From the Laboratory of Molecular and Cellular
Neuroscience, The Rockefeller University, New York, New York 10021, the
§ Division of Chemistry and Chemical Engineering, California
Institute of Technology, Pasadena, California 91125, the
¶ Department of Experimental Medicine, Section of Human
Physiology, University of Genova, Genova I-16132, Italy, and the
Department of Psychiatry, School of Medicine, Yale University,
New Haven, Connecticut 06508
Received for publication, June 10, 2002, and in revised form, October 31, 2002
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ABSTRACT |
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Spinophilin is a protein phosphatase 1 (PP1)- and
actin-binding protein that modulates excitatory synaptic transmission
and dendritic spine morphology. We report that spinophilin is
phosphorylated in vitro by protein kinase A (PKA).
Phosphorylation of spinophilin was stimulated by treatment of
neostriatal neurons with a dopamine D1 receptor agonist or with
forskolin, consistent with spinophilin being a substrate for PKA in
intact cells. Using tryptic phosphopeptide mapping, site-directed
mutagenesis, and microsequencing analysis, we identified two major
sites of phosphorylation, Ser-94 and Ser-177, that are located within
the actin-binding domain of spinophilin. Phosphorylation of spinophilin
by PKA modulated the association between spinophilin and the actin
cytoskeleton. Following subcellular fractionation, unphosphorylated
spinophilin was enriched in the postsynaptic density, whereas a pool of
phosphorylated spinophilin was found in the cytosol. F-actin
co-sedimentation and overlay analysis revealed that phosphorylation of
spinophilin reduced the stoichiometry of the spinophilin-actin
interaction. In contrast, the ability of spinophilin to bind to PP1
remained unchanged. Taken together, our studies suggest that
phosphorylation of spinophilin by PKA modulates the anchoring of
the spinophilin-PP1 complex within dendritic spines, thereby likely
contributing to the efficacy and plasticity of synaptic transmission.
Dendritic spines are specialized protrusions that receive the
majority of excitatory input in the central nervous system (1-3). Spines are highly motile and have been observed to change shape rapidly
in response to changes in behavior, hormonal status, and synaptic
activity (4-12). This dynamic behavior is believed to be fundamental
to the function of dendritic spines and to contribute to the efficacy
and plasticity of synaptic transmission. The ability of dendritic
spines to change shape has been attributed to a dense network of
proteins that facilitates the rapid assembly and disassembly of the
actin cytoskeleton (13, 14). Indeed, dendritic spines contain a rich
variety of receptors, ion channels, actin-regulating proteins, and
other biochemical machinery that support the organization of the actin
cytoskeleton (15, 16). Despite growing evidence that this protein
network may be involved in the dynamic behavior of spines, the
molecular mechanisms that underlie this process are not well understood.
Spinophilin (also known as neurabin II) is a
PP11- and actin-binding
protein that is enriched in dendritic spines (17, 18). Several
independent lines of evidence suggest that spinophilin may link
synaptic transmission to changes in the structure and function of
dendritic spines. Spinophilin has been shown to regulate excitatory
synaptic transmission by targeting PP1 to
Spinophilin is likely to influence the dynamic behavior of dendritic
spines by its ability to modulate the actin cytoskeleton. Spinophilin
has been shown to bind and cross-link actin filaments in
vitro (18). In vivo, spinophilin-deficient mice
exhibited a marked increase in spine density during development (19). Moreover, cultured neurons from spinophilin knockout mice had more
filopodia, or spine-like protrusions, but the same number of nerve
terminals as wild-type mice. These observations suggest that
spinophilin may either facilitate spine retraction or suppress the
initial outgrowth of spines from the dendrite.
In addition to actin, spinophilin interacts with a variety of other
proteins, including its homologue neurabin (21, 22), D2-class dopamine
receptors (23), Materials--
Male Sprague-Dawley rats (150-200 g) were
obtained from Charles River Laboratories (Wilmington, MA). Drugs were
obtained from the following sources: cyclosporin A, okadaic acid, and
calyculin A from Alexis Biochemicals (San Diego, CA); forskolin,
SKF-81297 and quinpirole from Sigma-Aldrich (St. Louis, MO). The
purified catalytic subunit of PP1 was kindly provided by Dr. Hsien-bin Huang (National Chung Cheng University, Taiwan). Radioisotopes were
purchased from PerkinElmer Life Sciences (Boston, MA), and cellulose
thin-layer chromatography plates were from Kodak (Rochester, NY).
Restriction enzymes were purchased from Invitrogen (Rockville, MD) and
polyvinylidene difluoride (PVDF) membrane was from Millipore (Bedford,
MA). Oligonucleotides were obtained from Operon Technologies (Berkeley,
CA), and peptides and phosphopeptides were synthesized at The
Rockefeller University Protein/DNA Technology Center (New York, NY).
Preparation and 32P Labeling of Neostriatal
Slices--
Neostriatal slices were prepared from male Sprague-Dawley
rats (8-12 weeks old) as described previously (26). Coronal sections (500 µm in thickness) were cut on a Vibratome (Ted Pella, Redding, CA), maintained at 4 °C, and placed in cold, oxygenated (95%
O2/5% CO2)
Krebs-HCO Immunoprecipitation of 32P-Labeled
Spinophilin--
Frozen tissue slices were sonicated in lysis buffer
(10 mM Na2HPO4, pH 7.0, 150 mM NaCl, 5 mM EDTA, 1% Triton X-100, and 0.2% SDS) containing protease inhibitors (1 mM
phenylmethylsulfonyl fluoride, 20 µg/ml leupeptin, 20 µg/ml
antipain, 5 µg/ml pepstatin, 5 µg/ml chymostatin, 1 mM
benzamidine) and phosphatase inhibitors (20 mM NaF, 1 mM Na3VO4, 0.05 mM
Na2MoO4). The total
[32P]phosphate incorporated into trichloroacetic
acid-precipitated protein was determined for each sample, and
homogenates containing equal amounts of 32P-labeled
proteins were mixed with pre-swollen protein A-Sepharose CL-4B (10 µl
per sample, Amersham Biosciences, Piscataway, NJ) for 1 h at
4 °C. The beads were then pelleted by centrifugation, and the
supernatant was transferred to tubes containing a rabbit polyclonal
antibody against spinophilin (3 µg of RU145 per sample (17)). The
sample was mixed at 4 °C overnight and then for an additional 2 h with pre-swollen protein A-Sepharose. The beads were pelleted by
centrifugation and washed four times with lysis buffer and once with 50 mM HEPES, pH 7.0. After the final wash, the beads were
resuspended in SDS-PAGE sample buffer (50 mM Tris-HCl, pH
6.7, 10% glycerol, 2% SDS, 10% 2-mercaptoethanol, and 0.01% bromphenol blue), boiled for 5 min, and centrifuged. The recovered proteins were resolved by SDS-PAGE on 8% acrylamide gels. The gels
were dried, and [32P]phosphate incorporation into
spinophilin was visualized using a PhosphorImager 400B and ImageQuaNT
software from Amersham Biosciences.
