1 Department of Oral Biology, College of Dentistry, University of Nebraska
Medical Center, Lincoln, NE 68583, USA
2 University of Minnesota Cancer Center, Minneapolis, MN 55455, USA
3 Department of Laboratory Medicine and Pathology, Minneapolis, MN 55455,
USA
4 Department of Orthopaedic Surgery, Minneapolis, MN 55455, USA
5 Eppley Institute for Research in Cancer and Allied Diseases, Department of
Biochemistry and Molecular Biology, and Department of Pathology and
Microbiology, University of Nebraska Medical Center, Omaha, NE 68198,
USA
* Author for correspondence (e-mail: weste047{at}umn.edu)
Accepted 23 January 2003
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Summary |
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Key words: Formin homology, FHOD1, Actin cytoskeleton, Stress fibers, Migration, Cell motility, FHOS
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Introduction |
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A significant advance in our understanding of cytoskeletal dynamics is the
realization that members of the formin (Fmn)/diaphanous (Dia) family of
proteins [here collectively referred to as formin-homology (FH) proteins] have
crucial roles in actin cytoskeleton reorganization. FH proteins are
evolutionarily conserved and regulate cytokinesis and cell polarity in
Drosophila melanogaster, Saccharomyces cerevisiae, Schizosaccharomyces
pombe and Aspergillus nidulans
(Afshar et al., 2000;
Castrillon and Wasserman, 1994
;
Chang et al., 1997
;
Harris et al., 1997
;
Kohno et al., 1996
). They also
regulate conjugation in S. pombe
(Petersen et al., 1995
) and
axial and bud site selection in S. cerevisiae
(Evangelista et al., 1997
;
Imamura et al., 1997
;
Zahner et al., 1996
).
Recently, it was demonstrated that FH proteins in S. cerevisiae
induce actin cable formation in an Arp2/3-independent manner prior to
tropomyosin stabilization (Evangelista et
al., 2002
; Sagot et al.,
2002
) and are nucleators of unbranched and polarized actin
filaments (Pruyne et al.,
2002
).
In mammalian cells, FH proteins regulate cytoskeletal organization,
embryonic patterning, cell survival, migration and gene transcription
(Habas et al., 2001;
Ishizaki et al., 2001
;
Jackson-Grusby et al., 1992
;
Sotiropoulos et al., 1999
;
Tominaga et al., 2000
;
Trumpp et al., 1992
;
Watanabe et al., 1997
;
Watanabe et al., 1999
;
Westendorf, 2001
;
Yayoshi-Yamamoto et al.,
2000
). Fmn mutations cause limb deformities and renal
failure in mice (Maas et al.,
1990
; Woychik et al.,
1985
; Woychik et al.,
1990
; Wynshaw-Boris et al.,
1997
). In humans, Dia mutations are linked with autosomal
dominant deafness (Dia1) and premature ovarian failure
(Dia2) (Bione et al.,
1998
; Lynch et al.,
1997
). Increased Dia2 expression is also directly
associated with the metastatic potential of rat osteosarcoma cells
(Fukuda et al., 1999
). Thus,
deregulated FH protein activity in mammalian cells causes developmental
defects and might contribute to cancer progression.
FH proteins regulate these diverse processes by interacting with actin
binding proteins and bridging Rho-family GTPases to Src tyrosine kinase and
Wnt signaling pathways (Evangelista et al.,
1997; Habas et al.,
2001
; Ishizaki et al.,
2001
; Kohno et al.,
1996
; Tominaga et al.,
2000
; Watanabe et al.,
1997
; Watanabe et al.,
1999
; Yayoshi-Yamamoto et al.,
2000
). Increasing evidence suggests that FH proteins play a
crucial role in cell motility processes. A truncated, active form of the RhoA
effector mDia1 is sufficient to restore force-induced focal contact formation
in cells containing inactive RhoA, thereby suggesting that mDia is crucial for
cell contraction (Riveline et al.,
2001
). Expression of a truncated version of mDia1 containing the
Rho-binding domain inhibits the directed migration of CHO cells in a wound
closure assay (Krebs et al.,
2001
). Similarly, expression of a truncated form of formin-related
gene in leukocytes (FRL) containing the Rac1-binding domain inhibits cell
spreading and chemokine-induced migration of an FRL-expressing macrophage cell
line by blocking cell adhesion to fibronectin
(Yayoshi-Yamamoto et al.,
2000
). Thus, FH proteins appear to regulate several stages of cell
migration.
