(Received for publication, December 5, 1996, and in revised form, January 23, 1997)
From the Department of Biochemistry and Molecular
Biology and the Molecular, Cellular and Integrative Neuroscience
Program, Colorado State University, Fort Collins, Colorado 80523 and
§ The Children's Medical Research Institute, 214 Hawkesbury Road, Westmead, New South Wales 2145, Australia
Myoblasts, transfected with a human gene encoding
a -actin point mutation, down-regulate expression of actin
depolymerizing factor (ADF) and its mRNA. Regulation is
posttranscriptional. Expression of cofilin, a structurally similar
protein, and profilin, CapG, and tropomodulin is not altered with
increasing mutant
-actin expression. Myoblasts expressing either
human
-actin or the mutant
-actin down-regulate the endogenous
mouse actin genes to keep a constant level of actin mRNA, whereas
the
-actin transfectants do not down-regulate ADF. Thus, ADF
expression is regulated differently from actin expression.
The mutant -actin binds to ADF with about the same affinity as
normal actin; however, it does not assemble into normal actin filaments. The decrease in ADF expression correlates with an increase in the unassembled actin pool. When the actin monomer pool in untransfected myoblasts is increased 70% by treatment with latrunculin A, synthesis of ADF and actin are down-regulated compared with cofilin
and 19 other proteins selected at random. Increasing the actin monomer
pool also results in nearly complete phosphorylation of both ADF and
cofilin. Thus, ADF and cofilin are coordinately regulated by
posttranslational modification, but their expression is differentially
regulated. Furthermore, expression of ADF is responsive to the
utilization of actin by the cell.
Regulation of the synthesis and assembly of cytoskeletal
components is critical to cell survival. Although it has been
recognized for many years that cellular demand for cytoskeletal
components can drive the synthesis of monomer (1, 2), the
autoregulatory mechanisms have been determined in detail only for
-tubulin (3-5).
Actin is also subject to feedback regulation although the mechanism is
not established (2). Overexpression of human genes encoding -actin
or a
-actin single mutant
(
sm-actin)1 (Gly-244 to Asp)
in mouse C2 myoblast cells results in the down-regulation of endogenous
mouse
- and
-actin genes, thus maintaining a constant level of
actin mRNA and protein (6). In contrast, overexpression of a human
gene encoding a highly unstable mutant actin protein fails to elicit
down-regulation of the endogenous mouse genes. This suggests that the
feedback regulation is directed by the protein product of that
introduced gene (6).
Feedback regulation of actin synthesis by an increased monomer pool may involve the control of mRNA stability. Synthesis of actin decreases in cells that have been treated with the actin depolymerizing drugs, latrunculin A (Lat A) and botulinum toxin C2 (7-9). The decline in rates of actin synthesis in Lat A and C2 toxin-treated cells can be largely accounted for by the observed decrease in mRNA stability within the cytoplasm (8, 9). On the other hand, de novo synthesis of actin mRNA increases in cells in which actin assembly has been promoted by incorporation of phalloidin (9), suggesting that two different autoregulatory control mechanisms, one nuclear and the other cytoplasmic, work to maintain actin homeostasis.
Actin is also capable of reprogramming the expression of
microfilament-associated proteins. High level expression of different human actin genes in mouse C2 cells impacts on the expression and
organization of different tropomyosin isoforms (10) and on the
expression of vinculin and talin (11). Vinculin expression is also
decreased by drugs that depolymerize actin, and vinculin down-regulation requires the presence of a nucleus (8). In addition,
changes in actin and vinculin expression in response to drug treatment
are independent of changes in cell shape (8). High level expression of
tropomyosin Tm1 (12), vinculin (13, 14), -actinin (15), and gelsolin
(16), which all alter cell morphology, have no impact on the expression
of other microfilament proteins. This suggests that the ability of
actin to reprogram expression of microfilament proteins may be unique
to actin.