Cloning and Expression of Spinophilin Fusion Proteins--
A
spinophilin construct containing an amino-terminal histidine tag was
prepared as follows: the spinophilin sequence encoding residues 1-305
was amplified by PCR using the primers 5'-CCC ACA TAT GAT GAA GAC GGA
GCC TCG-3' and 5'-CTT TCC TCA ACC TCC ACC GGT T-3'. The resulting
900-bp fragment was digested with NdeI/XhoI and
subcloned with a XhoI/XhoI fragment of
spinophilin (residues 220-817 (17)) into the
NdeI/XhoI site of the vector pET-15b (Novagen,
Madison, WI). Point mutations were introduced using the QuikChange
site-directed mutagenesis kit (Stratagene, La Jolla, CA) and the
spinophilin histidine tag construct as a template. The actin-binding
construct of spinophilin encoding residues 1-221 was amplified by PCR
using the primers 5'-CCC ACA TAT GAT GAA GAC GGA GCC TCG-3' and 5'-CGC
GGA TCC TAC CTC GAG TCG GCT TTC TCG A-3'. The resulting fragment was
digested with NdeI/BamHI and ligated in-frame
into the NdeI/BamHI site of the vector pET-15b. All mutants were confirmed by sequencing. Recombinant proteins were
expressed in bacteria and purified using nickel-nitrilotriacetic acid
agarose resin (Ni-NTA; Qiagen, Valencia, CA) as described previously
(27).
In Vitro Phosphorylation Reactions--
Phosphorylation
reactions were performed using the protein of interest (10 µM) and the catalytic subunit of PKA (40 µg/ml (28)) in
50 mM HEPES, pH 7.4, 10 mM MgCl2, 1 mM EGTA at 30 °C. Reactions were initiated by the
addition of ATP (50 µM) in the presence of
[ Phosphopeptide Mapping--
After autoradiography, gel pieces
containing 32P-labeled spinophilin were re-swollen in
destain (50% methanol/10% acetic acid in water), washed twice with
50% methanol in water, and dried. Gel pieces were then incubated with
L-1-tosylamido-2-phenylethyl chloromethyl ketone-treated
trypsin (50 µg/ml, Worthington, Lakewood, NJ) in 50 mM
NH4HCO3, pH 8.0 (1 ml), for 18 h at
37 °C. The supernatants containing the soluble phosphopeptides were
recovered after centrifugation. The extraction efficiency (~85%) was
quantified by Cerenkov counting of the gel pieces and supernatants
before and after digestion. Two-dimensional phosphopeptide mapping was
performed as described previously (29). For phosphopeptide mapping,
electrophoretic separation was at pH 3.5 for 90 min at 400 V, and
ascending chromatography was in pyridine/n-butanol/acetic
acid/water (15:10:3:12). The pattern of tryptic phosphopeptides was
detected by autoradiography using PerkinElmer Life Sciences (Boston,
MA) or Kodak (Rochester, NY) film.
Identification of the Phosphorylation Sites on
Spinophilin--
Wild-type and alanine mutant histidine fusion
proteins were expressed in BL21(DE3) Escherichia coli by the
induction of log-phase cells (5-ml cultures) with
isopropyl-1-thio-
Phosphorylation sites, determined by point mutations, were confirmed by
reverse phase capillary high performance liquid chromatography (HPLC)
and microsequencing analysis. Wild-type spinophilin (10 µg) was
phosphorylated by PKA in the presence of [ Generation of Phosphorylation State-specific
Antibodies--
Rabbit polyclonal phosphorylation state-specific
antibodies were generated and purified essentially as described (31).
Antibodies were raised against the following cysteine-containing
phosphopeptides conjugated to thyroglobulin: VRL(pS)LPRAC (residues
91-98), CLPRAS(pS)LNE (residues 95-103), and CQERA(pS)LQDRK (residues
173-182). Affinity purification was performed using dephospho- and
phosphopeptides coupled to SulfoLink gel (Pierce, Rockford, IL).
Treatment of Neostriatal Slices and
Immunoblotting--
Neostriatal slices were dissected as described
above and preincubated in fresh
Krebs-HCO Phosphorylation of Spinophilin in HEK 293T Cells and Cultured
Neurons--
Embryonic day 17 rat cerebrocortical or striatal tissue
was used to prepare primary cortical and striatal cultures as described (32, 33). Full-length spinophilin was transiently expressed in 293T
cells for 36 h following calcium phosphate transfection. Cells
were treated with forskolin (50 µM) for 5 min and lysed in 1% SDS containing protease inhibitors and phosphatase inhibitors. Equal amounts of total protein were resolved on 6% acrylamide gels and
analyzed by immunoblotting.
Actin Purification and Iodination--
Actin was prepared from
an acetone powder of rabbit skeletal muscle in buffer A (0.2 mM CaCl2, 0.2 mM ATP, 0.5 mM NaN3, 0.5 mM 2-mercaptoethanol,
2 mM Tris-HCl, pH 8.0) as described (34) and further
purified by gel filtration on a Sephadex G-150 column (35). G-actin was
polymerized in buffer A containing 2 mM
MgCl2/90 mM KCl. Polymerized actin was labeled
with Na125I (5 mCi per 2 mg of actin) using IODO-BEADs
(Pierce, Rockford, IL). 125I-Labeled actin was purified
over a desalting column (D-salt, Pierce, Rockford, IL) and
eluted with 5 mM Tris-HCl, pH 8.0, 0.2 mM ATP,
0.2 mM CaCl2, 0.5 mM dithiothreitol.
F-actin Overlays--
Protein samples were separated by SDS-PAGE
and transferred to nitrocellulose membranes (Schleicher & Schuell,
Keene, NH). Membranes were dried for 30 min and blocked in 10 mM Tris-HCl, pH 7.4, 90 mM NaCl, 0.5% Tween 20 containing 5% nonfat dry milk for 2 h at room temperature.
