(Received for publication, April 24, 1997, and in revised form, May 22, 1997)
From the Department of Physiology, University of
Texas Southwestern Medical Center, Dallas, Texas and the
§ Division of Medicinal Chemistry and Pharmaceutics, College
of Pharmacy, University of Kentucky,
Lexington, Kentucky 40536-0082
The actin cytoskeleton of nonmuscle cells undergoes extensive remodeling during agonist stimulation. Lamellipodial extension is initiated by uncapping of actin nuclei at the cortical cytoplasm to allow filament elongation. Many actin filament capping proteins are regulated by phosphatidylinositol 4,5-bisphosphate (PIP2), which is hydrolyzed by phospholipase C. It is hypothesized that PIP2 dissociates capping proteins from filament ends to promote actin assembly. However, since actin polymerization often occurs at a time when PIP2 concentration is decreased rather than increased, capping protein interactions with PIP2 may not be regulated solely by the bulk PIP2 concentration. We present evidence that PIP2 binding to the gelsolin family of capping proteins is enhanced by Ca2+. Binding was examined by equilibrium and nonequilibrium gel filtration and by monitoring intrinsic tryptophan fluorescence. Gelsolin and CapG affinity for PIP2 were increased 8- and 4-fold, respectively, by µM Ca2+, and the Ca2+ requirement was reduced by lowering the pH from 7.5 to 7.0. Studies with the NH2- and COOH-terminal halves of gelsolin showed that PIP2 binding occurred primarily at the NH2-terminal half, and Ca2+ exposed its PIP2 binding sites through a change in the COOH-terminal half. Mild acidification promotes PIP2 binding by directly affecting the NH2-terminal sites. Our findings can explain increased PIP2-induced uncapping even as the PIP2 concentration drops during cell activation. The change in gelsolin family PIP2 binding affinity during cell activation can impact divergent PIP2-dependent processes by altering PIP2 availability. Cross-talk between these proteins provides a multilayered mechanism for positive and negative modulation of signal transduction from the plasma membrane to the cytoskeleton.
Phosphoinositides are important in signal transduction, both as
precursors to signaling molecules and as physical anchors and
regulators of proteins (1, 2). Among these, the D4 phosphoinositide, phosphatidylinositol 4,5-bisphosphate
(PIP2),1 has been implicated as
a potential mediator of actin cytoskeletal rearrangements (3, 4).
PIP2 modulates many actin regulatory proteins. These
include the following: actin severing and/or capping proteins (gelsolin
(5), CapG (6), and capping protein (also known as Cap Z) (7)),
monomer-binding proteins (profilin (8) and cofilin (9)), and other
actin-binding proteins (-actinin (10) and vinculin (11)). It has
been hypothesized that PIP2 induces explosive actin
assembly by dissociating capping proteins from filament ends and
releasing actin monomers from actin-sequestering proteins (3, 7, 12).
The involvement of PIP2 in actin polymerization is
supported by recent experiments that show that Rac1 and RhoA, monomeric
GTPases of the Rho family that have well defined effects on the
cytoskeleton (13), stimulate the synthesis of PIP2
(14-16). Furthermore, manipulations that alter the availability of
PIP2 in cells have profound effects on agonist and/or
Rac1-induced filament end capping, actin polymerization, and cell
motility (16, 17). However, although the time courses of
PIP2 hydrolysis and recovery correlate in some cells (16,
18), they do not in most of the cells examined (19-21). Particularly
puzzling is the finding that, in many cells, actin polymerizes at a
time when PIP2 level is reduced, rather than increased, as
would be expected if uncapping and monomer desequestration are
initiated by PIP2. To explain this discrepancy, it is often
hypothesized that local PIP2 availability can be enhanced
by compartmentalization or differential turnover (22-24), even as the
bulk PIP2 mass is reduced. The equally attractive
possibility that PIP2 binding is regulated by signals generated during agonist stimulation has not been considered.
