of the SH2 and SH3 domains of PLC-1
were made as described earlier (2). Both domains were
amplified by PCR using primers containing restriction sites at the
ends. Amplicons were digested with their respective enzymes and cloned into pGEX-2T. Amino acid sequences of the NH2- and COOH-terminal SH2
and SH3 domains correspond to residues 550-657 (N-SH2),
668-745 (C-SH2), 550-745 [(N + C)-SH2], and
758-851 (SH3). Escherichia coli BL21 cells were
transformed with the constructs and cultured at 37°C. Expression of
the fusion proteins was induced with 0.1 mM
isopropyl-
-D-thiogalactopyranoside (IPTG), and the cells
were harvested by centrifugation at 2,000 g for 15 min,
sonicated in phosphate-buffered saline (PBS), and centrifuged at 5,000 g for 15 min. The resulting supernatant was mixed with 2 ml
of a 50% (vol/vol) slurry of glutathione Sepharose 4B and incubated at room temperature for 30 min. After centrifugation the supernatant was
removed, and the resin was washed with 50 bed volumes of PBS. This
procedure was repeated twice. Glutathione S-transferase
(GST)-fused recombinant proteins were eluted from the beads
using 5 mM reduced glutathione. The eluted proteins were dialyzed, and
the purity of the proteins was determined by separating the proteins by
SDS-PAGE and staining the gel with Coomassie blue or GelCode blue.
Full-length villin (human) cDNA cloned in pGEX-2T was also purified as
described above.
Polymerization kinetics of actin using recombinant and native villin. The effects of villin on the polymerization of actin are well documented (9). In the presence of low concentrations of neutral salts, G-actin polymerizes to form long, double-stranded F-actin filaments. The kinetics of filament formation can be followed by measuring the change in viscosity of the protein. Actin polymerization by recombinant and native villin (purified from chicken brush-border membranes) was measured using the Ostwald capillary viscometer (type 100 Cannon Instrument, PA). Negative controls included GST protein and the buffer cocktail. Specific viscosity is defined as flow time of sample solution divided by flow time of the corresponding buffer, minus 1 (9).
The kinetics of actin polymerization were also determined using a nonmuscle actin polymerization kit according to the instructions of the manufacturer. The basis of this assay is that the fluorescence intensity of pyrene actin is much greater for polymeric than monomeric actin. (29). The ability of villin to nucleate actin assembly or to sever actin filaments was determined by its effect on the rate and extent of increase or decrease, respectively, of fluorescence of pyrene-labeled actin. G-actin (6 µM) in buffer containing 5 mM Tris · HCl, 0.2 mM ATP, and 0.2 mM CaCl2 was preincubated with native or recombinant villin (60 nM) for 10 min on ice. Polymerization was induced by the addition of 150 mM KCl and 1 mM MgCl2. The increase in fluorescence that occurs when pyrene G-actin forms pyrene F-actin was measured over time. The rate at which actin polymerizes depends on the concentrations of free actin monomers and the filament ends. Because villin complexes with G-actin faster than spontaneous actin nuclei can form, the initial rate of polymerization determined from the rate of fluorescence increase is proportional to the number of pointed end nuclei formed and, therefore, the relative nucleation activity of villin (29). For assays of filament-severing activity, a sample of pyrene-labeled F-actin was diluted below its critical monomer concentration into solutions containing villin (60 nM). Because actin filaments depolymerize only from their ends, the rate of fluorescence decrease, proportional to the depolymerization rate, depends on the number of ends and, therefore, on the number of cuts introduced by villin (29). Fluorescence measurements were performed at 25°C using the Fluorolog 3 fluorimeter. The excitation wavelength was set at 365 nm, and the emission wavelength was set at 388 nm.In vitro kinase assay.
Recombinant villin (concentrations indicated in RESULTS)
was phosphorylated in vitro by c-src (2.5 U) in an assay mixture containing 50 mM Tris · HCl, pH 7.5, 150 mM NaCl, 0.5 mM EGTA, 20 mM MgCl2, 20 µM ATP, and 2-5 mM
-mercaptoethanol. Villin was phosphorylated for 60 min at 37°C,
and the reaction was stopped by the addition of Laemlli sample buffer.