The assembly of the actin cytoskeleton depends upon the production and
activities of a large number of actin binding proteins (reviewed in
Refs. 17-19). Among the most important of these proteins are those in
the profilin, thymosin, and actin depolymerizing factor (ADF)/cofilin
families that have been implicated in regulating actin assembly in a
number of different systems (reviewed in Refs. 20 and 21). How
responsive the cellular levels of these proteins are to actin
utilization is unknown. The mutant human sm-actin is
known to form aberrant actin filaments (22), and thus, clones of cells
expressing different amounts of this protein provide a good model
system to determine which of the actin assembly regulatory proteins are
responsive to its nonfilamentous accumulation. Clones of cells
expressing equivalent amounts of the assembly competent, normal human
- and
-actins provide useful controls.
In various C2 cell clones, expressing human sm-actin and
normal human
- and
-actins, we examined the levels of three actin monomer binding proteins ADF, cofilin, and profilin. In addition, we
looked at the expression of CapG (23) and tropomodulin (24), two
proteins present in myoblasts that can also alter actin assembly dynamics and filament lengths. The expression of only one of these proteins, ADF, is inversely proportional to the levels of the assembly
mutant,
sm-actin. Further analysis of the same C2 cell clones showed an increase in G-actin concomitant with a decrease in ADF
expression, suggesting that ADF may be a critical assembly regulatory
protein sensitive to the utilization of actin by the cell. To explore
this hypothesis further, untransfected C2 cells were treated with Lat A
to depolymerize F-actin and increase the unassembled actin pool. As
previously shown in Swiss 3T3 and HeLa cells (8), the Lat A treatment
induced a down-regulation in actin synthesis. Results presented below
demonstrate that the synthesis of ADF, but not cofilin, decreases when
the pool of unassembled actin increases. In addition, C2 cells respond
to Lat A by nearly complete phosphorylation (inactivation) of both ADF
and cofilin. Thus, although the posttranslational regulation of ADF and
cofilin is coordinated, their synthesis is regulated independently. We
propose that perturbations in the actin monomer pool regulate the
expression of ADF, but not cofilin, in C2 cells.
Purification and Quantification of Proteins
The following proteins were generous gifts of the individuals listed: calf spleen profilin and CapG from Dr. Helen Yin, University of Texas Southwestern Medical School; tropomodulin from Dr. Mark Sussman, University of Southern California. ADF was isolated from chick embryo brain (25). Recombinant chick ADF (26) and recombinant cofilin (27) were prepared in the laboratory as described previously. Skeletal muscle actin was purified from rabbit muscle acetone powder (28), and brain actin was purified from 18-day embryonic chick brain (29). Protein concentrations were determined by the filter paper dye-binding method, using ovalbumin as a standard (30).
Cell Culture
The C2C12 murine myoblast cell line was maintained as a monolayer in 75-cm2 tissue culture flasks (Corning Laboratories, Corning, NY) in growth medium containing Dulbecco's modified Eagle's medium (low glucose), 20% fetal bovine serum, and 0.5% chick embryo extract (all from Life Technologies, Inc.). For experiments, cells were cultured in 6- or 10-cm tissue culture dishes at less than 30% confluence. All actin transfected cell clones were previously described (31).
Nuclear Run-on Transcription
C2 cells (2-5 × 107) were washed with 4 °C
phosphate-buffered saline (PBS) and scraped off plates into 4 ml of 50 mM Tris, pH 8.0, 100 mM NaCl, 5 mM
MgCl2, 0.5% (v/v) Nonidet P-40 at 4 °C, and nuclei were
prepared (32). The nuclei were resuspended in 100 µl of 50 mM Tris, pH 8.3, 40% glycerol, 5 mM
MgCl2, 0.1 mM EDTA and assayed immediately. To
each nuclei sample was added 100 µl of 10 mM Tris, pH 8, 5 mM MgCl2, 0.3 M KCl, 0.5 mM DTT, 10 mM GTP, 10 mM ATP, and
10 mM CTP and 200 µCi of [32P]UTP (3000 Ci/mmol), and the mixture was incubated at 26 °C for 10 min (33).