Membranes were incubated with 125I-labeled actin (50 µg/ml) in blocking buffer containing phalloidin (5 µM,
Cytoskeleton, Denver, CO) at room temperature without agitation. After
2 h, the blots were briefly washed five times with 10 mM Tris-HCl, pH 7.4, 90 mM NaCl, 0.5% Tween 20 and dried. Actin binding was quantified using a PhosphorImager 400B and
ImageQuaNT software. An anti-His tag monoclonal antibody
(Clontech, Palo Alto, CA) was used to detect the
amount of spinophilin in each sample, and immunoreactive protein bands
were quantified using 125I-labeled protein A and
PhosphorImager analysis.
PP1 Overlays--
Protein samples were separated by SDS-PAGE and
transferred to PVDF membranes. Membranes were blocked for 1 h in
TBST (50 mM Tris-HCl, 150 mM NaCl, 0.05% Tween
20, pH 7.4) containing 5% nonfat milk, and then incubated with
bacterially expressed PP1 F-actin Sedimentation--
Phosphorylation reactions were
performed on a large-scale using purified recombinant spinophilin (3 µM, ~500 µg) and in the absence or presence of PKA
(40-100 µg/ml) in 50 mM HEPES, pH 7.4, 10 mM
MgCl2, 1 mM EGTA at 30 °C. Reactions were
initiated by the addition of ATP (50 µM). An aliquot of
the reaction mixture was incubated in parallel with
[
The indicated amounts of spinophilin (62.5-1000 nM, 30 µl) were incubated with 1 µM polymerized actin (0.1 nmol actin/sample, 70 µl) for 30 min at room temperature. The samples
were centrifuged at 200,000 × g for 30 min, and the
pellet was resuspended in SDS-PAGE sample buffer and loaded onto
acrylamide gels. The pelleted actin was stained with Coomassie
Brilliant Blue R-250 and quantified by laser scanning densitometry. The
spinophilin associated with the actin pellet was visualized by
autoradiography after immunoblotting with an anti-spinophilin antibody
and 125I-labeled secondary antibodies and was quantified by
radioactive counting of the excised bands. Samples were incubated in
parallel with spinophilin (62.5-1000 nM) in the absence of
actin to correct for a small amount of pelleted spinophilin in the
background. Standard curves of actin and spinophilin were constructed
for each analysis.
Subcellular Fractionation--
The striatum of Sprague-Dawley
rats was homogenized in ice-cold 0.32 M sucrose, and
subcellular fractions were prepared as described (37). Proteins were
resolved by SDS-PAGE and transferred to PVDF membrane. Immunoblots were
probed with the indicated antibodies and visualized by ECL development.
Spinophilin Is Phosphorylated by PKA--
The amino acid sequence
of spinophilin contains consensus phosphorylation sites for several
protein kinases, including PKA, protein kinase C, and
Ca2+/calmodulin-dependent protein kinases. To
examine whether spinophilin is regulated by phosphorylation in
vivo, striatal slices were labeled with
[32P]orthophosphate, and spinophilin was
immunoprecipitated. A significant level of basal phosphorylation of
spinophilin was detected (Fig. 1A). Moreover, phosphorylation
was increased following treatment with the adenylyl cyclase activator,
forskolin, suggesting that spinophilin might be phosphorylated by
PKA in vivo. Two-dimensional phosphopeptide maps of
spinophilin immunoprecipitated from 32P-labeled slices
revealed multiple phosphopeptides under basal conditions (Fig.
1C), as well as the appearance of novel
phosphopeptides upon treatment with forskolin (peptides labeled
1, 2, and 4, Fig. 1C).
To confirm that PKA directly phosphorylates spinophilin, we performed
phosphorylation reactions in vitro using recombinant spinophilin, PKA, and [ Identification of the PKA Phosphorylation Sites on
Spinophilin--
Spinophilin contains multiple consensus sites for PKA
phosphorylation within its 817-amino acid sequence. No significant
phosphorylation was detected using a truncated spinophilin fragment
consisting of amino acids 298-817 fused to glutathione
S-transferase. We therefore focused on a region at the amino
terminus of spinophilin (amino acids 1-221) that exhibits a high
degree of homology with the actin-binding domain in neurabin (18, 21).
This region was phosphorylated as efficiently as the full-length
protein. Tryptic phosphopeptide maps of the amino-terminal fragment
exhibited the same five major phosphopeptides as were detected using
wild-type spinophilin (Fig. 1E), indicating that all of the
PKA phosphorylation sites are located within the actin-binding domain
of spinophilin.
As shown in Fig. 2A, the
actin-binding domain contains nine consensus sites for PKA
phosphorylation. To identify specific sites of phosphorylation, we
compared the phosphopeptide map of wild-type spinophilin to maps of
alanine mutants at each putative serine phosphorylation site. Mutant
proteins were expressed as histidine fusion proteins in small-scale
bacterial cultures, purified using Ni-NTA agarose, and phosphorylated
with PKA and [
We confirmed the identity of the putative phosphorylation sites by HPLC
and microsequencing analysis. Recombinant spinophilin was
phosphorylated in vitro using PKA and
[ Phosphorylation of Ser-94 and Ser-177 of Spinophilin in Intact
Cells--
Having established the primary sites of phosphorylation
in vitro, we generated phosphorylation state-specific
antibodies to the Ser-94, Ser-100, and Ser-177 sites to analyze the
phosphorylation of spinophilin in greater detail. The
anti-phospho-Ser-94, anti-phospho-Ser-100 and anti-phospho-Ser-177
antibodies recognized spinophilin only upon phosphorylation by
PKA; no detectable binding to dephosphorylated spinophilin was observed
(Fig. 3A). Importantly, each
antibody did not cross-react with the other phosphorylated site of
spinophilin (Fig. 3B). The anti-phospho-Ser-177 antibody
failed to detect phosphorylated spinophilin upon mutation of Ser-177
(but not of Ser-94) to alanine. Similarly, mutation of Ser-94 to
alanine prevented anti-phospho-Ser-94, from detecting phosphorylated
spinophilin. Because Ser-100 is also a putative substrate site for
CaMKII, we examined the possible phosphorylation of spinophilin by
CaMKII. Indeed, CaMKII phosphorylated Ser-100 of spinophilin. However, the physiological significance of this in vitro
phosphorylation is presently unknown.