FHOD1 (formin-homology-2-domain-containing protein) is an FH protein that
was previously described as FHOS
(Westendorf, 2001;
Westendorf et al., 1999
). Its
designation was changed to meet the Guidelines for Human Gene Nomenclature.
FHOD1 is a characteristic FH family member in that it contains a GTPase
binding domain (GBD), FH1 and FH2 domains, a coiled coil, and a
Diaphanous-like autoregulatory domain (DAD)
(Alberts, 2000
;
Wasserman, 1998
). FHOD1
interacts with Rac1 in a guanine-nucleotide-independent manner
(Westendorf, 2001
).
Self-interactions between the N- and C-termini prevent full-length FHOD1 from
activating gene transcription from the serum response element (SRE). However,
deletion of the N- or C-termini allows FHOD1 to induce SRE transcription
(Westendorf, 2001
). The
induction of the SRE by the C-terminal FHOD1 mutant, FHOD1
C, was
independent of Rac1 (Westendorf,
2001
). The goals of this study were to determine the effects of
FHOD1 and FHOD1 C-terminal truncation mutations on cell behavior and actin
cytoskeleton organization. FHOD1 and FHOD1
C co-localized with F-actin
at the cell periphery and in newly formed stress fibers, respectively. FHOD1
expression also induced cell elongation and enhanced the migration of WM35
melanoma cells and NIH-3T3 fibroblasts to extracellular matrix components
without affecting integrin usage. These data show that expression of a
full-length FH protein affects cell motility and suggest that FH proteins are
crucial regulators of cell migration.
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Materials and Methods |
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FHOD1-N rabbit antiserum was generated by Rockland Laboratories
(Gilbertsville, PA) using glutathione-S-transferase (GST)-FHOD1
(1-328) fusion protein as an immunogen. FHOD1-C mouse antiserum have been
described previously (Westendorf et al.,
1999).
Generation of stable cells lines
WM35 and NIH-3T3 cells were transduced twice within 24 hours with
retroviral supernatants in the presence of 8 µg ml-1 polybrene.
eGFP expression was monitored by immunofluorescence microscopy. 2-3 days after
the last transduction, eGFP-positive cells were selected by
fluorescence-activated cell sorting on a FACSVantage flow cytometer (Becton
Dickinson, Mountain View, CA). WM35 cells were maintained in DMEM supplemented
with 10% fetal bovine serum (Biowhittaker, East Rutherford, NJ), 2 mM
L-glutamine, 50 U ml-1 penicillin and 50 µg ml-1
streptomycin. NIH-3T3 cells were maintained in DMEM supplemented with 10%
bovine calf serum (Biowhittaker), 2 mM L-glutamine, 50 U ml-1
penicillin and 50 µg ml-1 streptomycin. FHOD1 and eGFP levels
were regularly monitored with immunoblotting and microscopy, respectively.
Robert Kerbel (University of Toronto, Canada) generously provided the WM35
melanoma cells.
Cell measurements
Digital phase contrast images were imported into NIH-Image, where cell
lengths and widths were measured in arbitrary units. Length-to-width ratios
were calculated for each cell. Isolated cells (20-34) in a minimum of three
fields were measured for each cell line. Statistical significance was
determined with a one-way analysis of variance.