Tyrosine-phosphorylated proteins were separated by SDS-PAGE and
transferred to nitrocellulose membrane, and a Western analysis was done
using phosphotyrosine monoclonal antibodies.
Tyrosine phosphorylation of villin in TKX1 cells.
Full-length villin cDNA (human) cloned in pGEX-2T was expressed in the
Epicurian coli TKX1 cells. TKX1 cells carry a
plasmid with the elk tyrosine kinase (tk) gene controlled by the trp
promoter. The elk tyrosine kinase has broad specificity and has been
shown to tyrosine phosphorylate a number of proteins in Epicurian
coli (31). A two-step protocol involving
first the induction of expression of villin protein gene (by addition
of IPTG) followed by induction of the tk gene [by addition of
3--indoleacrylic acid (IAA)], allowed for the accumulation of
GST-tagged tyrosine-phosphorylated villin (VILT/WT). Phosphorylated
villin was affinity purified from bacterial lysates using a glutathione
Sepharose 4B column. The proteins were eluted with 5 mM glutathione in
1-ml fractions. Purity of the fractions was assessed by SDS-PAGE and
staining with GelCode Blue. Tyrosine phosphorylation of villin was
determined by Western analysis of the samples using phosphotyrosine
monoclonal antibodies. TKX1 cells cultured in the absence of IAA were
used to obtain nonphosphorylated villin (VIL/WT).
Preparation of lipids.
PIP2 and PE obtained as chloroform solutions were dried in
a stream of nitrogen, and micelles were prepared by dissolving the
dried lipid in 10 mM Tris · HCl, pH 8.0, at a concentration of
3.4 mg/ml. The lipids were sonicated in a Branson 1210 sonicator for 5 min at maximum power. The lipids were stored in aliquots at 80°C,
and before use the lipids were sonicated for an additional 5 min.
Association of villin with PIP2. The association of villin with PIP2 was determined by a procedure described by Fukami et al. (15). Briefly, nonphosphorylated villin (VIL/WT, 1 µM) and phosphorylated villin (VILT/WT, 10 µM) were incubated with PIP2 (100 µM) for 15 min at 37°C in a buffer containing 140 mM NaCl, 1 mM EGTA, 0.5 mM dithiothreitol (DTT), 0.5 mM NaN3, 0.05% Tween 20, 20 mM MOPS-Tris, pH 7.4, and 0.2% BSA. The recombinant proteins were separated by SDS-PAGE and transferred to nitrocellulose, and a Western analysis was done with monoclonal antibodies to PIP2 (1).
The lipid binding assay described by Touhara et al. (36) was used to determine the association of PIP2 with villin. Briefly, fusion proteins (VIL/WT, VILT/WT, and GST, 2 µg) were incubated with PIP2 vesicles (final concn 1.7 mg/ml), and the reaction mixture was diluted to a final volume of 40 µl with PBS. After incubation at room temperature for 10 min and on ice for 5 min, the tubes were centrifuged at 100,000 rpm for 15 min at 4°C. The supernatant (S) was saved, and the pellet (P) was rinsed once with PBS. The pellet was resuspended in 60 µl of laemmli sample buffer, and 20 µl of this was loaded on a gel. The total (T, 2 µg protein) and 10 µl of the supernatant were mixed with 10 µl of sample buffer and loaded on the same gel. The gels were stained with GelCode blue, and the amounts of protein in T, S, and P were compared using the Eagle Eye II imager system (Stratagene). Alternatively, the procedure described by Davletov and Sudhof (10), with minor modifications, was used to determine the association of PIP2 with villin. Twenty micrograms of fusion proteins bound to glutathione Sepharose beads were resuspended in 100 µl of buffer containing 50 mM HEPES, pH 7.2, 0.1 M NaCl, 0.5 mM EGTA (incubation buffer), and lipid vesicles containing 0.8 mg/ml of PIP2 and 3H-labeled PIP2 (20,000 cpm). The mixture was incubated at room temperature for 15 min with shaking. The fusion proteins bound to GST Sepharose were collected by centrifugation and washed three times with 1 ml of the incubation buffer. Lipid binding was quantified by liquid scintillation counting of the beads.Preparation of ileal brush-border membranes. Distal ileum from New Zealand male rabbits was used to prepare brush-border membrane fractions by a method of differential centrifugation and Mg2+ precipitation, as described earlier (22). Rabbit distal ileum was exposed to carbachol (1 µM, 30 s), conditions previously shown to lead to tyrosine phosphorylation of villin (21). Brush-border membranes from control tissue were made in parallel (22).