We then used the phospho-specific antibodies to examine the
phosphorylation of spinophilin in both transfected HEK 293T cells and
in cultured neurons. In either cell type, very low basal
phosphorylation of Ser-94 and Ser-177 was observed, whereas activation
of PKA with forskolin (50 µM) significantly increased the
phosphorylation at both sites (Fig.
4A). In contrast, 1,9-dideoxy
forskolin, an inactive form of forskolin, had no effect on spinophilin
phosphorylation under similar conditions (data not shown). Unlike
Ser-94 and Ser-177, basal phosphorylation of Ser-100 appeared to be
higher in cultured neurons, and incubation with forskolin had little
effect (Fig. 4B).
Involvement of a D1 Receptor/cAMP Cascade in Regulation
of Phosphorylation of Spinophilin in Neurons--
To understand
further the regulation and functional significance of spinophilin
phosphorylation in the brain, we identified signaling pathways that
modulate spinophilin phosphorylation in the neostriatum. Rat
neostriatal slices were incubated with various pharmacological agents,
and the phosphorylation of spinophilin was monitored by immunoblotting
of cell lysates. Because dopamine is known to stimulate adenylyl
cyclase activity in the neostriatum, we evaluated the potential
contribution of the two major classes of dopamine receptors, D1 and D2,
on spinophilin phosphorylation. The D1 class receptors stimulate
adenylyl cyclase and increase cAMP formation (41), whereas D2 class
receptors are coupled to multiple effector systems, including adenylyl
cyclase, Ca2+ and K+ channels, and
phospholipase C (42). Treatment of neostriatal slices with the D1
receptor agonist SKF81297 increased the phosphorylation of both Ser-94
(133 ± 17% of control) and Ser-177 (236 ± 2% of control),
whereas the D2 receptor agonist quinpirole had no effect on basal
phosphorylation levels (Fig.
5A). As expected, forskolin stimulation significantly increased the phosphorylation of both Ser-94
(440 ± 20% compared with control) and Ser-177 (1030 ± 46% of control; Fig. 5A). These results strongly suggest that
dopamine stimulates the phosphorylation of spinophilin via activation
of a D1 receptor/PKA pathway.
We next examined the role of serine/threonine protein phosphatases in
regulating spinophilin dephosphorylation. Neostriatal slices were
incubated with cyclosporin A, a selective protein phosphatase-2B
inhibitor, or with calyculin A or okadaic acid, both inhibitors of PP1
and protein phosphatase-2A (PP2A). Calyculin A, which has previously
been shown to inhibit both PP1 and PP2A activity in neostriatal slices
by ~40% (1 µM (43)), increased the phosphorylation of
Ser-94 and Ser-177 by 5.8- and 4.4-fold, respectively (584 ± 78%
and 439 ± 75% of control; Fig. 5B). Smaller, but
significant, increases of 3.3- and 1.9-fold (332 ± 13% and 185 ± 6% of control), respectively, were produced with okadaic acid at concentrations that inhibit PP2A activity by ~80% and PP1
activity by 5% (200 nM (43)). In contrast, treatment with cyclosporin A resulted in no detectable changes in basal
phosphorylation levels (102 ± 16% for Ser-94 and 95 ± 28%
for Ser-177 of control). These data suggest that PP1, and possibly
also PP2A, can reverse the PKA-mediated phosphorylation of
spinophilin in the neostriatum.
Phosphorylated Spinophilin Is Enriched in Specific Subcellular
Compartments--
We compared the subcellular distributions of
phosphorylated and unphosphorylated spinophilin by immunoblotting with
anti-spinophilin, anti-phospho-Ser-94, or anti-phospho-Ser-177
antibodies. As a control, the various subcellular fractions were also
probed with an antibody selective for the NMDA receptor NR1 subunit,
which is enriched in postsynaptic densities. Spinophilin was found in both cytosolic and membrane-associated fractions of rat neostriatum and
was highly concentrated in the PSD (Fig.
6). A single extraction (Triton
X-100) of the PSD pellet did not remove spinophilin, suggesting that a
significant fraction of spinophilin is tightly associated with the PSD.
Similar to other targeting proteins localized to dendritic spines (44),
further extraction with Sarkosyl detergent disrupted the
interaction of spinophilin with the PSD.
The phosphorylated forms of spinophilin showed striking localization to
specific subcellular compartments. Spinophilin phosphorylated at Ser-94
was concentrated in membrane fractions, including the PSD. In contrast,
spinophilin phosphorylated at Ser-177 was absent from the PSD, although
it remained associated with both the P3 and synaptic plasma membrane
fraction. Importantly, spinophilin phosphorylated at Ser-177 was highly
enriched in the cytosolic S3 fraction. These findings suggest that
phosphorylation of spinophilin at Ser-177 reduces the ability of
spinophilin to interact with one or more specific proteins in the
postsynaptic density. Because phosphorylated spinophilin remains
associated to some extent with the synaptic membrane, our data suggest
that phosphorylation does not induce large movements in the
localization of spinophilin; rather it may trigger subtle alterations
in the targeting of spinophilin within dendritic spines.
The differential distributions of phospho-Ser-94 and phospho-Ser-177
spinophilin suggest that phosphorylation at each site may be
differentially regulated in vivo. The phosphorylation
studies in slices support this notion, because the Ser-177 site
appeared to be more responsive to activation of PKA (10-fold
versus 4.5-fold) than the Ser-94 site, but both sites were
approximately equally responsive to inhibition of PP1/PP2A with
calyculin A (Fig. 5). The greater phosphorylation of Ser-177 may
reflect the different pools of PP1 and PP2A that are present in these
subcellular fractions, in particular the fact that PP1 is enriched in
the PSD fraction (45). Thus, the delicate balance of kinase and
phosphatase activities in neurons may regulate the steady-state levels
of phosphorylation at each site, thereby controlling the targeting of
spinophilin within dendritic spines.
Phosphorylation of Spinophilin Reduces Its Interaction with Actin
Filaments--
Because the two major PKA phosphorylation sites of
spinophilin, Ser-94 and Ser-177, are located within the actin binding
region, we examined whether phosphorylation might influence the
interaction between spinophilin and the actin cytoskeleton. Spinophilin
was phosphorylated in vitro to various stoichiometries and
examined for its ability to bind actin filaments using a
125I-labeled F-actin overlay assay. Under these conditions,
the stoichiometries of phosphorylation of Ser-94 and Ser-177 were
approximately equal at all time points as determined by either
phosphopeptide mapping (see Fig. 1D above) or by the use of
the phospho-specific antibodies to the two sites (Fig.
