Department of Physiology and Biophysics, College of Medicine, University of Tennessee, Memphis, Tennessee 38163
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The cause of reduced migration ability in polyamine-deficient
cells is not known, but their actin cytoskeleton is clearly abnormal.
We depleted polyamines with -difluoromethylornithine (DFMO) in
migrating cells with or without stimulation by epidermal growth factor
(EGF) and investigated filamentous (F-) actin, monomeric (G-) actin,
and thymosin
4 (T
4), using immunofluorescent confocal microscopy,
DNase assay, and immunoblot analysis. DFMO reduced F-actin in the cell
interior, increased it in the cell cortex, redistributed G-actin, and
increased nuclear staining of T
4. However, DFMO did not affect the
amount of T
4 mRNA. EGF caused a rapid increase in the staining of
F-actin in control cells, but DFMO prevented this response to EGF.
Despite the visible changes shown by immunocytochemistry, statistically
significant changes in the amount of either actin isoform or of total
actin did not occur. We propose that DFMO reduces migration by
interfering with the sequestration of G-actin by T
4 and the
association of F-actin with activated EGF receptors.
epidermal growth factor; -difluoromethylornithine; thymosin
4
messenger ribonucleic acid; monomer sequestration
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
THE ACTIN CYTOSKELETON IS the structural basis of the changes in cell shape required for migration and proliferation in animal cells (27). These processes require rapid cytoskeletal reorganization that is accomplished by a cycle of polymerization and depolymerization of actin monomers (G-actin) and filaments (F-actin) and their reorganization within the cell. The reorganization is carried out by an array of actin binding proteins in response to extracellular and intracellular signals. Epithelial cells typically form broad lamellipodia when migrating. These structures are protrusions of the cell membrane over dense, short actin filaments arranged with their rapidly polymerizing barbed ends toward the cell membrane. The assembly of new attachment sites on the extended edge of lamellipodia provides a purchase on which the cell contents can move forward propelled by force exerted through actin filaments. [For a commentary on this subject, see Heidemann and Buxbaum (13).] Actin filaments are produced by a combination of monomer sequestration, nucleation, filament severing, and the uncapping of filament ends by specific actin binding proteins (15, 20).
Studies with -difluoromethylornithine (DFMO), an irreversible
inhibitor of ornithine decarboxylase (the rate-limiting enzyme of
polyamine biosynthesis), have shown that the polyamines must also be
considered in these processes. Polyamines are essential for cell
proliferation (36), attachment (32), efficient migration in culture
(18), and healing in vivo (38). In IEC-6 cells, a normal rat intestinal
crypt cell (28), polyamine deficiency lowers epidermal growth factor
(EGF) receptor phosphorylation, changes its distribution within the
cell, and inhibits proliferation and migration (17). Inhibitors of
polyamine biosynthesis prevent the accumulation of mRNAs that encode
major cytoskeletal components in mouse splenocytes (16). Despite these
effects, DFMO does not reduce total protein (17) and is not toxic to
cells even at a concentration of 10 mM (unpublished data).
The distribution of F-actin has been described by many investigators. However, the distribution of G-actin has received less attention. In cultured cells, most of the G-actin is sequestered by monomer binding proteins (22). Punctate structures thought to represent transient storage of G-actin have been described in the region behind the lamellipodia (4) and in a ring around the nucleus (11). Extracellular ATP has been shown to induce nuclear accumulation of G-actin (19).
Actin binding proteins regulate the polymerization of actin by severing
and capping actin filaments and by sequestering actin monomers.
Sequestering monomers keeps the monomer level above the critical
concentration for actin filament assembly and provides a reserve that
can be rapidly mobilized where needed for polymerization when cell
migration is stimulated. The -thymosins, a family of highly
conserved 4.9-kDa polypeptides, comprise the bulk of the actin
monomer-sequestering capacity in nonmuscle cells (22). Thymosin
4
(T
4) is the major monomer binding protein in most nonmotile cells
and is present at concentrations near 1 × 10
5 M (25). When bound to
actin, T
4 strongly inhibits nucleotide exchange by blocking its
dissociation (8) and may alter the rate or location of polymerization
through its effect on the actin monomer supply. At the low levels
(<20 µM) found in most nonmotile cells, T
4 sequesters actin
monomers at a ratio of 1:1. At high levels such as those present in
circulating cells (>200 µM), the T
4-actin complex can
incorporate into the actin filament, reducing the sequestering ability
of T
4 (5). Whether the T
4-actin complex may reach high levels in
particular intracellular areas in nonmotile cells is not known. Other
actin monomer binding proteins, profilin in particular, can oppose the
action of T
4, thereby providing a regulating step in the cycle of
polymerization and depolymerization (8). T
4 gene expression has been
demonstrated in various cells in which differentiation is occurring:
mouse embryonic stem cells, neural and cardiovascular cells (10), embryonic brain tissue (6), and NIH/3T3 cells (41).