In vitro binding studies.
GST fusion proteins of the SH2 and SH3 domains of PLC-1
immobilized on glutathione Sepharose 4B were incubated for 1 h at 4°C with brush-border membrane extracts containing
tyrosine-phosphorylated villin (prepared from carbachol-treated ileum,
1 µM, 30 s) or nonphosphorylated villin (prepared from control
ileum). The recombinant proteins were collected by centrifugation and
washed several times using a buffer containing 1% Triton X-100 and 150 mM NaCl. The samples were separated by SDS-PAGE and transferred to
nitrocellulose, and a Western analysis was done using monoclonal
antibodies to villin, phosphotyrosine, or GST.
PLC activity assay.
The PLC-1 activity was measured using small unilamellar
vesicles of 3H-labeled PIP2 as described
earlier (2). The assay mixture contained ~10 µg of
PIP2, 0.8 mM sodium desoxycholate, ~20,000 cpm of
[3H]PIP2, 3 mM MgCl2, 100 mM
NaCl, 2 mM EGTA, 2 mM CaCl2 (final free Ca2+
concn of 45 µM), and 0.1 mM DTT in 50 mM HEPES buffer, pH 7.0. For
preparation of the substrate solution, PIP2 (0.75 mg) and [3H]PIP2 (1.5 µCi) were mixed and dried
under a stream of nitrogen. Five milliliters of 65 mM HEPES (pH
7.0)-100 mM NaCl were added to the dried lipids, and the samples were
sonicated for six 3-min intervals interspersed with periods of cooling
on ice under a stream of nitrogen. Purified PLC-
1 (40 ng) was used as the source of the enzyme. Recombinant villin was
incubated with the substrate before the addition of
PLC-
1 for 5 min at 37°C. After the reaction mixture
was incubated with PLC-
1 for an additional 10 min at 37°C, the reaction was stopped by adding 500 µl of
chloroform:methanol (1:1), and the lipids were extracted in the
presence of 1 N HCl containing 5 mM EGTA. The aqueous and organic
phases were separated by centrifugation, and a 350-µl portion of the
upper aqueous phase was removed for liquid scintillation counting. The
PLC activity was measured in the presence and absence of recombinant
villin, either phosphorylated or not. The PLC activity was also
measured using native villin and a recombinant protein expressing GST alone.
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RESULTS |
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Recombinant villin demonstrates biological activity like native
villin.
Villin cDNA cloned in pGEX-2T was expressed as a GST fusion protein
(Fig. 1A). The effects of
villin on the polymerization of actin and the kinetics of filament
formation were followed by measuring the change in viscosity of the
protein as described in METHODS. Change in viscosity of
actin was measured using three different molar concentrations of
recombinant villin (recombinant villin:G-actin molar ratios of 1:65,
1:139, 1:270). These concentrations were chosen on the basis of prior
studies of the effects of villin on actin polymerization and are
physiologically meaningful based on the ratio of villin to actin in
purified brush border (9). Viscosity measurements using
native villin (native villin:G-actin molar ratios of 1:139 and 1:270)
were done in parallel. GST (GST:G-actin molar ratio, 1:65) and the
buffer cocktail were used as negative controls. As shown in Fig.
1B, recombinant villin polymerizes actin nearly as well as
the native protein.
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Villin can be tyrosine phosphorylated in vitro.
Native villin (760 ng) and recombinant villin (3 µg) were tyrosine
phosphorylated in vitro using c-src kinase. Unlike the native villin,
recombinant villin was not tyrosine phosphorylated in vitro by c-src
(Fig. 2A). Previous studies on
phosphorylation of gelsolin suggested that addition of PIP2
to the kinase assay significantly increased the tyrosine
phosphorylation of gelsolin in vitro (11). Thus it was
possible that PIP2 might alter the ability of c-src to
phosphorylate recombinant villin in vitro. However, addition of
PIP2 to the kinase assay mixture did not phosphorylate
recombinant villin (data not shown). In contrast, in vitro
phosphorylation of recombinant villin required the presence of the
reducing agent, -mercaptoethanol (at concentrations
0.5 mM; Fig.