7C). F-actin binding decreased
in parallel to the increase in the level of spinophilin
phosphorylation, reaching a maximum reduction of ~70% observed at a
sub-maximal stoichiometry of ~1.2 mol/mol (Fig. 7, A and
B). These results suggest that phosphorylation by PKA
directly disrupts the association between spinophilin and F-actin.
We further analyzed the interaction of phosphorylated spinophilin with
actin filaments by performing co-sedimentation studies. Spinophilin was
phosphorylated by PKA to a stoichiometry of 1.6 mol/mol and incubated
with polymerized actin. For comparison, unphosphorylated spinophilin
was analyzed in parallel. Following ultracentrifugation, the amount of
spinophilin associated with the actin pellet was measured by
quantitative immunoblotting (Fig. 8). The
analysis of the binding isotherms indicated that phosphorylation did
not alter the binding affinity of spinophilin for F-actin: dephospho-
and phospho-spinophilin exhibited similar dissociation constants of
455 ± 88 and 490 ± 35 nM, respectively. These
values are close to the Kd of 500 nM
previously reported for dephospho-spinophilin (18). In contrast, the
stoichiometry of the spinophilin-actin interaction decreased by 53%
upon phosphorylation (Bmax of 12 ± 0.7 mol/mol for dephospho-spinophilin, 5.7 ± 0.8 mol/mol for
phospho-spinophilin, p < 0.003). Phosphorylation of spinophilin did not appear to perturb the G- to F-actin equilibrium (data not shown), consistent with the binding of spinophilin to the
sides rather than the ends of actin filaments (18).
In addition to associating with actin, spinophilin binds to PP1 and
inhibits its activity in vitro. We therefore investigated whether phosphorylation might modulate the interaction of spinophilin with PP1. Phosphorylation of spinophilin by PKA, however, had no
detectable effect on PP1 binding in overlay assays (Fig.
7D). Taken together, these results strongly suggest that the
phosphorylation of spinophilin by PKA regulates its association with
the actin cytoskeleton, without disrupting the spinophilin-PP1 complex.
We have provided the first demonstration that protein kinases
modulate the function of spinophilin in neurons. Our data indicate that
spinophilin is phosphorylated by PKA in vitro at Ser-94 and Ser-177, which are located within the actin-binding domain of spinophilin. Ser-94 and Ser-177 are contained within consensus sites
for phosphorylation by PKA and are phosphorylated in neurons in
response to activation of D1 dopamine receptors or forskolin, consistent with both sites being physiological targets for PKA in
intact cells. Ser-100 was identified as a minor site for PKA in
vitro but was not regulated by activation of PKA in
vivo. Interestingly, Ser-94 and Ser-177 of spinophilin are not
conserved in neurabin, a homologue of spinophilin that shares ~48%
amino acid identity with spinophilin. This observation suggests that
the functional activities of spinophilin and neurabin may be
differentially regulated through protein phosphorylation. Indeed,
neurabin is phosphorylated in vitro by PKA at a specific
site that is not found in spinophilin, and phosphorylation appears to
decrease the association of neurabin with PP1 (46).
Phosphorylation of spinophilin by PKA disrupts its ability to associate
with actin filaments. Interestingly, phosphorylated spinophilin bound
to F-actin with high affinity, but with reduced stoichiometry. These
findings suggest that the actin binding region of spinophilin contains
at least two distinct recognition sites for actin. In support of this
idea, we have observed that the actin-binding domain of spinophilin
(amino acids 1-221) is sufficient to bundle actin
filaments.2 Phosphorylation
of spinophilin may disrupt one of these sites, reducing the number of
actin molecules bound to spinophilin without perturbing the binding
affinity of spinophilin for F-actin. Several actin bundling proteins,
including Phosphorylation of spinophilin was associated with decreased binding to
the post-synaptic density. In contrast to unphosphorylated spinophilin,
spinophilin phosphorylated at Ser-177 was absent from the PSD and was
enriched in the cytosolic fractions. This striking difference in
subcellular localization suggests that phosphorylation triggers changes
in the targeting of spinophilin within dendritic spines. Although it is
possible that the pool of spinophilin associated with the PSD cannot be
phosphorylated by PKA, we consider this possibility unlikely. Several
protein kinase A anchoring proteins have been shown to target PKA to
the PSD (50, 51) and enable the kinase to phosphorylate proteins such
as the GluR1 subunit of AMPA receptors (26), NMDA channels (52, 53),
and other receptors. Moreover, phosphorylation of spinophilin at the
Ser-94 site was shown to occur in the PSD. Taken together with our
actin-binding studies, the altered subcellular distribution of
phosphorylated spinophilin supports the notion that PKA activation
dynamically modulates the localization of spinophilin within dendritic spines.
Our previous studies have shown in striatal neurons that spinophilin
plays a role in the regulation of AMPA channels by the dopamine D1
receptor/cAMP/DARPP-32 cascade (19, 20). This is believed to result
from an amplification process that involves both direct phosphorylation
of the channel by PKA and inhibition of PP1 activity via PKA-mediated
phosphorylation of DARPP-32. In these studies, we also observed that
AMPA currents were stabilized by perfusion into neurons of a synthetic
spinophilin peptide that disrupts the targeting of PP1 to spinophilin
(20). This latter result implicated spinophilin in the localized
targeting of active PP1 close to AMPA channels. In the present study,
we have found that phosphorylation by PKA perturbed the interaction of
actin with spinophilin, but that it had no effect on the binding of PP1
to spinophilin. Thus, activation of cAMP-dependent
signaling would be expected to alter the association of the
spinophilin-PP1 complex with the actin cytoskeleton. Phosphorylation of
spinophilin by PKA may therefore contribute to the increased
phosphorylation of AMPA channels by also altering the localization of
PP1 (Fig. 9).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-amino-3-hydroxy-5-methyl-4-isoxazoleproprionic acid (AMPA)-
and N-methyl-D-aspartate (NMDA)-type glutamate
channels and promoting channel down-regulation through
dephosphorylation (19, 20). Moreover, spinophilin-deficient mice
exhibited more persistent AMPA- and NMDA-receptor currents and greatly
reduced long term depression (19).