Previously, we have found significant inhibition of migration and other
responses to EGF in polyamine-deficient IEC-6 cells. These changes were
accompanied by marked alterations in F-actin and EGF receptor
distribution (17). In the present investigation, we show that
cytoskeletal alterations also involve G-actin and T4. We propose
that these alterations interfere with the regulation of actin
polymerization and are responsible, at least in part, for the lowered
migration ability of these cells and their failure to respond normally
to a stimulus by EGF.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cell culture and general experimental plan. The IEC-6 cell line (CRL-1592) was purchased from the American Type Culture Collection (ATCC; Manassas, VA). Medium and other cell culture reagents were obtained from GIBCO (Grand Island, NY). Fetal bovine serum (FBS), dialyzed FBS (dFBS), and all other chemicals and biochemicals were obtained from Sigma (St. Louis, MO), except as noted. The cell stock was maintained in DMEM containing 5% heat-inactivated FBS, 10 µg insulin, and 50 µg/ml gentamicin sulfate (DMEM-FBS) in 90% air-10% CO2. Stock was split once a week at 1:30 and used for no more than four passages. Cells were tested for mycoplasma every 6 mo.
Treatment began with plating and lasted for 4 days. Cells were fed on day 2 and serum was removed on day 3. Treatment groups included controls (DMEM-dFBS), DFMO (DMEM-dFBS plus 5 mM DFMO), and DFMO-putrescine (DMEM-dFBS plus 5 mM DFMO supplemented with 10 µM putrescine). DFMO was kindly provided by the Merrell Dow Research Institute of Marion Merrell Dow (Cincinnati, OH). On day 4, cells were removed from about one-third of the monolayer with a razor blade, the medium was changed to remove cellular debris, and the cells were allowed to migrate for 3 h. EGF (Collaborative Research, Bedford, MA) was added in fresh medium at a concentration of 10 ng/ml 2 and 30 min before the end of the 3-h period. Control medium was changed as well. We have shown previously that 5 mM DFMO reduces intracellular putrescine to undetectable levels in 6 h, spermidine to undetectable levels by day 2, and spermine to 30% by day 4 and that 10 µM putrescine is the optimal dose to maintain DFMO-inhibited cell migration and proliferation at control levels (18).Immunocytochemistry. Cells were plated at 1.2 × 104 cells/cm2 on Matrigel-coated coverslips, treated, wounded, and allowed to migrate for 3 h as above. Matrigel was purchased from Collaborative Research. Cells were fixed with 3.7% formaldehyde, permeabilized with 0.1% Triton X-100, stained for G-actin with Gc globulin (Calbiochem, La Jolla, CA), anti-Gc globulin (Dako, Carpenteria, CA), and donkey anti-rabbit fluorescein-conjugated IgG (Jackson ImmunoResearch, West Grove, PA) for 1 h each, and finally stained for F-actin with rhodamine phalloidin (Molecular Probes, Eugene, OR) for 45 min, mounted, and sealed.
TF- and G-actin assay. F- and G-actin were assayed by DNase inhibition according to Heacock and Bamburg (12) with minor modifications. In this assay, actin filaments are bound by activated myosin to prevent depolymerization, immediately separated from unpolymerized actin by centrifugation, extracted, and depolymerized (if polymerized) for quantitation by DNase inhibition. Briefly, a buffer containing 0.5 M KCl, 50 mM K2PO4, and rabbit muscle myosin at an estimated molar ratio to actin of 1:1 (pH 6.2) was added to the cells while on plates. The plates were placed on ice, and the cells were quickly lysed in a buffer consisting of 10 mM Tris, 2 mM MgCl2, 0.2 mM dithioerythritol (DTE), 15% glycerol, and 1.0% Triton X-100 (pH 7.6). An aliquot was removed for the determination of total protein and total actin. The lysate was immediately centrifuged at 12,000 g for 1 min, and the resulting pellet (F-actin) was depolymerized to G-actin in actomyosin extraction buffer containing (in mM) 2 Tris, 0.2 CaCl2, 1.0 ATP, and 0.5 DTE (pH 8.0). The depolymerized pellet (originally the F-actin fraction) and the supernatant (G-actin) were then assayed by DNase inhibition. Fifty-microliter aliquots from each fraction were assayed for protein by the Bradford assay (2). Total actin from the original lysate was assayed in a guanidinium chloride buffer containing 1.5 M guanidinium chloride, 1 M Na acetate, 20 mM Tris, 7 mM CaCl2, and 1 mM ATP (pH 7.5).