Agonist-stimulated cells exhibit complex Ca2+ oscillations and pH transients. These signals alter the binding of gelsolin and CapG to actin, by inducing a conformational change (6, 25-27). In this study, we tested the effect of Ca2+ and pH on the binding of the gelsolin family proteins to PIP2 and found that they affect PIP2 binding in an interdependent manner. We identified the domains in gelsolin that impart such regulation and elucidated the relation between the NH2-terminal and COOH-terminal halves of the protein. Since gelsolin modulates the activity of many PIP2-regulated proteins with important signaling functions in vivo (28) and in vitro (29-31), our results have important implications for how the gelsolin family proteins are regulated during agonist signaling and how the activity of other PIP2-dependent cytoskeletal and noncytoskeletal proteins can be coordinated.
Gelsolin has six semihomologous domains (S1-6), which can be further divided into two functional halves (32). The expression vectors for the gelsolin NH2-terminal half (S1-3), gelsolin S1, gelsolin S2-3, and CapG have been described previously (33-35). The full-length gelsolin expression vector (encompassing the entire human plasma gelsolin coding sequence) was constructed by ligating gelsolin cDNA to pet3a via the BamHI site. Recombinant proteins were expressed in bacteria and purified using sequential anion and cation exchange chromatography (34). Protein concentration was determined by the method of Bradford (36), and protein purity was assessed by SDS-polyacrylamide gel electrophoresis.
The COOH-terminal half expression vector was constructed by using
polymerase chain reaction to generate a fragment encompassing human
plasma gelsolin nucleotides 1298-1753. The forward primer contains a
XhoI site (ACC TCC ACT CTC GAG GCC GCC), and the reverse primer has a SmaI site (CAA CAG CCC GGG TGG CT). The
polymerase chain reaction product was cloned into Bluescript KS+ via
the XhoI/SmaI sites. This construct was digested
with SmaI and blunt end-ligated with a downstream gelsolin
fragment. The fragment was excised with BamHI from
full-length gelsolin cDNA in Bluescript KS+ (gelsolin
SmaI site at nucleotide 1750 and vector multiple cloning
SmaI site downstream of the termination codon). The
resultant cDNA was digested with SpeI (in the 3
multiple cloning region, downstream of SmaI) and filled in
with CT nucleotides to create a site with a two-base overhang
compatible with that of HindIII. The other end was released
by digestion with XhoI and ligated to PGEX K6 vector that
was linearized with HindIII (site partially filled in with
nucleotides AG to generate a two-base overhang compatible with the
partially filled in SpeI) and XhoI. The fusion protein contained a 30-kDa GST followed by a 40-kDa gelsolin
COOH-terminal half. The COOH-terminal gelsolin was cleaved from GST
bound to a column with thrombin.
PIP2 was purchased from Calbiochem. Micelles were prepared by dissolving the dried lipid in water to a final concentration of 2 mg/ml and sonicating for 5 min. at maximum power (model W185; Heat Systems Ultrasonics, Inc., Farmingdale, NY). Large unilamellar vesicles at a 5:1 phosphatidylcholine:PIP2 ratio were made with an extruder (Lipex Biomembranes, Vancouver, Canada) as described by Machesky et al. (37).
Small Zone Gel FiltrationThe assay was similar to that
described previously for studying lipid binding to most actin
regulatory proteins (33, 35, 38). This is because small proteins bound
to PIP2 micelles or mixed vesicles migrate faster than the
unbound proteins. Proteins were incubated with lipid for 30 min at room
temperature, and 100 µl of the mixture was chromatographed at 4 °C
through a Superdex 75 HR 10/30 column (Pharmacia Biotech Inc.),
equilibrated with pH 7.0 or 7.5 buffers containing 25 mM
Hepes, 100 mM KCl, 0.5 mM -mercaptoethanol,
0.4 mM EGTA with or without CaCl2. Lipid was
not included in the elution buffer. Fractions were eluted at 0.5 ml/min, and 0.5-ml fractions were collected. The elution profile was
monitored by absorbance at 280 nm. The amount of unbound protein was
determined from the protein absorbance peak. The lipid-bound protein
was calculated as the difference between the total protein applied
minus the unbound protein. The apparent dissociation constant (Kd) was calculated as follows.