2-adrenergic receptors (24), and p70 S6
kinase (25). Thus, spinophilin may function as a scaffold protein to
regulate cross-talk between various physiological stimuli in dendritic
spines. However, it remains to be determined whether spinophilin is
regulated by specific neurotransmitter systems in the brain. Notably,
spinophilin contains consensus sequences for phosphorylation by several
protein kinases, including protein kinase A (PKA),
Ca2+/calmodulin-dependent protein kinase II
(CaMKII), and protein kinase C. In the present study, we demonstrate
that spinophilin is phosphorylated in vitro, and likely,
in vivo, by PKA, and we have studied the functional
consequences of spinophilin phosphorylation on its interactions
with actin and PP1.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
80 °C until assayed.
-32P]ATP. The reaction was terminated at various time
points by dilution of the reaction mixture into SDS-PAGE sample buffer,
and the stoichiometry of phosphorylation (moles of Pi/mole
of protein) was assessed after SDS-PAGE and autoradiography, by
measurement of 32P incorporation and normalization to the
amount of protein used.
-D-galactopyranoside at 37 °C. After
1 h, cells were resuspended in ice-cold buffer (50 mM
NaH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8.0, containing protease inhibitors) and
lysed by sonication. The lysate was centrifuged at 12,000 × g for 5 min at 4 °C, and the supernatant was mixed with
Ni-NTA resin (20 µl per sample) for 30 min at 4 °C. The resin was
washed three times with ice-cold buffer (50 mM
NaH2PO4, 300 mM NaCl, 20 mM imidazole, pH 8.0) and once with 50 mM
HEPES, pH 7.4, 10 mM MgCl2. Phosphorylation
reactions were carried out as described above with the proteins bound
to the resin. After 1 h, reactions were terminated with SDS-PAGE
sample buffer, and the phosphorylated proteins were loaded onto 8%
acrylamide gels for autoradiography and phosphopeptide mapping analysis.
-32P]ATP.
The reaction mixture was separated by SDS-PAGE and electrophoretically transferred to a PVDF membrane. The 32P-labeled spinophilin
was localized by autoradiography, and the membrane was excised and
subjected to proteolytic digestion with trypsin or GluC proteases.
Eluted peptides were separated by HPLC, and fractions eluting from the
capillary column were spotted directly onto a strip of PVDF membrane
using a 153 microblotter (PerkinElmer Life Sciences, Boston, MA (30)).
Membrane pieces containing the 32P-labeled phosphopeptides
were excised and subjected to automated amino-terminal protein
microsequencing and mass spectrometry.
80 °C until assayed.
Frozen tissue samples were sonicated in boiling 1% SDS containing
protein phosphatase and protease inhibitors. The protein concentrations
of the homogenates were determined using the BCA protein assay method
(Pierce, Rockford, IL) with bovine serum albumin as a standard. Equal
amounts of protein (100 µg) were separated by SDS-PAGE using 6%
acrylamide gels and transferred to PVDF membranes (0.2 µm). Membranes
were blocked for 1 h in TBS (50 mM Tris-HCl, 150 mM NaCl, pH 7.4) containing 5% (w/v) nonfat dry milk and
0.1% Tween 20 and then incubated with rabbit polyclonal antibodies
against spinophilin, phospho-Ser-94 spinophilin, phospho-Ser-100
spinophilin, or phospho-Ser-177 spinophilin. Following washes with TBS
containing 1% nonfat milk, 0.1% Tween 20, and 0.5% bovine serum
albumin, proteins were visualized by incubation with anti-rabbit
horseradish peroxidase-conjugated secondary antibodies (1:5000
dilution, Pierce, Rockford, IL) and enhanced chemiluminescence (ECL)
development (Amersham Biosciences, Piscataway, NJ). Chemiluminescence was detected by exposure of blots to photographic film, and bands were
quantified by analysis of scanned images using Image 1.52 software
(National Institutes of Health). Because the linear range for
quantification of signal density by the ECL detection method is limited
to <10-fold, we routinely exposed chemiluminescent membranes to film
for various time periods to obtain signals within the linear range.
Data were analyzed by the Mann-Whitney U test, with
significance defined as p < 0.05.
(1.5 µg/ml) in 50 mM
Tris-HCl, pH 7.0, 0.1 mM EGTA, 15 mM
2-mercaptoethanol, 0.01% brij-35, 0.3 mg/ml bovine serum albumin at
4 °C overnight. After washing twice with TBST containing 0.25%
nonfat milk, membranes were probed with an anti-PP1
antibody (1.5 µg/ml in TBST containing 1% nonfat milk (36)) for 1 h.
Following several washes, proteins were visualized by incubation with
anti-rabbit horseradish peroxidase-conjugated secondary antibodies and
ECL development.
-32P]ATP for determination of the phosphorylation
stoichiometry (1.6 mol/mol phosphorylated approximately equally at
Ser-94 and Ser-177). After 1 h, the reaction mixtures were
exhaustively dialyzed into 20 mM Tris-HCl, 50 mM NaCl, pH 7.4. Immediately prior to the actin sedimentation experiments, proteins were centrifuged at 200,000 × g for 30 min. Protein concentrations were determined using
the BCA protein assay method with bovine serum albumin as a standard.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Phosphorylation of spinophilin by PKA.
A, spinophilin was immunoprecipitated from
32P-labeled neostriatal slices treated without or with
forskolin. The incorporation of [32P]phosphate into
spinophilin was visualized by autoradiography. B, time
course for the in vitro phosphorylation of spinophilin by
PKA, with comparison to inhibitor-1. Proteins were phosphorylated
in vitro using PKA and [ -32P]ATP. The
stoichiometry of phosphorylation was assessed after SDS-PAGE and
autoradiography. C, phosphopeptide maps of spinophilin
immunoprecipitated from neostriatal slices incubated without (basal
slice) or with forskolin (forskolin-treated slice). Proteins were
excised from gels as shown in panel A and digested with
trypsin. The resultant phosphopeptides were separated by
two-dimensional phosphopeptide mapping and visualized by
autoradiography. D, phosphopeptide maps of spinophilin
phosphorylated for 60 min (left panel) or 1 min (right
panel). The autoradiograms were exposed for different times to
give approximately equivalent intensities of the phosphopeptides.