In situ immunofluorescence analysis
NIH-3T3 cells were grown on coverslips and transfected with expression
vectors encoding HA-tagged wild-type or truncated FHOD1 proteins. Cells were
washed with phosphate buffered saline (PBS) 24 hours after transfection and
fixed with 4% paraformaldehyde for 5 minutes at room temperature. Cells were
washed three times with PBS prior to blocking and permeabilization with 0.2%
Triton X-100 in 5% normal goat serum in PBS for 30 minutes. For assays of
F-actin and FHOD1 co-localization, coverslips were sequentially incubated for
30 minutes with phalloidin-rhodamine (1 U per coverslip; Molecular Probes,
Eugene, OR) followed by a 1 hour incubation with rabbit anti-HA antibody
(1:400, Santa Cruz Biotechnology, Santa Cruz, CA) and 1 hour incubation with
fluorescein isothiocyanate (FITC)-conjugated goat anti-rabbit antibody (1:400;
Jackson ImmunoResearch Laboratories, West Grove, PA).
For RacN17 experiments, NIH-3T3 cells were transiently transfected with
pCMV5-FHOD1 C and pCMV-T7-Rac N17. 24 hours after transfection, cells
were washed in PBS and fixed with 4% paraformaldehyde (pH 7.1) in PBS for 5
minutes at room temperature prior to washing and blocking in 5% normal goat
serum in PBS. For Rho pathway inhibition experiments, cells were exposed for
18 hours to C3 transferase (Rho inhibitor) or for 10 hours to Y-27632 (ROCK
inhibitor). Cells were washed in cytoskeleton buffer (CB; 10 mM MES, 150 mM
NaCl, 5 mM EGTA, 5 mM MgCl2, 5 mM glucose, pH 6.1) and fixed in 3%
paraformaldehyde in CB for 10 minutes at room temperature. Cell were
subsequently washed with CB, 0.1% Triton X-100 in CB for 1 minute prior to
washing and blocking in 5% normal goat serum in PBS. For assays of F-actin and
FHOD1 co-localization, coverslips were sequentially incubated for 30 minutes
with phalloidinrhodamine (1 U per coverslip; Molecular Probes) in PBS,
followed by a 1 hour incubation with rabbit anti-HA antibody (1:400; Santa
Cruz) and a 1 hour incubation with FITC-conjugated goat anti-rabbit antibody
(1:400; Jackson ImmunoResearch Laboratories). Between and after antibody
incubations, coverslips were washed in PBS and then mounted on glass slides in
Vectashield mounting medium (Vector Laboratories, Burlingame, CA) and
visualized at 400x or 600x using a Nikon confocal microscope. For
assays of FHOD1
C, RacN17 and F-actin co-localization, the above
protocols were followed except that incubation with anti-HA antibody was
followed by washing and a 1 hour incubation with mouse anti-T7 antibody
(1:1000; Novagen, Madison, WI). Secondary antibodies were Cy5-labeled donkey
anti-rabbit and FITC-labeled goat anti-mouse antibodies (Jackson
ImmunoResearch Laboratories).
Immunoprecipitations
HEK293T cells were transfected with HA-tagged FHOD1 plasmids or empty
vector (pCMV5) by calcium phosphate precipitation. 24 hours after
transfection, the plates were placed on liquid nitrogen for 1 minute and 1 ml
lysis buffer (50 mM HEPES pH 7.8, 1% Triton X-100, 1 mM EDTA, 30 mM sodium
pyrophosphate, 1 mM sodium vanadate, 10 mM sodium fluoride, 1 µM
phenylmethylsulfonyl fluoride, 20 ng ml-1 aprotinin) was added.