Following DNase I and proteinase K digestions, the RNA was extracted
and precipitated (34). The RNA was dissolved in 500 µl of rapid-hyb
buffer (Amersham Corp.) and hybridized to slot blots containing 10 µg
of cut, denatured plasmids containing probe sequences for actin
(pTRI--actin-mouse) and human 18 S ribosomal RNA (pT7 RNA 18 S)
(Ambion, Inc., Austin, TX) or human ADF (courtesy of Alan Weeds,
Cambridge, UK). The blots were prehybridized overnight at 65 °C, and
3.5 ml of fresh rapid-hyb buffer was added prior to the addition of the
labeled RNA. Blots were hybridized for 48 h and then washed 2 × for 60 min with 50 ml of 2 × SSC (0.3 M NaCl, 0.03 M sodium citrate, pH 7.0) at 65 °C, 1 × for 60 min
in 8 ml of 2 × SSC with 8 µl of 10 mg/ml RNase A at 37 °C, and 1 × for 30 min at 65 °C in 2 × SSC and 0.1% SDS.
Radioactivity was detected and quantified with a PhosphorImager
(Molecular Dynamics, Sunnyvale, CA). The solution of labeled RNA from
the first hybridization was rehybridized to a duplicate blot to confirm
complete hybridization of specific RNA in the first incubation.
Gel Electrophoresis and Immunoblotting
Sample PreparationCultured C2 cells were washed rapidly 4 × in 4 °C PBS, lysed into SDS extraction buffer (10 mM Tris, pH 7.5, 2% SDS, 10 mM NaF, 5 mM dithiothreitol (DTT), 2 mM EGTA) by scraping, and the extracts were heated in boiling water for 3 min. After cooling, the samples were sonicated briefly and the proteins precipitated (35). The proteins were dissolved in 2 × sample preparation buffer (1 × buffer contains 0.125 M Tris, pH 6.8, 0.5% SDS, 5% glycerol, 5% 2-mercaptoethanol, 0.005% bromphenol blue) for SDS-PAGE.
SDS-PAGE and ImmunoblottingSDS-PAGE was performed by the method of Laemmli (36) on 15% total acrylamide (2.7% cross-linker) isocratic mini-slab gels. Proteins were transferred electrophoretically to polyvinylidene difluoride (Immobilon P, Millipore Corp., Bedford, MA) for 1 h at 0.3 A in the buffer of Towbin et al. (37), using a Genie Electroblotter (Idea Scientific, Minneapolis, MN), blocked, washed, and immunostained as described previously (25). Alkaline phosphatase-conjugated secondary antibody (Sigma) was diluted in the wash buffer. Blots were developed with Lumiphos (Boehringer Mannheim) after a quick rinse in 100 mM Tris, pH 9.5, 100 mM NaCl, 50 mM MgCl2, or with CDP-star (Tropix Inc., Bedford, MA) after a quick rinse in 50 mM Tris, pH 9.5, 100 mM NaCl, 1 mM MgCl2 to get exposures within the linear range of the Hyperfilm ECL (Amersham Corp.). Following chemiluminescent detection, blots were immunostained with NBT/BCIP (Life Technologies, Inc.) according to manufacturer's directions. Exposures of the chemiluminescence images and the stained blots were analyzed with a Microscan 2000 image analysis system (Technology Resources Inc., Knoxville, TN). Internal standards of proteins were included on every immunoblot.
Antibodies
Mouse monoclonal antibody (C4) to actin was purchased from ICN
Pharmaceuticals, Inc. (Costa Mesa, CA). -Actin-specific rabbit antiserum was a gift from Dr. J. Chloe Bulinski, Columbia University, NY. Monoclonal antibody to the
-actin isoform was from Sigma. Rabbit
polyclonal antibody to ADF was prepared by us (38). Mouse monoclonal
antibody (MAb22) to cofilin was a gift from Drs. Hiroshi Abe and
Takashi Obinata, Chiba University, Japan (39). Polyclonal rabbit
antibodies to human profilin and CapG were a gift from Dr. Helen Yin,
University of Texas Southwestern Graduate School, Dallas, TX. Affinity
purified rabbit IgG to tropomodulin was a gift from Dr. Mark Sussman,
University of Southern California, Los Angeles, CA.