E, phosphopeptide map of the actin-binding domain of
spinophilin (residues 1-221). For D and E,
proteins were phosphorylated in vitro with PKA and
[
-32P]ATP and analyzed by SDS-PAGE and two-dimensional
phosphopeptide mapping. Peptides were labeled 1-5 based on the pattern
observed for full-length spinophilin phosphorylated by PKA in
vitro, with the number just to the right of the relevant
phosphopeptide. In the peptide maps obtained from spinophilin
phosphorylated in intact cells, peptide 5 was not consistently
detected.
-32P]ATP. Phosphorylation
proceeded in a time-dependent manner and reached a maximal
stoichiometry of 2.3 mol/mol within 60 min (Fig. 1B). The
initial rate of phosphorylation was comparable to that of inhibitor-1,
a physiological substrate of PKA known to be phosphorylated at a single
site. Two-dimensional phosphopeptide maps of spinophilin phosphorylated
by PKA to low (1-min incubation) or higher (60-min incubation)
stoichiometry revealed the presence of five major phosphopeptides
(labeled 1-5, Fig. 1D). The phosphopeptides
phosphorylated in vitro by PKA corresponded to the pattern
of phosphopeptides phosphorylated in spinophilin in slices in response
to forskolin (Fig. 1C) (with the exception that peptide 5 could not be detected in spinophilin phosphorylated in slices).
Together, these results support the conclusion that PKA phosphorylates
spinophilin in vivo at a subset of the total sites
phosphorylated in intact cells.
-32P]ATP. Mutation of Ser-94 led to the
specific disappearance of phosphopeptides 1 and 2 (Fig. 2B,
left panel); mutation of Ser-100 removed phosphopeptide 3 from the maps (Fig. 2B, middle panel), and
mutation of Ser-177 eliminated phosphopeptides 4 and 5 (Fig. 2B, right panel). The disappearance of two
phosphopeptides upon mutation of a single serine/threonine site has
previously been observed (38) and most likely results from multiple
tryptic cleavage sites. Interestingly, the phosphopeptide map of the
Ser-94 mutant showed increased phosphorylation at Ser-100, suggesting that elimination of this primary site of phosphorylation enhances the
phosphorylation kinetics of secondary sites. Mutation of all other
putative phosphorylation sites (Ser-17, Ser-59, Ser-87, Ser-99,
Ser-122, and Ser-126) had no effect on the phosphopeptide map patterns
(data not shown).
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Fig. 2.
Identification of the PKA phosphorylation
sites on spinophilin. A, the actin-binding domain of
spinophilin contains nine consensus sites for PKA phosphorylation
(underlined). The identified sites of phosphorylation
(Ser-94, Ser-100, and Ser-177) are boxed. B,
two-dimensional phosphopeptide maps of alanine mutants of spinophilin
(S94A, S100A, S177A).
-32P]ATP, and the tryptic phosphopeptides were
purified by HPLC. Microsequencing analysis of the radiolabeled peptides
identified two major peptides encompassing the Ser-94 and Ser-100 sites
(amino acids 93-97 and 98-111; Table
I). A third, weakly radiolabeled HPLC
fraction was found to contain the peptide 59-85. Peptide 59-85 has
likely co-purified with a phosphopeptide of low abundance (too low to
sequence); however, we cannot rule out the possibility that this
peptide contains a minor phosphorylation site that was not detected by
phosphopeptide mapping. Although initial studies using the protease
trypsin did not identify a peptide encompassing Ser-177, digestion with
the protease Glu-C produced a phosphopeptide that could be resolved by
HPLC and identified as amino acids 175-194. Together with the
phosphopeptide mapping studies, these results indicate that Ser-94 and
Ser-177 of spinophilin are phosphorylated efficiently by PKA at
comparable rates. In contrast, Ser-100 of spinophilin is phosphorylated
by PKA at a significantly lower rate.
Sequencing of phosphopeptides from spinophilin phosphorylated by
PKA
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Fig. 3.
Specificity of the phosphorylation state
specific antibodies. A, spinophilin was phosphorylated
in vitro without (dephos) or with PKA (PKA
phos) or with CaMKII (CaMKII phos) as indicated, and
phosphorylation state-specific antibodies to phospho-Ser-94,
phospho-Ser-100, or phospho-Ser-177 were used to detect phosphorylated
spinophilin by immunoblotting. B, the antibodies displayed
no cross-reactivity toward other phosphorylated sites of spinophilin.
Wild-type (WT) spinophilin and mutants in which Ser-94
(S94A), Ser-100 (S100A), or Ser-177 (S177A) were replaced with alanine
were phosphorylated in vitro without (dephos) or
with PKA or CaMKII (phos) as indicated. Proteins were
resolved by SDS-PAGE and transferred to PVDF membranes. Samples were
analyzed by immunoblotting with the phospho-Ser-94, phospho-Ser-100, or
phospho-Ser-177 antibodies as indicated.
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Fig. 4.
Spinophilin is phosphorylated in response to
activation of PKA at Ser-94 and Ser-177 in HEK 293T cells and cultured
neurons. A and B, 293T cells, transiently
transfected with spinophilin, or cultured neurons were incubated in the
absence (basal) or presence of forskolin (50 µM) for 5 min. Proteins were resolved by SDS-PAGE and transferred to PVDF
membranes. Spinophilin was detected by immunoblotting with the
indicated antibodies.
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Fig. 5.
Regulation of spinophilin phosphorylation in
the neostriatum. A, effect of forskolin, the D1
receptor agonist SKF81297 and the D2 receptor agonist quinpirole on the
phosphorylation of spinophilin. Neostriatal slices were incubated with
forskolin (50 µM), SKF81297 (1 µM), or
quinpirole (1 µM) for 5 min. B, effect of
phosphatase inhibitors on the phosphorylation of spinophilin.
Neostriatal slices were incubated with calyculin A (200 nM), okadaic acid (1 µM), or cyclosporin A (5 µM) for 60 min. For both A and B,
the homogenates were subjected to SDS-PAGE and immunoblotted with the
indicated phospho-specific antibodies. The amount of phosphorylated
spinophilin was quantified by densitometry, and the data were
normalized to the values obtained with untreated slices. Data represent
means ± S.E. for three or four experiments. *, p < 0.05; Mann-Whitney U test compared with control.
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Fig. 6.
Phosphorylated spinophilin is localized to
discrete subcellular compartments. Rat neostriatal homogenates
were fractionated as described under "Experimental Procedures," and
the subcellular fractions were analyzed by immunoblotting with the
indicated antibodies. 25 µg of total protein was loaded in each lane,
with the exception of the PSD fractions, which contained 15 µg. As
expected, NMDA receptor immunoreactivity was enriched in the PSD
fractions.