Lysates were vortexed and centrifuged to pellet detergent-insoluble cell
debris. Protein concentrations of lysates were determined using the BCA
Protein Assay (Pierce, Rockford, IL). Equal amounts of protein were incubated
for 2 hours with gentle rocking at 4°C with antibodies specific for HA
(Covance, Berkeley, CA) or actin (Santa Cruz). Protein-A/Sepharose beads
(Amersham Biosciences, Inc., Piscataway, NJ) were added for 20 minutes with
gentle rocking at 4°C. Immune complexes were collected by centrifugation,
washed twice with lysis buffer supplemented with 0.1% sodium dodecyl sulfate
and then washed twice with 50 mM HEPES prior to the addition of 1x
sample buffer. Proteins from immunoprecipitations or whole cell extracts were
resolved by 8-12% SDS-PAGE, electrophoretically transferred to
polyvinyldifluoride (PVDF; Schleicher & Schuell, Keene, NH) or
nitrocellulose membranes (Amersham Pharmacia Biotech) and immunoblotted with
primary antibodies for HA or actin, followed by the appropriate secondary
antibodies. Proteins on PVDF membranes were detected using
alkaline-phosphatase-conjugated secondary antibodies in a colorimetric
reaction. Nitrocellulose membranes were developed with
horseradish-peroxidase-conjugated secondary antibodies using the enhanced
chemiluminescence system (Amersham Pharmacia Biotech).
Flow cytometry
Transduced WM35 cells were detached with 3 mM EDTA in PBS and washed in
fluorescence buffer [1x PBS, 2.5% bovine calf serum (BCS)]. Cells
(3x105) were incubated with 5 µg ml-1 of the
indicated primary monoclonal antibody (mAb) for 30 minutes on ice. The cells
were washed once with fluorescence buffer and then incubated for 30 minutes on
ice with 5 µg ml-1 secondary antibodies conjugated to
phycoerytherin or Cy5 (Jackson ImmunoResearch Laboratories). After two washes,
the cells were resuspended in 1% paraformaldehyde and analyzed on FACScalibur
flow cytometer. Mean fluorescence intensities of eGFP positive cells were
determined using CELLQuest software (Becton Dickinson). Antibodies recognizing
integrins 1,
3,
4,
ß4 and
vß3 were purchased from Chemicon
(Temecula, CA). Anti-
2 and 9EG7 ß1 mAbs were purchased
from Pharmingen (San Diego, CA). Tucker LeBien and Yoji Shimizu (University of
Minnesota, Minneapolis, MN) kindly provided the P5D2 and 15/7ß1 Abs,
respectively. Control mouse and rat antibodies were obtained from Cappell
(Cochranville, PA).
Cell adhesion assays
Immulon 1B 96-well plates (Dynex Technologies, Chantilly, VA) were coated
overnight on ice with 100 µl of rat-tail type-I collagen (Becton Dickinson)
at the indicated concentrations. Wells were blocked with PBS containing 3%
(w/v) bovine serum albumin (BSA) for 1 hour at 37°C. Cells were released
with 3 mM EDTA, washed twice in serum-free medium containing 20 mM HEPES, pH
7.1 (SFM/HEPES) and 1% BSA (w/v) and then resuspended to
1x105 cells ml-1 in the same medium. For
integrin-blocking assays, cells were suspended in SFM/HEPES to
1x106 cells ml-1, pre-incubated with 2.5 µg
ml-1 of the indicated mAb for 15 minutes at 37°C and then
diluted to 1x105 cells ml-1. Cells
(1x104 in 100 µl) were added to the plates and allowed to
adhere to the wells for 20 minutes at 37°C. Wells were washed twice with
SFM/HEPES containing 1% BSA and then twice with SFM/HEPES. Growth medium [DMEM
containing 10% fetal bovine serum (FBS); 100 µl] was added to the wells.
PMS/MTS solution (CellTiter96 Aqueous Non-radioactive Cell Proliferation
Assay, Promega, Madison, WI) was then added according to the manufacturer's
instructions (20 µl per well). Cells were incubated at 37°C for 3-4
hours. Absorbance at 490 nm was measured with an ELISA plate reader. The
number of adherent cells was determined by normalizing to a standard curve
generated with increasing volumes of the starting cell dilutions. Values
represent the mean of triplicate samples.