Immunoprecipitation
Supernatants from the homogenates of C2 myocyte cultures were prepared by scraping the washed cells from a 6-cm culture dish in 300 µl of 10 mM HEPES, pH 7.4, 0.15 M NaCl, 1 mM DTT, and 1% Triton X-100 (IP buffer), sonicating the cell suspension for 3 s, and centrifuging at 10,000 × g for 5 min at 4 °C. The supernatant was then preincubated on a rotator with 240 µl of a 1:1 suspension of protein A-agarose in IP buffer at 4 °C for 5 min. The sample was centrifuged 10 s at 10,000 × g and the supernatant divided into two tubes. Anti-ADF IgG (2 mg/ml; 120 µl) or preimmune serum was added to each tube, and the tubes were rotated at 4 °C overnight. Protein A-agarose (60 µl of a 1:1 suspension) in IP buffer was added to each tube (rotated at 4 °C) for 1 h. The resin was centrifuged, washed once with 300 µl of IP buffer, and the bound antibody complexes extracted with 30 µl of 1% SDS in a boiling water bath. After centrifugation, the supernatant was mixed with an equal volume of 4 × sample preparation buffer for SDS-PAGE and immunoblotting.
Affinity Chromatography
The actin isoforms binding to ADF were also examined by using ADF-Affi-Gel resin. All steps were carried out at 4 °C and under nitrogen. Recombinant ADF (5.9 mg) was dialyzed against degassed 0.1 M MOPS, pH 7.5, for 4 h and then added to 1 ml of Affi-Gel 10 resin (Bio-Rad) hydrated in degassed MOPS buffer. After mixing for 2 h, 0.1 ml of 1 M ethanolamine, pH 8, was added, and the sample was mixed for 1 h. DTT (1 M; 3 µl) was then added (nitrogen atmosphere was not needed after this step), and the resin was placed in a column and washed with 25 ml of MOPS buffer containing 1 mM DTT. Supernatants of extracts from cultured C2 cells or C2 transfectants were prepared as described under immunoprecipitation except that the lysis buffer was 0.1 M Tris, pH 8.0, and 1 mM DTT (600 µl/6-cm dish). Supernatant (100 µl) was added to 60 µl of a 1:1 suspension of the ADF resin in 0.1 M MOPS, pH 7.5, 1 mM DTT, or to the same volume of Sepharose 4B resin as a control. The samples were incubated on a rotator for 5 min at 4 °C. After microcentrifuging for 10 s, the resin was washed once with 300 µl of 1 M Tris, pH 8.0, 1 mM DTT. The bound proteins were extracted with 40 µl of 1% SDS in a boiling water bath. Samples were diluted with an equal volume of 4 × sample preparation buffer, and actin isoforms were identified by SDS-PAGE and immunoblotting.
RNA Blotting and Analysis
RNA was isolated from duplicate sets of four 10-cm plates of C2
cells and each of the transfected clones using the guanidinium thiocyanate method (34). RNA from the duplicate samples was separated
by agarose-formaldehyde electrophoresis, transferred to Hybond-N+
membrane (Amersham Corp.), and hybridized with a 260-base pair fragment
of the human ADF cDNA (339-599 HindIII fragment; Ref.
40) which had been labeled with 32P by the random primer
method (41) or with an 18 S ribosomal RNA probe (used in excess),
end-labeled with T4 polynucleotide kinase, for normalization of loading
the gel (6). Hybridization of the ADF probe was carried out in 4 × SSC, 5 × Denhardt's solution (42), 50 mM
NaH2PO4, and 10% dextran sulfate at 65 °C
overnight and for the 18 S probe in the same solution at 55 °C
overnight. Filters were washed 4 × for 30 min at 50 °C in
0.5 × SSC, 0.1% SDS for the ADF probe, and 2 × for 20 min
at room temperature and 3 × for 20 min at 55 °C in 4 × SSC, 0.1% SDS for the 18 S rRNA probe. Autoradiography was performed
at 70 °C with either Kodak XAR-5 film or Amersham Hyperfilm-MP.
RNA bands on the autoradiogram (subsaturating levels) were quantified
using the Microscan 2000 image analysis system. The integrated density
of each ADF mRNA band was normalized to the amount of 18 S RNA in
that sample with the value from wild type C2 cells taken as 100%.