2B). Lower concentrations of
-mercaptoethanol (<2 mM) did not induce significant phosphorylation, whereas concentrations >5
mM did not lead to any further increase in the tyrosine phosphorylation of recombinant villin (data not shown). The native villin was stored in
0.5 mM
-mercaptoethanol. However, such a small amount (1 µl) of
the purified protein was used (which was further diluted in the kinase
reaction mixture) for phosphorylation by c-src that the final
concentration of
-mercaptoethanol in the kinase mixture was
insignificant. That the native protein is reduced in vivo by some
physiological equivalent of
-mercaptoethanol cannot be ruled out.
The addition of exogenous
-mercaptoethanol did not significantly
increase the tyrosine phosphorylation of native villin (data not
shown). These results suggest that the recombinant villin may form
disulfide bonds and assume a conformation that masks the
phosphorylation site, and reducing agents such as
-mercaptoethanol may be required to make this accessible for in vitro phosphorylation of
recombinant villin; or alternatively, the recombinant villin may be
folded differently from the native villin. However, it is possible that
the cleavage of disulfide linkages within the villin protein may
actually be important for the tyrosine phosphorylation of villin. A
physiological role for disulfide bonds has been shown in gelsolin and
severin (25, 39).
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Tyrosine phosphorylation of villin in vitro is not affected by
PIP2.
Previous studies with gelsolin suggested that PIP2
augmented the tyrosine phosphorylation of the protein by c-src
(11). To further examine a role for PIP2 in
mediating the tyrosine phosphorylation of villin, we tested the effect
of PIP2 on phosphorylation of villin in the presence of
nonsaturating concentrations of -mercaptoethanol. Recombinant villin
(1 µM) was tyrosine phosphorylated in vitro in the presence of
-mercaptoethanol (1 mM) and varying concentrations of
PIP2 (0-50 µM). As shown in Fig.
3A, the addition of exogenous PIP2 to the in vitro phosphorylation assay had no
significant effect on the tyrosine phosphorylation of villin. Similar
data were obtained using native villin purified from chicken brush border (Fig. 3B). In fact, the addition of PIP2
appears to inhibit the phosphorylation of villin, with significant
(78%, P < 0.05, n = 5) inhibition at
50 µM PIP2 (Fig. 3A). PE also did not increase the phosphorylation of villin and, like PIP2, appears to
inhibit (at 200 µM, 49%, P < 0.05, n = 3) the tyrosine phosphorylation of villin (Fig.
3C). The concentrations of PIP2 and PE used in this study were based on similar studies done with gelsolin
(11). The data indicate that villin differs from gelsolin
in its requirement for exogenous PIP2 for tyrosine
phosphorylation in vitro.
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Stoichiometry of villin phosphorylation.
The amount of phosphate incorporated in villin was determined in an in
vitro kinase assay (as described in METHODS) using 2.6 nmol
of recombinant villin. Villin was phosphorylated in vitro by c-src in a
reaction mixture containing 30 µCi of [-32P]ATP (20 µM ATP, sp act = 3,000 Ci/mmol) and 5 mM
-mercaptoethanol in
a final volume of 50 µl. The phosphorylated recombinant villin was
affinity purified using glutathione Sepharose, and the radioactivity associated with the phosphorylated villin was measured in a liquid scintillation counter. Phosphorylation of GST was negligible and was
subtracted from the phosphorylated villin samples. Using this method,
we obtained 0.61 mol phosphate/mol of villin. These amounts are
comparable to those reported for gelsolin (0.86 mol phosphate/mol gelsolin) (11) and suggest nearly quantitative
phosphorylation, provided only one major site is phosphorylated in
villin. Similar numbers were obtained with native villin (0.70 mol
phosphate/mol of villin).
Tyrosine phosphorylation of villin in TKX1 cells.