Standards were prepared from rabbit muscle actin at a concentration of 1 mg/ml in actin monomer buffer consisting of (in mM) 2 Tris, 0.1 ATP, and 0.2 DTE (pH 7.6). Standard curves for each fraction were run in the appropriate buffer for that fraction. Calf thymus DNA was prepared at a 0.1 mg/ml concentration in DNase buffer consisting of (in mM) 125 Tris, 5 MgCl2, 2 CaCl2, and 3 NaN3 (pH 7.5). DNase 1 from bovine pancreas was used at a concentration of 0.1 mg/ml in DNase buffer. To carry out the reaction, 900 µl of DNA, 20 µl of DNase, and 100 µl of sample or standard plus buffer were added to a cuvette, mixed, and immediately read every 10 s for 90 s. Sample values were calculated from standard curves and expressed as nanograms of actin per microgram protein. The entire experiment was carried out a total of 10 times, but not all samples in each experiment could be used, resulting in individual sample replication of between 4 and 9.Immunoblotting analysis. The cells were plated in 60-mm dishes at 6.25 × 104 cells/cm2 in duplicate and treated as described above. On day 4, 10 ng EGF/ml medium were added in fresh medium (also containing the treatments) for 2 and 30 min. The cells were extracted and analyzed for total protein by the Bradford method (2). Total protein (50 µg) was separated by electrophoresis on 15% SDS-polyacrylamide gels and transferred to nitrocellulose membranes for Western blotting. Equal loading of protein samples was confirmed by Ponceau S staining of the membrane. The actin band was identified with rabbit polyclonal antiserum (Sigma, St. Louis, MO). A peroxidase-labeled secondary antibody was used for visualization (Sigma) with purified actin as the standard. The experiment was repeated five times.
RNA isolation and Northern blot analysis. Total RNA was extracted with guanidinium isothiocyanate solution and purified by CsCl density gradient ultracentrifugation. The resulting RNA pellet was dissolved in 10 mM Tris · HCl (pH 7.4) containing 5 mM EDTA and 1% SDS.
The purified RNA was precipitated from the aqueous phase with 0.1 vol of 3 M sodium acetate and 2.5 vol of ethanol in sequence and dissolved in water. Total RNA (30 µg) was denatured and fractionated electrophoretically on a 1.2% agarose gel containing 3% formaldehyde and transferred by blotting to a nitrocellulose membrane. The blot was prehybridized for 24 h at 42°C with 5× Denhardt's solution, 5× standard saline citrate (SSC), 50% formamide, 25 mM potassium phosphate, and 50 µg/ml denatured salmon sperm DNA. Hybridization was carried out overnight at 42°C in the same solution containing 10% dextran sulfate and DNA probes for TStatistics.
Results were analyzed by ANOVA and unpaired, two-tailed Student's
t-test or by nonparametric tests when
SD varied widely. Differences were considered significant when
P 0.05.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
F- and G-actin distribution.
The effects of polyamine depletion on the cytoskeletal response to EGF
are shown in Figs.
1-3,
in which column
1 shows merged images of F-actin (red)
and G-actin (green). Cells show areas of yellow where the two coincide.
Columns
2 and
3 of Figs. 1-3 show F- and
G-actin, respectively. Figures 1-3 depict control cells
(A), DFMO-treated cells
(B), and DFMO-putrescine-treated
cells (C).
|
|
|
F- and G-actin concentration.
In DFMO-treated cells, F-actin concentrations tended to be higher than
in control cells or those supplemented with putrescine (Fig.
4). This was the result of sporadic high
values in all 10 experiments, causing a variability that prevented
statistical significance. Similar variation did not occur in the
concomitant control and DFMO-putrescine-treated groups. F-actin
accounted for ~65% of the total actin in the control and
putrescine-supplemented groups and for 80% in DFMO-treated groups.
These values are within the range reported by Heacock and Bamburg (12)
for Chinese hamster ovary (CHO) cells. EGF, whether at 2 or at 30 min,
did not affect the concentration of F-actin in any of the three groups.