Quantification of G-actin in Cultured Cells
Cultured cells were washed free of medium with four washes of 4 °C PBS and lysed in 10 mM Tris, pH 7.5, 2 mM MgCl2, 0.5 mM DTT, 2 mM EGTA, 1% Triton X-100, and 7.5% glycerol. Material was scraped to the edge of the plate and transferred with a wide bore pipette to a microcentrifuge tube for G-actin quantification or to an airfuge tube for preparation of supernatant and cytoskeletal fractions. The amount of G-actin in each lysate was determined by the DNase I inhibition assay (43, 44) using DNase I calibrated with purified skeletal muscle G-actin. Supernatant and cytoskeletal (pellet) fractions were prepared from the cell extracts by centrifugation of the lysates at 170,000 × gmax for 20 min. Actin in each fraction was determined by immunoblot analysis.
Preparation of Triton-soluble and Cytoskeletal Fractions
Lat A (a generous gift from Ilan Spector, SUNY, Stony Brook, NY) was added to C2 cell cultures to 5 µM for 0, 6, 12, and 18 h at 37 °C. The cells were washed four times with PBS (2 ml) at room temperature and extracted at room temperature with 50 mM MES, pH 6.5, 1 mM EGTA, 50 mM KCl, 1 mM MgCl2, 1 mM phenylmethylsulfonyl fluoride, 10 mM NaF, 0.5% Triton X-100, and 0.5% protease inhibition mixture (45) (0.45 ml/6-cm dish). The Triton-soluble fraction was transferred to a microcentrifuge tube containing 50 µl of 20% SDS and heated to boiling for 5 min. The cytoskeletal fraction was solubilized in SDS extraction buffer, transferred to a microcentrifuge tube, and heated in boiling water for 5 min. After cooling to room temperature, the cytoskeletal fraction was briefly sonicated to reduce the viscosity due to DNA. The proteins in both fractions were precipitated (35) and redissolved in 50 µl of 3 × sample preparation buffer. For Western blotting, 1 µg of protein/fraction and 10-50 ng of actin standards were loaded.
Measurement of Rates of Protein Synthesis
C2 cells on 6-cm dishes were treated with Lat A (5 µM) for 0, 6, 12, and 18 h at 37 °C. Thirty minutes prior to the end of the Lat A treatment, the cells were washed twice with prewarmed methionine-, cysteine-, and serum-free Dulbecco's modified Eagle's medium (labeling medium). The cells were then incubated with 200 µCi/ml [35S]methionine/cysteine (Promix; Amersham Corp.) in labeling medium containing Lat A (except for control dishes) for 30 min at 37 °C. The labeling solution was aspirated and the cells washed twice with 2 ml of PBS at room temperature. The cells were lysed in 150 µl/dish SDS extraction buffer and the extracts immediately heated to boiling for 3 min. After cooling, the samples were sonicated briefly, the proteins precipitated (35), and then dissolved in 9.5 M urea, 5% 2-mercaptoethanol, and 2% Nonidet P-40 or Igepal (Sigma).
Proteins were separated by two-dimensional PAGE using nonequilibrium pH gradient electrophoresis in the first dimension (46) and SDS-PAGE in the second dimension. Equal amounts of protein were loaded. ADF, cofilin, and actin were identified by Western blot analysis. Amounts of radioactivity in each spot were quantified using a PhosphorImager and ImageQuant or Photometrics Imaging software. Incorporation of radiolabeled amino acids into ADF, cofilin, and actin were compared with those of 19 randomly selected proteins at each time point.
Wild-type C2 cells and four different clones of C2 cells
transfected with the human sm-actin gene were shown to
express different levels of
sm-actin (Fig.
1A). The single mutation in actin alters its
mobility on SDS-PAGE so that it can be identified on one-dimensional gels (2). These same clones were analyzed for ADF protein by Western
blotting (Fig. 1A). Two forms of ADF can be visualized on
these one-dimensional blots, and both decrease with increasing expression of the transfected gene. The upper species is the
phosphorylated form of ADF (45).
Northern blots of total RNA, extracted from these same
sm-actin expressing cell lines, show a decline in ADF
mRNA levels (Fig. 1B). Quantitative analysis of both
Northern and Western blots shows that there is a strong inverse
relationship between both the relative amounts of ADF mRNA and ADF
protein and the amount of
sm-actin expressed in each
clonal line (Fig. 1C).