![]() |
(Eq. 1) |
The method of Hummel and Dreyer (39), as modified by Machesky et al. (37) was used. A Superose 12 HR 10/30 column (Pharmacia) was equilibrated with CapG (ligand) in a buffer containing 25 mM Hepes, 75 mM KCl, 0.5 mM dithiothreitol, 1.8 mM NaN3, 0.05 µM CaCl2, pH 7.5, at room temperature. 100 µl of the equilibration buffer containing CapG was incubated with PIP2 micelles for 30 min and loaded onto the column. The column was developed with the equilibration buffer containing CapG at 0.25 ml/min, and 0.3-ml fractions were collected. CapG concentration in the column fractions was monitored by UV absorption. The amount of CapG bound to PIP2 was determined from the trough in the absorbance peak. Multiple runs using equilibration CapG concentrations of 0.96, 1.3, 2.6, and 3.9 µM and PIP2 concentrations of 34, 46, 68, and 91 µM were done. Kd was determined by the equation,
![]() |
(Eq. 2) |
Fluorescence
spectra were recorded at 30 °C with a QM-1 fluorometer (Photon
Technology International, Canada). 2 ml of a protein solution (0.3 µM, 30 °C) in 25 mM Hepes, 100 mM KCl, 0.4 mM EGTA, 0.5 mM
-mercaptoethanol, pH 7.5, with or without 36 µM free
Ca2+ were placed in a 1-cm square quartz cuvette and
stirred with a minimagnetic stirrer. After allowing 5 min for
equilibration, the tryptophan fluorescence spectrum was recorded by
excitation at 292 nm. The excitation and emission beam slits were set
at 3 and 2 nm bandwidth, respectively. PIP2 micelles (at
final PIP2 concentrations ranging from 0.042 to 32.3 µM, depending on the protein studied) were added at
2-µl increments, and the fluorescence spectra were recorded 5 min
after each addition. The total volume of micelles added did not exceed
2% of the initial protein solution volume. The decrease in
fluorescence emission at 320 nm was plotted as a function of
PIP2 concentration, and the fluorescence change was assumed
to be proportional to the concentration of the protein-phosphoinositide complex. Data were analyzed as described by Ward (40). The apparent dissociation constant, Kd, was calculated using the
equation,
![]() |
(Eq. 3) |
![]() |
(Eq. 4) |
The concentrations of free Ca2+ in EGTA containing solutions with varying amounts of Ca2+ were measured with Ca2+-sensitive dyes. 5 µM Fura-2 was used to determine Ca2+ concentrations below 1 µM. Free Ca2+ concentration was calculated (26) assuming the Kd of the Fura-2-Ca2+ complex is 229 nM at pH 7.0 and 144 nM at pH 7.5. Calcium green 5N (Molecular Probes, Eugene, OR) was used to measure Ca2+ concentrations higher than 1 µM, and free Ca2+ concentration was calculated assuming a Kd of 14 µM.
Small zone gel filtration
analyses showed that CapG bound to PIP2 micelles in a
dose-dependent manner. Micelle-bound CapG eluted in the
void volume that was well separated from the free protein peak (Fig.
1). Binding to phosphatidylcholine-PIP2
vesicles gave similar results (data not shown), suggesting that
micelles could be used to assess binding, although it is not a
physiological substrate. To facilitate comparison under different
binding conditions and between different proteins, we attempted to
calculate a Kd. Equilibrium binding studies suggest
that each CapG binds two PIP2 molecules (see below).
Assuming this stoichiometry, the apparent Kd for
binding to PIP2 micelles (calculated using Equation 1) was
69.0 µM in 1 mM EGTA, and 29.4 µM in the presence of 36 µM
Ca2+ at pH 7.5 (Table I). These values
represent the upper limit, since measurements were not made under
equilibrium conditions.
|
To determine if there is indeed a Ca2+-induced change,
equilibrium binding studies based on the quenching of CapG intrinsic tryptophan fluorescence by PIP2 were performed. This method
has been used to study the binding of profilin (43), phospholipase C
(44), and dynamin pleckstrin homology domain (45) to PIP2. CapG had an emission maximum of 327 nm, and 36 µM
Ca2+ produced a small reduction in fluorescence intensity
(the ratio of peak fluorescence in EGTA/Ca2+ is 0.92 ± 0.05 (mean ± S.E., n = 5) (Fig.
2, A and B). PIP2
induced a dose-dependent and saturable decrease in
intrinsic fluorescence, without shifting the emission maximum. Micelles
alone without CapG did not have significant emission (data not shown).