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Fig. 7.
Phosphorylation disrupts the interaction of
spinophilin with actin filaments but not with PP1. Spinophilin was
phosphorylated in vitro to various stoichiometries and
examined for its ability to bind (A) actin filaments or
(D) PP1. An anti-histidine tag antibody was used to detect
total spinophilin levels. B, the phosphorylation
stoichiometry and extent of actin binding were quantified using a
PhosphorImager 400B and ImageQuaNT software. The values for actin
binding represent the fraction of the total amount of actin bound at
time zero. Data represent means ± S.E. for three experiments.
C, the relative rates of phosphorylation of Ser-177 and
Ser-94 were determined by immunoblotting aliquots of samples of
spinophilin phosphorylated at different times.
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Fig. 8.
Phosphorylation decreases the stoichiometry
of the spinophilin-actin interaction. A, polymerized
actin (1 µM) was incubated with the indicated amounts of
dephosphorylated or phosphorylated spinophilin (phosphorylated to 1.6 mol/mol). Following ultracentrifugation, the amount of spinophilin
associated with the actin pellet was measured by quantitative
immunoblotting. B and C, binding isotherms
(B) indicate that phosphorylation markedly decreases the
binding stoichiometry (Bmax) without affecting
the dissociation constant (Kd) of the
spinophilin-actin interaction (C). Data represent means ± S.E. for three experiments.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-actinin, dystrophin, and spectrin, possess multiple
actin-binding domains, which contact actin molecules via
-helical
structures (47-49). Under the conditions used in the present study,
both Ser-94 and Ser-177 were phosphorylated to approximately equal
levels. Ser-94 and Ser-177 reside in regions predicted to adopt
-helices, and phosphorylation of spinophilin at either or both sites
may prevent the binding of actin molecules through destabilization of
the protein-protein interface.
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Fig. 9.
Model for the regulation of AMPA-type
glutamate receptors by spinophilin. Spinophilin anchors PP1
in the vicinity of the AMPA channel by binding to actin filaments.
Under basal conditions, the spinophilin-PP1 complex maintains the AMPA
channel in a dephosphorylated state in which it is relatively
insensitive to its neurotransmitter, glutamate. After D1 receptor
stimulation, AMPA channel phosphorylation is increased due to
phosphorylation of spinophilin by PKA, likely targeted close to the
synapse via protein kinase A-anchoring proteins. In turn, the
spinophilin-PP1 complex is removed from the vicinity of the channel and
leads to synergistic increases in phosphorylation of the AMPA
receptor.
The ability of cAMP-dependent signaling cascades to alter dynamically the localization of the spinophilin-PP1 complex may also have important consequences for synaptic plasticity. PP1 has been found to play a role in synaptic plasticity likely by modulating AMPA type glutamate channels important for synaptic transmission, and targeting of PP1 has been proposed to play a role in long term depression (54, 55). Spinophilin knockout mice exhibit altered glutamatergic transmission and reduced long term depression (19), suggesting that spinophilin is responsible for the regulation of AMPA receptors by PP1. Phosphorylation of spinophilin by PKA may contribute to regulation of ion channel conductances critical for the maintenance and plasticity of dendritic spines (56, 57). Phosphorylation of spinophilin may also influence trafficking of AMPA receptors by modulating the actin cytoskeleton in dendritic spines.
Spinophilin phosphorylation could have other important consequences for the structure and function of dendritic spines. Changes in the number, size, and shape of dendritic spines have been associated with learning, electrophysiological, developmental, and hormonal changes (4-12). The ability of dendritic spines to change shape has been attributed to a dense network of proteins that facilitates the rapid assembly and disassembly of the actin cytoskeleton (13, 14). Spinophilin has been shown to cross-link actin filaments in vitro and to regulate the development of dendritic spines in vivo (19). Thus, phosphorylation of spinophilin may influence the morphology of spines by modulating the bundling or polymerization of actin filaments. Alternatively, phosphorylation may serve to control PP1-mediated changes in the actin cytoskeleton. Previous studies have shown that PP1 regulates cellular morphology through dephosphorylation of the actin cytoskeleton (58-60).
In conclusion, we have demonstrated that phosphorylation of spinophilin
is increased in response to dopamine D1 receptor/cAMP signaling
cascades. Spinophilin phosphorylation, in turn, modulates its ability
to interact with the actin cytoskeleton. Our findings suggest that
phosphorylation of spinophilin by PKA regulates anchoring of the
spinophilin-PP1 complex within dendritic spines. These studies may
provide greater insight into the molecular mechanisms that underlie
synaptic plasticity and dendritic spine morphology. Future studies will
examine the dynamic targeting of the spinophilin-PP1 complex in living
neurons and its impact on the structure and function of dendritic spines.
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ACKNOWLEDGEMENTS |
---|
We thank Martin Lan, Mercedes Paredes, Dr. Stacie Grossman, Jean Whitesell (Cocalico Biologicals, Inc.), and Dr. Joseph Fernandez (Rockefeller University Protein/DNA Technology Center) for valuable assistance.
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FOOTNOTES |
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* This work was supported by United States Public Health Services Grants MH40899 and DA10044 and by Fellowship DRG-1451 of the Cancer Research Fund of the Damon Runyon-Walter Winchell Foundation (to L. C. H.-W.).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.
** To whom correspondence should be addressed: Laboratory of Molecular and Cellular Neuroscience, The Rockefeller University, 1230 York Ave., New York, NY 10021. Tel.: 212-327-8871; Fax: 212-327-7888; E-mail: nairn@mail.rockefeller.edu.
Published, JBC Papers in Press, November 1, 2002, DOI 10.1074/jbc.M205754200
2 L. C. Hsieh-Wilson and P. Greengard, unpublished observations.
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ABBREVIATIONS |
---|
The abbreviations used are:
PP1, protein
phosphatase-1;
PKA, protein kinase A;
PSD, postsynaptic density;
AMPA, -amino-3-hydroxy-5-methyl-4-isoxazoleproprionic acid;
NMDA, N-methyl-D-aspartate;
CaMKII, Ca2+/calmodulin-dependent protein kinase II;
HPLC, high-performance liquid chromatography;
PP2A, protein
phosphatase-2A;
PVDF, polyvinylidene difluoride;
Ni-NTA, nickel-nitrilotriacetic acid.
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