To determine if this type of regulatory response is specific to ADF or
if other actin-binding proteins behave in an identical manner, we
examined these cell lines transfected with the human sm-actin gene for expression of cofilin, profilin, CapG,
and tropomodulin by Western blotting. Cofilin expression remained constant with increasing
sm-actin expression (Fig.
2) demonstrating that the feedback regulatory signals
differentially impact on the expression of these two structurally and
functionally similar actin assembly regulatory proteins. Expression of
profilin, CapG, and tropomodulin also remained constant with increasing
sm-actin expression (Fig. 2). The mutant
sm-actin is therefore highly specific in its effect on
actin binding proteins.
Possible Mechanisms for the Observed Changes in ADF Expression
To determine if there is a relationship between the
mechanisms responsible for changes in ADF expression that occur in the sm-actin expressing clones and those that alter
endogenous actin gene expression and/or cell morphology, we examined
clonal cell lines of C2 cells transfected with the normal human
-actin gene. Previous studies have revealed a remarkable similarity
between the impact of
sm-actin and
-actin gene
transfections in the C2 cells. Both genes lead to down-regulation of
the endogenous
- and
-actin genes and a similar decrease in cell
surface area (6, 31). In addition, both genes produce down-regulation of the tropomyosin isoforms Tm2,3 (10), and the focal adhesion proteins
vinculin and talin (11). No significant difference in the level of ADF
expression (as a percentage of total protein) was found in either the
high or low expressing clones resulting from transfection of the human
-actin gene (Fig. 3). This demonstrates that neither
morphology nor endogenous actin gene expression was directly coupled to
the ADF down-regulatory response. The down-regulation of ADF was not
due to intrinsic activity of an exogenous
-actin promoter. Clones
expressing mRNA encoding a normal human
-actin at levels
comparable with those of the highest
sm clone (6) did
not impact on ADF expression (Fig. 3).
To examine the level at which ADF expression was down-regulated, we
compared the transcriptional activity of ADF, actin, and 18 S RNA
genes in wild type C2 cells with that in the highest sm-actin-expressing clone (clone 522) by nuclear run-on
assays (Fig. 4). Normalizing the transcriptional
activity to 18 S RNA, the ratio of expression (522 cells/wild type)
for ADF is 1.6, and for actin it is 0.56. Neither of these ratios is
significantly different from 1, but the ADF ratio is significantly
different from 0.2, the value expected if ADF expression is totally
regulated at the level of transcription, since the ADF mRNA level
in the 522 cells is about 20% of that in wild type C2 cells. These
results suggest that ADF down-regulation is controlled
posttranscriptionally.
ADF Recognizes and Binds to Mutant
It seemed possible that ADF expression could be
down-regulated by the decreased levels of normal actin expressed by the
cell. Indeed, the level of normal -actin closely parallels the level of ADF in these cells. ADF might not recognize the
sm-actin pool, and thus, the level of ADF could be
regulated by the size of the pool of normal actin. To determine if ADF
would recognize and bind to the
sm-actin, we used two
approaches. First we lysed the C2 cell line expressing the highest
level of
sm-actin (clone 522) into immunoprecipitation
buffer and immunoprecipitated the ADF and associated proteins with a
rabbit ADF antiserum. The immunoprecipitates were washed, solubilized
in SDS, and the actin isoforms associated with the ADF were identified
by immunoblotting using a monoclonal antibody that recognizes all of
the actin isoforms (Fig. 5A). For the second
approach, we used an ADF affinity resin to bind actin in lysates of the
cell line expressing the highest level of
sm-actin. The
bound actin was extracted with SDS, and isoforms were identified by
immunoblotting (Fig. 5B). Both methods demonstrate that ADF
binds the
sm-actin in the same ratio to total actin as
occurs in the whole cell lysate.