Cell migration assays
Migration assays were performed in a modified Boyden chamber (NeuroProbe,
Gaithersburg, MD). Cells were detached with 3 mM EDTA in PBS and washed in
SFM/HEPES. Ligands [rat-tail type-I collagen, mouse type-IV collagen
(Invitrogen), purified human plasma fibronectin, mouse laminin (Invitrogen) or
BSA (Sigma)] were diluted in SFM/HEPES and added (33 µl) to the lower
wells. SFM/HEPES was placed in control wells. Cells (2.2x104
in 55 µl) were placed in the upper wells and separated from the lower wells
by polycarbonate filter with 8 µm pores (Osmonics, Minnetonka, MN). For
antibody inhibition studies, cells were pretreated with 2.5 µg
ml-1 of the indicated integrin or isotype-matched control antibody
(in SFM/HEPES) for 15 minutes at 37°C with rotation prior to placement in
the migration chamber. Antibodies to ß1 integrin were added at
1 µg ml-1 as this was determined to be optimal in titration
experiments (data not shown). Migration assays were incubated at 37°C for
4-5 hours. Filters were fixed and stained with Diff-Quik solutions (Dade
Behring, Newark, DE), and unmigrated cells were wiped from the filters.
Filters were mounted on glass slides in immersion oil, and migrated cells in
five random fields were counted at 400x magnification.
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Results |
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An alteration in cellular morphology was observed in eGFP-positive (i.e. post-sort) populations expressing full-length FHOD1. NIH-3T3 (Fig. 2A) and WM35 (data not shown) cells that expressed FHOD1 were elongated in both confluent and subconfluent conditions. These observations were quantified by measuring the lengths and widths of subconfluent eGFP-positive NIH-3T3 cells in phase contrast images using NIH-Image software. The length-to-width ratio was calculated for each cell (Fig. 2B). As would be expected for unsynchronized cell populations, wide ranges were observed. Cells transduced with viruses expressing just eGFP (MSCV control) or truncated FHOD1 (1-421) had a mean length-to-width ratios of 13.7 and 13.1, respectively. The median value for each of these populations was 10.6. Cells expressing full-length FHOD1 displayed larger range of values with significantly different means of 20.2 (P<0.05). Thus, FHOD1-expressing cell populations contained elongated cells.
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FHOD1 proteins co-localize with F-actin structures
Cell shape is regulated by the actin cytoskeleton. To determine whether
FHOD1 proteins interact with and/or affect the actin cytoskeleton, NIH-3T3
cells were transiently transfected with FHOD1 expression constructs. In situ
immunofluorescence microscopy revealed that full-length FHOD1 was expressed in
perinuclear regions and at cell peripheries
(Fig. 3A), which is consistent
with previously reported results
(Westendorf, 2001).
FHOD1-expressing cells had noticeably more bundled F-actin, as detected with
rhodamine-conjugated phalloidin and confocal microscopy, than adjacent
nontransfected cells. FHOD1 co-localized with F-actin at the cell periphery;
however, FHOD1 did not co-localize with F-actin in perinuclear regions.
Moreover, dense F-actin structures away from the membrane did not appear to
contain FHOD1. These data indicate that FHOD1 induces or stabilizes F-actin
formation but that interactions between FHOD1 and F-actin are only present at
the cell periphery.
|
We previously demonstrated that N- and C-terminal domains in FHOD1 mediate
an intramolecular interaction(s)
(Westendorf, 2001). Deletion
of the self-interaction domains creates active FHOD1 proteins that stimulate
transcription via alternative signaling pathways. Differential cellular
localization of the FHOD1 mutants appears to be responsible for these effects
(Westendorf, 2001
). To expand
these studies, we examined how FHOD1 truncation mutants interact with the
actin cytoskeleton. Cells expressing FHOD1
C demonstrated markedly
thick stress fibers. FHOD1
C co-localized with the stress fibers
(Fig. 3A,
Fig. 4B). Cells expressing
FHOD1 (1-421) did not display stress fibers but, like cells containing
full-length FHOD1, appeared to have more F-actin than untransfected cells in
the same field. FHOD1 (1-421) also co-localized with F-actin at the cell
membrane and in the perinuclear region. These results indicate that the
N-terminus of FHOD1 is sufficient for localization to the membrane and
perinucleus. These results demonstrate that an activated form of FHOD1 induces
or stabilizes stress fiber formation.