The phosphorylation of villin by c-src is shown in these studies to
characterize the phosphorylation of villin by PIP2, to determine the stoichiometry of villin phosphorylation, and to compare
in vitro phosphorylation of villin with other proteins of its family,
including gelsolin. However, since differences in folding between
recombinant and native villin cannot be ruled out, an alternative
approach to obtain tyrosine-phosphorylated recombinant villin was used.
This approach obliterates the need to use either exogenous kinase or
-mercaptoethanol to phosphorylate the recombinant villin.
Full-length villin cDNA was expressed in the TKX1 cells, which carry an
inducible tyrosine kinase gene (see METHODS). Recombinant
villin was purified from the TKX1 cells as a GST-tagged
tyrosine-phosphorylated protein (Fig. 4).
Batch purification was not used to collect tyrosine-phosphorylated
villin from GST Sepharose because the eluted protein contains
phosphorylated bands in addition to villin (see fraction 4,
Fig. 4B). Phosphorylated villin (VILT/WT) collected from
fractions 7-13 was pooled and used in these studies.
VILT/WT polymerized actin like the native protein (data not shown).
Villin phosphorylated in TKX1 cells was used in all the subsequent
experiments.
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Tyrosine-phosphorylated villin does not bind PIP2.
Although direct interaction of villin with PIP2 has not
been demonstrated, two PIP2-binding domains have been
identified in villin (17). This and our previous
observation that tyrosine-phosphorylated villin associates with
PLC-1 led us to speculate that tyrosine phosphorylation
may regulate the ability of villin to bind both the enzyme and the
substrate involved in phosphoinositide signal transduction, namely
PLC-
1 and PIP2. We sought to determine if the ligand-binding properties of villin could be regulated by its
tyrosine phosphorylation. To determine the association of villin with
PIP2, recombinant villin phosphorylated (VILT/WT) or not
(VIL/WT) was incubated in vitro with PIP2. The recombinant proteins incubated with (Fig. 5,
A and B) or without (Fig. 5C) PIP2 were analyzed by Western analysis using
PIP2 monoclonal antibodies.
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Tyrosine-phosphorylated villin binds to the COOH-terminal SH2
domain of PLC-1.
We have previously reported that tyrosine-phosphorylated villin
associates with PLC-
1 in vivo (22).
PLC-
1 contains two SH2 domains and one SH3 domain. SH2
domains from several different proteins, including
PLC-
1, have been shown to bind specifically to
tyrosine-phosphorylated cellular proteins (32). We
hypothesized that tyrosine-phosphorylated villin may associate with the
SH2 domain of PLC-
1. Plasmids containing the SH2 domains
of PLC-
1 were expressed as GST fusion proteins (Fig.
6A). To determine the
association of tyrosine-phosphorylated villin with SH2 domains of
PLC-
1, we exposed rabbit distal ileum to carbachol (1 µM, 30 s), conditions previously shown to lead to tyrosine
phosphorylation of villin. Brush-border membranes from control tissue
were made in parallel as described in METHODS. The
recombinant PLC-
1 proteins immobilized on glutathione
Sepharose beads were incubated with brush-border membrane extracts from
carbachol-treated ileum, the proteins bound to the beads were separated
by SDS-PAGE, and Western blot analysis was done using villin monoclonal
antibodies. As shown in Fig. 6B, villin associates with the
COOH-terminal SH2 domain of PLC-
1 and with the
recombinant protein containing both the COOH- and the
NH2-terminal domains of PLC-
1. The tyrosine phosphorylation of the villin associated with the SH2 domain of PLC-
1 is shown in Fig. 6C.
Tyrosine-phosphorylated villin does not associate with the
NH2-terminal SH2 domain of PLC-
1 (Fig. 6,
B and C). Brush border extracts from control
ileum incubated with PLC-
1 recombinant proteins showed
no association of villin with PLC-
1 (Fig.
6D). Figure 6E shows Western analysis of the same
samples as shown in Fig. 6, B, C, and
D, using GST monoclonal antibodies. The
villin-PLC-
1 complex was also probed for other known
ligands of PLC-
1, including nck, actin, and grb-2, all of which were negative (data not shown). A GST fusion protein expressing the SH3 domain of PLC-
1 showed no association
with either nonphosphorylated or phosphorylated villin (data not
shown).
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Nonphosphorylated villin inhibits PLC-1 catalytic
activity in vitro.