A plot of CapG fluorescence quenching versus
PIP2 concentration showed that saturation was reached at a
lower PIP2 concentration in the presence of
Ca2+ than in EGTA (Fig. 3A). The
Kd values for binding at pH 7.5, calculated
according to Equation 3, were 31.9 and 8.4 µM in EGTA and
Ca2+, respectively, for the experiment shown. Similar
values (24.4 and 6.0 µM) were obtained when the data were
analyzed using Equation 4 (Table I). These Kd values
were 3-4 times lower than the small zone gel filtration values,
suggesting that CapG-PIP2 complexes dissociate during
nonequilibrium gel filtration. Using a similar protocol, human platelet
profilin binds PIP2 with a Kd of 35 µM (43), and binding is not affected by
Ca2+.
The stoichiometry of CapG binding was 1.7 in either Ca2+ or EGTA (Table I). Since CapG has one known PIP2 binding site (6, 33), this site appears to bind two PIP2 molecules. The two PIP2 bound independently and noncooperatively, as indicated by the Hill coefficients of close to 1 (1.02 ± 0.05 and 1.09 ± 0.02 in EGTA and Ca2+, respectively) (Fig. 3B, Table I). The exact meaning of this stoichiometry is not clear, because each micelle contains multiple PIP2 and CapG can potentially bind more than one micelle. Nevertheless, the calculated stoichiometry is useful for comparison among different proteins.
Equilibrium gel filtration validated the Kd derived
by fluorescence titration. The column was preequilibrated with CapG,
and PIP2 incubated with CapG in the equilibrating buffer was added. The column was then developed with CapG containing equilibration buffer. CapG bound to PIP2 migrated faster,
increasing the CapG content above the equilibration level (peak) and
depleting the amount in the trailing fractions (trough) (Fig.
4, A and B). Assuming that each
CapG bound two PIP2 molecules (see Table I), the
Kd obtained from five experiments performed with a
range of CapG and/or PIP2 concentration was 8.1 ± 0.9 µM (mean ± S.E.). This is comparable with the
spectroscopic titration result, affirming the validity of the two
independent methods.
Gelsolin Binding to PIP2
Tryptophan titration
could not be used to study gelsolin binding to PIP2 because
the full-length gelsolin signal (without phosphoinositide) fluctuated
and did not reach a steady level even after 20 min. The reason for this
instability was not investigated further. Gel filtration experiments
showed that gelsolin binding to PIP2 was enhanced by
Ca2+ (Fig. 5, A-F). At pH 7.5, the apparent Kd values were 305.4 and 40.2 µM with and without Ca2+ (Table
II). The latter value is similar to that of CapG,
indicating that gelsolin and CapG have comparable PIP2
binding affinity in the presence of Ca2+. However, in EGTA,
gelsolin has a much higher Kd than CapG, suggesting
that Ca2+ induces a larger change in binding affinity. This
could be due to a disproportionate increase in
koff relative to kon. 10 µM Mg2+ did not substitute for
Ca2+ (data not shown), consistent with previous results
(46).
|
The effect of Ca2+ was amplified when the pH was shifted from 7.5 to 7.0 (Fig. 5, compare A-C with D-F). The relations among Kd, Ca2+, and pH are shown in Fig. 5G. In the absence of Ca2+, decreasing pH from 7.5 to 7.0 had minimal effect (Kd of 300 and 350 µM, respectively). This is not surprising, since PIP2 protonation is not expected to change substantially within this narrow pH range (47) and a broader pH range does not affect binding of profilin to PIP2 either (37). However, at pH 7.0, less Ca2+ was required to increase binding. 0.2 µM Ca2+ decreased the Kd by half at pH 7.0, while 4.5 µM Ca2+ was required to produce the same effect at pH 7.5. Both Ca2+ concentrations are well within the range achieved following agonist stimulation, particularly at the cytoplasm immediately subjacent to the plasma membrane.
Ca2+ and pH Regulation of Gelsolin DomainsTo determine which part of gelsolin contributes to the Ca2+ and/or pH dependence of PIP2 binding, we examined the PIP2-binding characteristics of several gelsolin domains. Gelsolin contains six segmental repeats, S1-6 (32). The NH2-terminal half encompassing S1-3 binds actin independently of Ca2+ (48) and has two known PIP2 binding sites and potentially a third unmapped site (33, 49, 50). The COOH-terminal half (S4-6), which requires Ca2+ to bind actin (51), has not been examined previously for PIP2 binding.