|
To determine whether FHOD1 proteins also interact with globular (G) actin,
HEK293T cells were transiently transfected with full-length or truncated
HA-FHOD1 proteins and assayed by immunoprecipitation and immunoblotting
analysis. Full-length FHOD1 co-immunoprecipitated with Triton X-100 soluble
actin (Fig. 3B). However, the
truncation mutants, FHOD1 (1-421), [1-1010 (C)] and (469-1165), did not
interact with G-actin. These results suggest that FHOD1 associates with
G-actin and that multiple regions of FHOD1 contribute to the interaction.
FHOD1-C-dependent stress fibers require Rac1, RhoA and ROCK
activities
We previously demonstrated that FHOD1 interacts with Rac1 but not RhoA
(Westendorf, 2001). To
determine whether Rac1 is required for FHOD1-
C-dependent stress fiber
formation, we transiently co-transfected NIH-3T3 cells with FHOD1
C and
dominant-negative RacN17. Actin stress fiber organization in cells expressing
both FHOD1
C and RacN17 was similar to neighboring, non-transfected
cells that displayed normal actin-rich lamellipodia
(Fig. 4A). Thus, RacN17
prevented the appearance of stress fibers in FHOD1
C-expressing cells.
These data suggest that FHOD1 is upstream of Rac1.
We also tested the effects of RhoA and ROCK inhibitors, C3 transferase and
Y-27632, on FHOD1-dependent stress fibers. Stress fiber formation requires
RhoA and its downstream effector ROCK
(Amano et al., 1997;
Leung et al., 1996
;
Ridley and Hall, 1992
). C3
transferase (Rho inhibitor; Fig.
4B) and Y-27632 (ROCK inhibitor;
Fig. 4C) inhibited
FHOD1-
C-dependent stress fibers. These inhibitors induced a phenotype
seen with Rho inhibition by RhoGAP (Tatsis
et al., 1998
; Vincent and
Settleman, 1999
). Together these data demonstrate that
FHOD1-dependent stress fibers required Rac1, RhoA and ROCK enzymatic
activities.
FHOD1 enhances cell migration without affecting adhesion, integrin
usage or integrin activation
The altered morphology of FHOD1-expressing cells suggested that FHOD1 might
affect cell adhesion and/or motility. To determine whether FHOD1 affects cell
adhesion, transduced WM35 cells were allowed to adhere to various
concentrations of type-I collagen. The number of adherent cells increased in a
type-I collagen concentration-dependent manner and peaked at 15 µg
ml-1 (Fig. 5A).
FHOD1-expressing cells bound to the substrate similar to control (empty pMSCV
vector) and FHOD1 (1-421)-positive cells. Likewise, the adherence of
FHOD1-positive and control NIH-3T3 cells to fibronectin was comparable
(Fig. 5B). These data indicate
that FHOD1 does not affect the overall adherence of cells to these
extracellular matrix proteins.
|
Although cell adhesion was not affected by FHOD1 expression, motility towards extracellular matrix proteins was significantly enhanced in both WM35 and NIH-3T3. An approximate two- to threefold increase in the migration of FHOD1-positive WM35 melanoma cells to type-I collagen was observed (Fig. 6A). Migration was concentration dependent, but FHOD1-positive cells had increased migration at each concentration relative to control (MSCV) and FHOD1 (1-421)-expressing cells. WM35 cells did not migrate to BSA or fibronectin (data not shown).
|
FHOD1-positive NIH-3T3 cells also displayed increased migration to extracellular matrix proteins. FHOD1-expressing cells had increased migration to fibronectin and laminin compared with control (MSCV) and FHOD1 (1-421)-expressing cells (Fig. 6B). Similar results were observed when 10% BCS, which contains fibronectin and laminin along with other stimulatory molecules, was used as a migratory signal. None of the NIH-3T3 cells migrated in response to BSA (Fig. 6B), or type-I or type-IV collagen (data not shown). Together, these data indicate that FHOD1 expression enhances cell migration in both fibroblasts and melanoma cells.