Having shown that villin can associate with both PIP2 and
PLC-
1, we next sought to determine if villin could
regulate the activity of PLC-
1. Purified
PLC-
1 (40 ng) used in these studies was determined to be
98% pure by HPLC as described earlier (2). PLC-
1 catalytic activity was measured in the presence or
absence of different concentrations of recombinant villin (VIL/WT,
0-30 µM). As shown in Fig.
7A, villin inhibits
PLC-
1 activity in a dose-dependent manner
(n = 11). Fifty percent of the PLC-
1
activity was inhibited by villin at a concentration of 12.4 µM. This
concentration dependency is comparable to that observed for the
inhibitory effect of gelsolin on PLC-
activity (34).
The PLC-
1 catalytic activity was also measured in the
presence of native villin, and similar results were obtained. There was
no change in the PLC-
1 activity in the presence of GST.
PLC-
1 activity was measured in the presence of 10 µM
villin and increasing concentrations of PIP2 (Fig.
7B). Increasing concentrations of PIP2 reversed
the inhibitory effect of villin on the PLC-
1 catalytic
activity. The simplest interpretation of these data is that villin
inhibits PLC-
1 by substrate competition.
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Tyrosine-phosphorylated villin does not inhibit
PLC-1 activity in vitro.
To determine the effect of tyrosine-phosphorylated villin on
PLC-
1 catalytic activity, we used recombinant villin
phosphorylated in TKX1 cells (VILT/WT). Nonphosphorylated villin
(VIL/WT) was obtained from TKX1 cells grown in the absence of IAA. The
PLC-
1 activity was measured in vitro in the presence of
villin (5 and 10 µM) that was phosphorylated or not. As seen in Fig.
7C, recombinant villin inhibited PLC-
1
activity at both concentrations. In contrast, phosphorylated villin did
not inhibit PLC-
1 at either concentration used. These
data show that tyrosine phosphorylation abolishes the ability of villin
to inhibit PLC-
1 catalytic activity in vitro. On the
basis of these data, we propose that tyrosine phosphorylation of villin
is fundamental to the regulation of PLC-
1 catalytic activity by villin, because it determines villin's ability to associate with both PIP2 and PLC-
1.
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DISCUSSION |
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Villin, an actin-regulatory protein described in the epithelial
cells of the intestine and kidney, is now being reported in other
epithelial cells such as the merkel cells (37) and the alveolar cells (20), suggesting that villin may be a
protein ubiquitous to all epithelial cells. We have previously shown
that tyrosine phosphorylation of villin and its association with
PLC-1 in the intestinal epithelial cells is involved in
the cytoskeletal rearrangement associated with receptor activation
(21, 22). Since our first report, tyrosine phosphorylation
of villin and its association with PLC-
1 has been shown
to be involved in rearrangement of the actin cytoskeleton in the
opossum kidney cell line (28). Villin is abundantly
expressed in renal proximal tubule and intestinal microvilli, and its
association with PLC-
1 has been shown to regulate ion
transport in both the renal and intestinal epithelial cells. More
recently, a role for villin in renal ischemia has been reported
(40). The actin cytoskeleton of proximal tubule cells
contributes significantly to development of acute, ischemic kidney tubule cell damage (30). Alterations in the actin
cytoskeleton associated with the actin-severing properties of villin
thus appear to be important both in physiology and pathophysiology of
epithelial cells of the intestine and kidney.
Villin belongs to a family of proteins (that includes gelsolin and severin) that share F-actin-severing activity. Several actin-binding proteins, including gelsolin, CapG, fragmin, and profilin, have been shown to be tyrosine phosphorylated in vitro (11). Although these proteins have not been demonstrated to be tyrosine phosphorylated in vivo, tyrosine kinases of the Src family have been shown to regulate the physiological functions of these proteins (6, 26). These observations suggest that tyrosine phosphorylation of this family of proteins may be a general mechanism that may lend new properties to these proteins and add yet another level of regulation that may be recognized by future studies involving the identification of the phosphorylated tyrosine residues and other functional assays.