ADF Activity and Expression Are Perturbed by Increasing the Actin Monomer Pool
Since the sm-actin mutation does not
affect ADF binding, it seemed likely that the levels of ADF might
depend upon the utilization of actin by the cell and be regulated by
signals dependent upon the monomer or filamentous actin pools. To
assess the distribution of actin isoforms between the soluble and
cytoskeletal pool, 170,000 × g supernatant and pellet
fractions were prepared from extracts of C2 cells and three of the
sm-actin expressing cell lines used above. The
distribution of
-actin,
sm-actin, and
-actin in the supernatant and pellet fractions were determined from Western blots, using a polyclonal antibody specific for the
-actin isoform and a monoclonal antibody specific for the
-actin and the aberrantly migrating
sm-actin isoforms (Fig. 1A). The
results of this analysis (Table I) show that in cells
expressing more of the
sm-actin isoform, a higher
proportion of the total actin exists in the supernatant
fraction. Neither of the highest expressing clones transfected with the
wild-type human
-actin or
-actin showed any significant
difference from C2 cells in the distribution of actin between the
soluble and particulate pool (data not shown). By assaying cell
extracts for G-actin using the DNase I inhibition assay (Table I), we
also confirmed that cell lines expressing a higher amount of the
sm-actin contained a higher amount of unassembled actin.
The increase in G-actin measured by the DNase I assay is not as large
as the increase observed in the supernatant actin pool, perhaps
indicating that some of the increase in non-sedimentable actin results
from aggregates or oligomers that do not inhibit DNase I
stoichiometrically. The percentage increase in actin in the soluble
pool roughly parallels the decline observed in ADF expression,
suggesting that ADF expression may be sensitive to feedback regulation
as a result of increased monomer and/or decreased polymer.
|
To explore this hypothesis further, we examined the effect of the
cell-permeable actin depolymerizing agent, Lat A, on the synthesis of
ADF. By binding to monomeric actin (47), Lat A increases monomeric
actin pools in Swiss 3T3 and HeLa cells, a change which is accompanied
by a decrease in actin synthesis (8). C2 cells rounded up within 20 min
of Lat A addition to 5 µM. Unassembled actin in Triton
X-100-soluble extracts of C2 cells treated with this same concentration
of Lat A increased from 34 ± 3% of total actin to 58 ± 4%
(constant from 6 to 18 h after Lat A addition), a shift comparable
with that observed previously (8). Over the 18-h time course of Lat A
treatment, ADF and cofilin did not change significantly as a percent of
total protein (not shown). However, even within 6 h of Lat A
treatment, nearly all of the ADF and cofilin became phosphorylated
(Fig. 6). The phosphorylated forms of ADF and cofilin
are inactive in binding actin (45, 48). Thus, the immediate response of
C2 cells to an increase in unassembled actin is to inactivate both ADF
and cofilin.
The amounts of radioactivity incorporated into ADF, cofilin, and actin
by pulse labeling before and after Lat A treatment were determined by
PhosphorImaging and compared with 19 randomly selected protein spots.
Exposure of C2 cells to Lat A for greater than 12 h caused a
significant decline in overall protein synthesis. As shown in Fig.
7, cofilin synthesis closely follows the average levels
of protein synthesis during the first 12 h. However, ADF and actin
synthesis decrease much more rapidly following Lat A addition,
paralleling the increase in unassembled actin in Lat A-treated cells.
These results demonstrate that the expression of ADF, but not cofilin,
is sensitive to the utilization of actin by the cell.
The ability of an actin mutant to specifically down-regulate ADF
suggests the existence of a unique regulatory pathway linking ADF with
an unknown aspect of actin function. The specificity of this pathway is
highlighted by the finding that sm- and
-actin transfections parallel each other in all aspects except ADF regulation (6, 10, 11, 31). We doubt, however, that ADF is responding to the
decreased levels of normal actin per se because the binding studies suggest that ADF can equally bind normal and
sm-actin. Unlike
-actin, the mutant
sm-actin cannot form normal actin filaments but rather
assembles into ribbon-like structures (22). ADF regulation may
therefore be responding to either the decrease in F-actin available for
binding or to the increase in the G-actin pool. The ability of Lat A to
induce decreased synthesis of ADF but not cofilin confirms that the
G-/F-actin ratio can certainly regulate ADF metabolism.