To begin to understand the mechanism of FHOD1-enhanced cell migration, we
examined integrin expression levels and the adhesion properties of
FHOD1-expressing and control WM35 melanoma cell lines. WM35 cells expressed
1,
2,
3,
4, ß1 and
vß3 at varying levels. FHOD1-positive cells
had slightly elevated integrin levels as compared to control (MSCV) and FHOD1
(1-421)-expressing cells (Fig.
7A) but the mean fluorescence intensities varied by less than
10%.
|
To determine whether FHOD1 expression affected integrin function in
adhesion and migration, we preincubated control (MSCV) and FHOD1-positive WM35
cells with anti-integrin blocking antibodies prior to placing them in adhesion
and migration assays. The 1ß1,
2ß1,
3ß1
and
vß3 integrins mediate binding to type-I
collagen (van der Flier and Sonnenberg,
2001
). These integrin subunits were all expressed in the WM35
cells (Fig. 7A). Blocking
antibodies to ß1 integrin completely inhibited the adhesion
and migration of control (MSCV) and FHOD1-positive cells
(Fig. 7B,C). Blocking
antibodies to
1 and ß2 each partially
blocked adhesion and migration of control (MSCV) and FHOD1-expressing WM35
cells. Antibodies blocking
3
(Fig. 7B,C) and
vß3 (data not shown) did not block the
adhesion or migration of any WM35 cell populations. Thus, WM35 cell adhesion
to type-I collagen is largely mediated by
1ß1 and
2ß1 integrin complexes. Importantly, FHOD1
expression did not alter the ability of these integrins to mediate cell
adhesion or migration. FHOD1 also did not affect the appearance of
activation-dependent ß1 integrin conformation epitopes, 15/7
(Yednock et al., 1995
) or 9EG7
(Lenter et al., 1993
), in WM35
cells (Fig. 8). Divalent
cations (MnCl2) increased these epitopes to similar levels in both
MSCV- and FHOD1-expressing WM35 (data not shown). These data indicate that
FHOD1 does not affect the avidity or affinity of integrins. Thus, FHOD1
probably enhances cell migration by activating events downstream of
integrins.
|
![]() |
Discussion |
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Together with previous reports, our data indicate that FH proteins might
affect multiple phases of cell migration. Truncated active forms of mDial that
lack the N-terminal RhoA-binding domain restore force-induced focal contact
formation in cells in which RhoA is inactivated, suggesting that mDia is
crucial for cell contraction and acts downstream of RhoA
(Riveline et al., 2001). Krebs
and colleagues demonstrated that expression of a truncated version of mDial
containing the FH3 domain and GBD inhibits the directed cell migration of CHO
cells in a wound closure assay (Krebs et
al., 2001
). Third, a truncated form of FRL containing the FH3 and
Rac1-binding domains inhibits cell spreading and chemokine-induced migration
of an FRL-expressing macrophage cell line by blocking cell adhesion to
fibronectin (Yayoshi-Yamamoto et al.,
2000
). These latter two reports suggest that the N-termini of FH
proteins are sufficient to interfere with migration, possibly by inhibiting
the respective endogenous FH protein. We did not observe significant
inhibition of migration upon FHOD1 (1-421) expression. However, this fragment
lacks the GBD and thus might not block the appropriate signaling pathways.
One novel aspect of our study is the long-term expression and analysis of a
full-length FH protein. All previous studies examining the roles of FH
proteins in cell migration were performed with fragments of the respective FH
protein that have not been identified in nature. Attempts to overexpress Fmn
and FRL were reportedly unsuccessful, leading the investigators to conclude
that stable expression of these FH proteins is toxic
(Vogt et al., 1993;
Yayoshi-Yamamoto et al.,
2000
). We were able to obtain long-term cultures of
FHOD1-overexpressing NIH-3T3 and WM35 cells. Our results might be dependent on
our choice of expression systems but it is also possible that FHOD1 has a
distinct role and that its overexpression does not place the cells at a
selective disadvantage. Interestingly, our attempts to generate stable FHOD1
C expression were unsuccessful. The predominant stress fibers found in
cells transiently transfected with this truncated and activated FHOD1 protein
probably prohibit long-term survival. These results suggest that individual FH
proteins might perform specialized functions and link the actin cytoskeleton
to different signaling pathways to control cell activities.