We now report for the first time the in vitro tyrosine phosphorylation
of villin. Recombinant villin can be tyrosine phosphorylated in vitro
by tyrosine kinases such as elk (Fig. 4), c-src (Fig. 2), and c-yes
(Khurana, unpublished observation). The brush border of intestinal
epithelial cells expresses both c-src and c-yes (23).
Because it is suggested that phosphorylation of gelsolin is regulated
by c-src, it is reasonable to speculate that villin phosphorylation in
vivo may also be regulated by the Src family tyrosine kinases, c-src
and/or c-yes. Although the recombinant villin protein exhibits
actin-regulatory properties similar to those of the native protein
(Fig. 1), it required 2 mM of
-mercaptoethanol for phosphorylation
(Fig. 2B). Tyrosine phosphorylation of gelsolin and CapG in
vitro by c-src is significantly enhanced in the presence of
PIP2 (11). The mechanism of PIP2
stimulation is hypothesized to involve exposure of key tyrosine
residues due to a phospholipid-dependent conformational change in the
proteins. This effect was specific for PIP2 and could not
be duplicated by other anionic and neutral phospholipids, including
phosphatidylserine or PE. In our studies, we could not substitute
-mercaptoethanol with PIP2 or any other phospholipid,
including PE, to tyrosine phosphorylate recombinant villin in vitro.
Also, unlike gelsolin, PIP2 did not enhance the phosphorylation levels of villin in vitro. This observation could mean
one of two things: that villin may actually differ from these proteins
in its requirement for PIP2 for phosphorylation or,
alternatively, that the
-mercaptoethanol present in the kinase
buffer may function like the phospholipid in unmasking the tyrosine
residues. These differences in the phosphorylation of villin and
gelsolin by PIP2 may even be physiologically relevant and
perhaps important to our understanding of the unique function performed
by each of these proteins in vivo. This is important in light of the
fact that both villin and gelsolin are expressed in intestinal
epithelial cells.
Two phosphoinositide binding sites have been identified in villin
(amino acids 111-118 and 137-145), which are also the site of
F-actin binding to villin before severing (18). Like
villin, gelsolin and CapG, members of an actin filament-severing and
capping protein family, are also activated by Ca2+ and
inhibited by phosphoinositides, particularly PIP2
(18). There is emerging evidence that actin-regulatory
proteins binding to PIP2 may have functions beyond a direct
effect on the cytoskeleton. For instance, in vitro gelsolin alters the
activity of phosphoinositide-specific PLC (3, 7),
phospholipase D (4, 33), and phosphoinositide 3-OH kinase
(6, 7). The association of villin with the
phosphoinositide, PIP2, and the phosphoinositide
hydrolyzing enzyme, PLC-1, suggests that villin may be
involved in regulating the phosphoinositide-mediated signal
transduction pathways. These studies suggest that proteins of this
family, including villin, may be components of a signaling complex that
transduces external stimuli to the cortical cytoskeleton.
We have previously shown that tyrosine-phosphorylated villin associates
with PLC-1 in vivo, suggesting that tyrosine
phosphorylation plays a key role in the interaction between villin and
PLC-
1. Using recombinant proteins to the SH2 domain of
PLC-
1, we now show that phosphorylated villin binds
specifically to the COOH-SH2 domain of PLC-
1. Gelsolin,
which shares villin's actin-severing but not actin-bundling
properties, has also been shown to associate in vivo with PLC-
(5). However, the site of interaction between gelsolin and
PLC-
remains to be determined. Future studies with other members of
this family may demonstrate that PLC enzymes may also be common ligands
for this family of proteins.
The association of villin with both the substrate and the enzyme,
namely PIP2 and PLC-1, suggests that
villin's interaction with PLC-
1 may have a functional
consequence. To determine the effect of villin association on
PLC-
1 activity, we measured the catalytic activity of
purified PLC-
1 in the presence or absence of villin.