The difference in regulation of expression of ADF and cofilin is of
considerable interest because they are members of the same family of
18.5 kDa, calcium-independent, pH-sensitive
F-actin-binding/depolymerizing and G-actin-sequestering proteins (26,
27, 40, 49-52). Both proteins have identical regulatory sites for
phosphorylation (48, 53) and have been identified as proteins that
undergo rapid dephosphorylation in response to external stimuli which
result in changes in cytoskeletal organization and assembly (54-57).
The results presented here demonstrate that both proteins respond identically to posttranslational regulation in response to increased monomeric actin pools. This is consistent with a model in which elevated G-actin initially inactivates both ADF and cofilin by phosphorylation. If this G-actin elevation persists, a second level of
regulation, decreased synthesis, comes into play only for ADF. In
addition, both ADF and cofilin contain identical nuclear localization
signals (58-60) that serve to target these proteins to the nucleus
(with cytoplasmic actin), especially in times of stress (61, 62). The
nuclear translocation of cofilin in T-cells is strictly regulated by
co-stimulatory signals that stimulate both clonal growth and expression
of the cell's functional repertoire, including the production of
interleukin-2 (54). Inhibiting this translocation, albeit indirectly
through blocking dephosphorylation of cofilin, leads to enhanced
apoptosis (63). Therefore, these proteins would be ideal candidates for
providing a nuclear signal for the assembly state of actin in the
cytoplasm, a likely first step in the process of maintaining actin
homeostasis. The difference in their response to
sm-actin suggests that if this is one of their
functions, then they are monitoring different properties of actin
metabolism.
ADF and cofilin are also differentially regulated during myogenesis.
ADF is down-regulated in developing muscle in vivo (38, 45)
when the muscle-specific -actin isoform is synthesized and G-actin
levels decline (64). However, during in vitro myogenesis (45), ADF levels do not decline, but rather the ADF is inactivated by
phosphorylation. These results suggest that control of ADF activity is
necessary for regulating actin assembly in muscle cells and that
extracellular signals, provided by the in vivo environment,
are necessary for repression of ADF synthesis. Cofilin levels also
decline in developing muscle but to a lesser extent than ADF (39), due
in part to the expression of a muscle cofilin isoform (65). Levels of
cofilin, but not ADF, are found to increase in denervated muscle (66)
and in dystrophic muscle (67), demonstrating that differential
up-regulation of these proteins can also occur. Together with the data
presented in this paper, this work suggests that ADF and cofilin are
independently regulated although they are usually co-expressed. This
suggests different functional requirements for these two gene
products.
Actin appears to be unique in its ability to regulate the expression of
other gene products associated with microfilament function. Both wild
type and sm-actin gene transfections can change both the
protein and mRNA levels of endogenous actins, selective
tropomyosins, vinculin, talin, and now ADF (6, 10, 11, 31). In
contrast, transfection of gene constructs encoding vinculin,
-actinin, Tm-1, and gelsolin, which all impact on cell morphology,
do not affect the expression of other microfilament associated products
(12, 13, 15, 16). This therefore raises the question of whether actin
expression is a master regulator of other microfilament components and
can send signals to, but does not respond to, the expression of these
other gene products. Recent studies in which the single ADF/cofilin
gene product in Dictyostelium discoidium was overexpressed
suggest that this is not true (68). A 7-fold increase was achieved in
Dictyostelium cofilin expression, but this was accompanied
by a 3-fold increase in actin expression, suggesting that for certain
actin-binding proteins, especially those that are involved in setting
or maintaining the monomeric actin pool, compensatory changes in actin
synthesis are triggered. This suggests that it is direct alteration in
actin metabolism that may regulate actin and other actin-binding
proteins.
What then is the biological function of ADF? These data certainly do not support a role as a simple actin monomer sequestering protein. Its biosynthetic regulation is consistent with what one would expect for a protein whose function is to set and maintain the G-/F-actin ratio. Indeed, it is the proteins of the ADF/cofilin family that are necessary to turn over the actin in the tails of Listeria monocytogenes, the intracellular bacterium that utilizes actin assembly to propel itself around the cytoplasm of infected cells (69). We propose that through both posttranslational and biosynthetic regulation, ADF plays a role in regulating actin polymer levels in C2 cells.