Unlike full-length gelsolin, the gelsolin NH2-terminal half
behaved well during fluorescence titration (Fig.
6A). It bound PIP2 with high
affinity, and saturation was reached at a slightly lower
PIP2 concentration in EGTA than in Ca2+ (the
opposite of full-length gelsolin and CapG). The Kd values for the experiment shown in Fig. 6A were 1.2 and 2.9 µM, respectively. The stoichiometry of binding, derived
from Fig. 6B, was 3.4. This value is twice that of CapG,
confirming that gelsolin NH2-terminal half has more
PIP2 binding sites (33). Gel filtration studies confirmed
that Ca2+ increased the Kd. The Hill
coefficient of 1.1 ± 0.03 (Fig. 6C, Table II)
suggested that binding was noncooperative and that the sites bound
PIP2-independently. S1, which has one PIP2
site, bound 1.6 mol of PIP2 with a Kd of
4.2 µM in EGTA, while S2-3 bound 2.1 µmol of
PIP2 with a Kd of 1.0 and 2.9 µM in EGTA and Ca2+, respectively (Table
II).
The gelsolin COOH-terminal half bound PIP2 with much lower
affinity (approximately 7-fold higher Kd by
fluorescence measurements) than the NH2-terminal half
(Table II). It is therefore probably not involved in PIP2
binding per se. As with the NH2-terminal half,
binding to the COOH-terminal half was reduced in Ca2+ (Fig.
7C). This is in sharp contrast to the large
Ca2+-enhancement of PIP2 binding to full-length
gelsolin. The opposite effects of Ca2+ on full-length and
half-length gelsolins therefore cannot simply be due to nonspecific
lipid aggregation. The pronounced enhancement of PIP2
binding to full-length gelsolin most likely reflects a Ca2+-dependent exposure of the
NH2-terminal half PIP2 binding sites through a
change in the COOH-terminal half. This conclusion is based on the
observation that neither the NH2- nor COOH-terminal halves
are activated by Ca2+ to bind PIP2, and only
the COOH-terminal half is known to undergo Ca2+-induced
conformational change (51).
Gelsolin NH2-terminal half binding to PIP2 was enhanced by lowering pH. The Kd dropped from 8.2 to 3.4 µM between pH 7.5 and 7.0 in the presence of EGTA (Fig. 7A). In contrast, the gelsolin COOH-terminal half was not affected by pH (Fig. 7B).
Actin polymerization in response to agonist activation is frequently associated with a rise in cytosolic Ca2+, changes in PIP2 content, and intracellular pH. There is also compelling evidence that gelsolin, which severs and caps actin filaments in response to changes in Ca2+ and PIP2 concentration and pH, is involved in actin remodeling (17, 52-54). In this paper, we show that gelsolin and CapG binding to PIP2 is affected by physiologically relevant changes in Ca2+ and pH. The effects are not due to alterations in PIP2 structure per se but reflect changes in the proteins. This is the first report that PIP2 binding to any protein is directly modulated by signals generated during agonist stimulation and has implications for divergent PIP2-dependent processes beyond a direct effect on the cytoskeleton.
The finding that gelsolin binding to PIP2 is promoted by Ca2+ is consistent with the current model for how gelsolin is activated by Ca2+ to bind actin (48, 51). Our deletion studies suggest that the extreme COOH terminus of gelsolin is critical to the inhibition of the NH2-terminal actin binding sites, because gelsolin lacking the COOH-terminal 23 residues no longer requires Ca2+ to bind actin (56). We do not know at present whether actin binding and PIP2 binding are regulated identically. This question can now be addressed, because the actin and PIP2-binding sites of gelsolin have been mapped (33, 50, 56-58) and the crystal structures of gelsolin S1 complexed with actin (57) and full-length gelsolin in EGTA2 have been solved recently.
Less is known about how pH affects gelsolin conformation. Selve and Wegner (59) first reported that pH 6 increases the rate of gelsolin binding to actin in the presence of Ca2+. Lamb et al. (26) subsequently showed that the Ca2+ requirement for gelsolin severing is reduced at pH 6.5 and abolished at pH below 6.0. pH 5 induces gelsolin unfolding, as determined by dynamic light scattering (26). We find that a less extreme pH drop potentiates Ca2+ activation of PIP2 binding to full-length gelsolin. Acidic pH increases the NH2-terminal half binding to PIP2 even without Ca2+ but has no effect on COOH-terminal half binding. Therefore, mild acidification probably promotes PIP2 binding by directly altering the NH2-terminal PIP2 binding sites.