Villin inhibits PLC-
1 activity in a dose-dependent manner. This is consistent with the idea that proteins of the villin
family may regulate PLC activity, since gelsolin and CapG have been
demonstrated to inhibit PLC-
activity both in vitro and in vivo
(5, 34, 35). The simplest hypothesis to explain this
observation is that since villin binds PIP2, sequestration of the substrate by villin may be the mechanism for the observed inhibition of PLC activity by villin. Such an observation has been made
with gelsolin (34). Reversal of villin's inhibitory effect on PLC-
1 activity in the presence of increasing
concentrations of PIP2 (Fig. 7B) lends support
to this hypothesis. The inability of tyrosine-phosphorylated villin to
inhibit PLC-
1 activity (Fig. 7C) would then
suggest that tyrosine phosphorylation of villin may modulate its
ability to bind PIP2. Indeed, using in vitro binding
studies (Fig. 5, A-C), a centrifugal liposome-based
binding assay (Fig. 5E), as well as association of fusion
proteins to 3H-labeled PIP2 vesicles (Fig.
5D), we demonstrate that, while nonphosphorylated villin
binds PIP2, tyrosine phosphorylation of villin abolishes
its ability to associate with the phospholipid. Our studies demonstrate
that phosphorylation of villin determines its association with
PLC-
1 and PIP2. Thus tyrosine
phosphorylation of villin regulates its interaction with both the
substrate and the enzyme associated with phosphoinositide signal
transduction. The present work presents biochemical proof of the
functional relevance of tyrosine phosphorylation of a cellular protein.
The data presented in this study indicate that there is significant
cross-talk between components of the transmembrane signal machinery and
the actin cytoskeleton at multiple levels, including the generation of
important second messengers. Our working model (Fig.
8) is that binding of villin to
PLC-1 brings the enzyme in proximity of the substrate
(PIP2), thus activating PLC-
1. However, the
enzyme may be activated only when villin no longer sequesters
PIP2, i.e., when villin is tyrosine phosphorylated. The
identified PIP2 binding site in villin is also the site for actin binding to villin before severing (17); thus removal
of PIP2 from this site would make it accessible for actin
to bind to villin and for villin to sever F-actin filaments. Decades of in vitro work with villin have suggested that very high
Ca2+ concentrations (>5 µM) are required to activate
villin's actin-severing function (27). Because this is
not a physiologically relevant intracellular Ca2+
concentration ([Ca2+]i), it has been assumed
for years that villin's actin-bundling and not actin-severing
properties are physiologically significant. Studies done with the
villin-knockout mice report normal bundling of F-actin filaments in the
intestinal microvilli, suggesting that in the absence of villin, the
actin-bundling properties associated with villin can be substituted by
other proteins in the microvilli (28). A second study with
the villin-knockout mice reports that villin is required for F-actin
severing rather than bundling in the microvilli of intestinal cells
(12). On the basis of these observations, it seems it may
be time to revisit villin's actin-severing rather than actin-bundling
functions.
|
According to our model, the regulation of villin's actin-severing
function may be intimately linked to its tyrosine phosphorylation. Tyrosine phosphorylation of villin would lead to the recruitment of
PLC-1 and alter the ability of villin to sequester
PIP2. Removal of PIP2 from villin would vacate
this site for F-actin to bind to this severing site (17).
Activation of PLC-
1 in proximity of villin would
generate D-myo-inositol 1,4,5-trisphosphate
(IP3), which would release Ca2+ from
intracellular stores, leading to a localized increase in [Ca2+]i. Intracellular Ca2+
stores are now identified in the apical membrane of epithelial cells of
the intestine (24). Tyrosine phosphorylation of villin could also modulate its ability to regulate the cortical cytoskeleton in submicromolar concentrations of Ca2+. Concurrent with
this model, we have recently reported a decrease in the F-actin content
as well as a redistribution of villin in response to carbachol
treatment in the intestinal cell line, Caco-2 (21).
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Susan W. Craig (Johns Hopkins University, Baltimore, MD) for assistance with the actin polymerization experiments and the kind gift of villin purified from chicken brush-border membranes, and Dr. Tony Guerrerio (Johns Hopkins University) for assistance with the fluorimetric measurements of actin polymerization.
![]() |
FOOTNOTES |
---|
These studies were supported by a grant from the American Digestive Health Foundation (Industry Research Scholar Award) and by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-54755 to S. Khurana.
Address for reprint requests and other correspondence: S. Khurana, Univ. of Tennessee Health Science Center, Dept. of Physiology, 894 Union Ave., Nash 402, Memphis, TN 38163 (E-mail: skhurana{at}utmem.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 31 January 2001; accepted in final form 3 May 2001.
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