FHOD1 was localized to cell membranes and perinuclear regions in both
stable and transient transfectants (Fig.
1C, Fig. 3A). This
staining pattern is similar to that observed for mDial
(Watanabe et al., 1997).
Interestingly, FHOD1 co-localized with F-actin in lamellae but not with
F-actin in the cytoplasm. These results suggest that FHOD1 participates in
filamentous actin polymerization or nucleation at the cell membrane. The
induction of stress fiber formation in cells expressing an activated FHOD1
protein lacking the C-terminal autoregulatory domain has also been observed by
others (Oliver Fackler, personal communication) and indicates that FHOD1 might
have a role in actin polymerization. These results agree well with the recent
findings that FH proteins in S. cerevisiae control cell polarity by
nucleating actin cable assembly independent of Arp2/3 proteins
(Evangelista et al., 2002
;
Pruyne et al., 2002
;
Sagot et al., 2002
). The
nature of FHOD1 interactions with F-actin remains to be determined.
Full-length FHOD1 did interact with G-protein monomers in vivo, but truncated
FHOD1 proteins did not. These data suggest that FHOD1 associates indirectly
with actin and/or that multiple regions of FHOD1 are necessary to interact
with actin or actin-binding proteins. One likely candidate is profilin, which
binds to the proline-rich FH1 domain of other FH proteins
(Chang et al., 1997
;
Evangelista et al., 1997
;
Umikawa et al., 1998
;
Watanabe et al., 1997
;
Yayoshi-Yamamoto et al.,
2000
).
Many studies indicate that FH proteins are involved in actin organization
and the appearance of stress fibers. Truncated and activated versions of
mDia1, a downstream effector of RhoA
(Watanabe et al., 1997),
induce thin actin stress fibers that are subsequently organized into thick
stress fibers by another RhoA effector, ROCK
(Watanabe et al., 1999
). In
this report, we demonstrate that a truncated version of FHOD1 (residues
1-1010) also induces stress fibers. These actin structures were inhibited by a
dominant-negative Rac1 (N17) protein. The ROCK and RhoA inhibitors Y-27632 and
C3 transferase also prevented stress fiber formation. These data suggest that
FHOD1 acts upstream of Rac1, RhoA and ROCK. We previously placed Rac1
downstream of FHOD1 in pathways activating transcription from the SRE
(Westendorf, 2001
). Thus,
there might exist multiple classes of FH proteins, those that act upstream of
Rho family GTPases and those that are Rho GTPase effectors. FHOD1 and Daam1, a
Wnt-activated FH protein that regulates RhoA
(Habas et al., 2001
), are
members of the former class, whereas mDia belongs in the latter. The signals
that activate FHOD1 remain to be identified. However, it is clear that FH
proteins have multiple, complex roles in the organization of actin
structures.
In summary, we found that FHOD1 co-localized with some F-actin structures
at the cell periphery. We also found that long-term FHOD1 expression induced
cell elongation and enhanced migration of fibroblasts and melanomas cells. A
FH gene (a mDia2 homolog) was identified as being overexpressed
differently in highly metastatic rat osteosarcoma cells
(Fukuda et al., 1999). Thus,
increased FH protein expression might play a role in tumor metastasis by
facilitating actin polymerization. Subsequent stabilization of actin filaments
would lead to the formation of contractile bundles that are important for cell
motility (Feldner and Brandt,
2002
). Our data indicate that FHOD1 expression increases migration
without altering integrin expression, affinity or avidity. Future studies will
address the roles of FHOD1 in regulating events downstream of integrins to
better understand its role in cell motility and potentially tumor
metastasis.
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Acknowledgments |
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References |
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