The significance of an increase in PIP2 affinity described here depends on the PIP2 concentration in the plasma membrane. This is difficult to estimate precisely because PIP2 may be partitioned and sequestered. One estimate, based on PIP2 accounting for 1% of plasma membrane lipid, suggests that the PIP2 concentration in the plasma membrane of a spherical cell with a radius of 10 µm is 10 µM (44). In platelets, the PIP2 concentration is estimated to be about 300 µM when averaged over the entire cell volume (internal and plasma membranes) (60), and PIP2 concentration decreases by 30% following stimulation (16). Cytosolic [Ca2+] rises during agonist stimulation, and the 4-8-fold increase in CapG and gelsolin binding affinity described here is sufficiently large to promote their increased association with the plasma membrane despite a modest decrease in membrane PIP2. The magnitude of the increase depends on the PIP2 concentration before and after stimulation. Immunogold labeling studies show that 4 and 6.5% of gelsolin is associated with the plasma membrane in resting and activated platelets, respectively (42). This represents a 63% increase in membrane association after stimulation. Our finding that Ca2+ increases PIP2 binding affinity can explain how PIP2 uncaps gelsolin and CapG even as the plasma membrane PIP2 content decreases following agonist stimulation.
Since only a handful of the currently identified
PIP2-binding proteins are Ca2+- and
pH-sensitive, our finding is consistent with a selective regulation of
the gelsolin family. Nevertheless, increased gelsolin and CapG binding
will impact multiple PIP2-dependent processes by altering PIP2 availability to other binding proteins,
especially when PIP2 concentration is decreased during
agonist stimulation. Some actin-binding proteins are inhibited by
PIP2 (profilin, cofilin, capping protein), while others are
activated (-actinin and vinculin). Gelsolin and CapG can therefore
exert positive as well as negative effects indirectly by controlling
PIP2. We postulate that as the cytosolic
[Ca2+] rises during stimulation, gelsolin severs
filaments and PIP2 dissociates it from the filament end.
Increased gelsolin binding to PIP2 displaces capping
protein and profilin, neither of which are Ca2+-sensitive,
from the plasma membrane. Profilin catalyzes polymerization (16), and
the reaction is terminated by capping protein-mediated filament capping
(55). Multiple rounds of severing, uncapping, and facilitated actin
addition at the barbed ends fuel explosive amplification of filament
growth observed during lamellipodial extension and membrane
ruffling.
Our findings also have implications beyond a direct effect on the
cytoskeleton. Many important signaling proteins are regulated by
PIP2 as well. It is significant that several pleckstrin
homology proteins (reviewed in Ref. 2) bind PIP2 with
similar affinity as the gelsolin family. For example, the
Kd values of -adrenergic receptor kinase type 1, pleckstrin, dynamin, and phospholipase C
are 50, 50, 4, and 1 µM, respectively. Therefore, gelsolin and CapG can
potentially compete with them for PIP2, particularly when
the [Ca2+] rises and PIP2 level drops during
agonist stimulation. This possibility is supported by in
vitro and in vivo experiments. In vitro,
gelsolin stimulates and inhibits inositol-specific phospholipase C
isozymes in a biphasic manner (29).3
Gelsolin stimulates phosphoinositide 3-OH-kinase (31), although we find
that gelsolin and CapG also inhibit it.4
Gelsolin activates phospholipase D (30) in a
PIP2-dependent manner. Modest overexpression of
CapG (28) or gelsolin has profound effects on phospholipase C
and
phospholipase C
activated through two distinct receptor-mediated
pathways.3
In conclusion, these observations show that gelsolin and CapG binding to PIP2 is selectively regulated by second messengers. This regulation provides an additional level of control above that of a bulk change in PIP2 content. Differential modulation and cross-talk between the PIP2-binding proteins allow control to be exerted at multiple points in the signaling cascade.
We thank Drs. J. Albanesi, D. Hilgemann, and P. Thomas for helpful discussions and L. Segura for excellent technical assistance.