The discrepancy between the assay results and the apparent increases
shown by immunocytochemistry could be due to the loss of measurable
filaments to depolymerization in the assay. However, this is unlikely
since there were no concomitant increases in G-actin. Other
possibilities are rapid spikes of polymerization only occasionally
captured and increased visibility of previously invisible filaments.
|
|
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Structural and functional changes occur in IEC-6 cells in which
polyamine biosynthesis has been interrupted. Cell shape changes and
striking rearrangements of filamentous actin follow DFMO treatment, and
cell migration into a denuded area is reduced by 70-80% (18). We
have previously shown that polyamine-depleted cells also have an
abnormal distribution of the EGF receptor and fail to respond to an EGF
stimulus for growth and migration (17). All of these changes can be
prevented by providing the DFMO-treated cells with a polyamine
supplement. To gain an understanding of the mechanisms involved in the
disruption of migration, we investigated the distribution and quantity
of F-actin and G-actin and the localization of T4, a major
actin-monomer binding protein, in migrating polyamine-deficient cells
after stimulation with EGF. We also investigated the effect of DFMO on
the mRNA of T
4.
Confocal images of the confluent cell monolayer 3 h after wounding show
that both F- and G-actin were concentrated on the migrating margin of
control cells. T4 was distributed at low levels around the nucleus.
In DFMO-treated cells, interior actin stress fibers were greatly
reduced, whereas F-actin in the cell cortex was increased. This pattern
of F-actin distribution has been noted by us previously and earlier by
Pohjanpelto et al. (26) in a polyamine-deficient CHO cell variant.
Others have made the same observation under other experimental
conditions. Several possibilities have been proposed to explain the
abnormally dense cortex, namely, that peripheral actin filaments are
more resistant to depolymerization because they are composed primarily of
-actin, whereas inner stress fibers consist of more easily depolymerized
- and
-actin (14, 24). Yu and co-workers (40) have
suggested that there is less T
4 in the cell cortex to sequester actin monomers (although this has not been demonstrated) and also that
cortical actin is protected by other actin binding proteins. In
transfected cells that overexpress T
4, central F-actin disappears and peripheral actin is retained (35, 40). Adhesion plaques and
cytoskeletal proteins increase (9). We found nuclear staining of T
4
in DFMO-treated cells but not in control cells unless they were treated
with EGF. Nuclear staining of T
4 has been attributed to diffusion
from the cytoplasm (39) or to increased cell thickness over the nucleus
(40). Because Northern blots did not show an increase in total T
4
mRNA after DFMO, we believe the increased nuclear staining of T
4 is
more likely to be due to its translocation from the cytoplasm than to
an actual increase in quantity.
After 2 min of EGF treatment, control and DFMO-putrescine-supplemented
cells showed a striking increase in the staining of F-actin and T4
throughout the cell. In DFMO-treated cells, any increase in F-actin
occurred mainly in the cortex. Similar examples of rapid actin
polymerization have been reported in polymorphonuclear leukocytes
exposed to a chemoattractant (3) and in A431 cells after EGF treatment
(29). In A431 cells, newly assembled actin filaments localize and bind
selectively to the tyrosine-phosphorylated EGF receptor in the plasma
membrane (30), where they provide force to advance the lamellipodia
(21). In view of the fact that DFMO reduces tyrosine phosphorylation of
the EGF receptor in IEC-6 cells (17), the association of newly
polymerized actin filaments with the EGF receptor at the membrane may
be reduced as well. Decreased phosphotyrosine content in several
cellular substrates has been reported in DFMO-treated thymoma cells
also (23). Thirty minutes after EGF, the early changes in actin and T
4 in control and DFMO-putrescine-supplemented cells were returning to their pretreatment condition. The DFMO-treated cells, however, showed little evidence of change.
Changes in F- and G-actin quantity 2 min after EGF were suggestive but
not statistically significant in spite of the visual evidence that
suggested an increase in F-actin and a decrease in G-actin. Although it
is possible for F-actin to be underestimated due to depolymerization in
actin assays that depend on centrifugation alone (35), the Heacock and
Bamburg (12) DNase assay avoids depolymerization by complexing F-actin
with activated myosin at lysis. Visibly, the perceived density of actin
filaments may have been enhanced by reinforcement, bundling, and
cross-linking of preexisting very fine filaments by actin binding
proteins that made the filaments increasingly visible. Decreased
F-actin in the cell interior may interfere with the normal transport of
monomers and mRNA and contribute to increased cortical actin and
changes in the cellular distribution of T4. Western blot analysis
showed no changes in total actin caused by either DFMO or EGF, and the assayed amount of total actin per cell was within the reported range of
other cell lines (1, 12, 34).
T4 binds and sequesters free actin monomers, simultaneously
preventing premature actin polymerization and providing a reserve of
monomers for new polymerization in areas of F-actin remodeling (22). In
control and DFMO-putrescine-supplemented cells, T
4 responded rapidly
to EGF by increasing from few stainable areas over the majority of the
monolayer to many areas showing bright fluorescence in the nucleus and
cytoplasm. Rapid induction of T
4 has been observed by others as
well, namely, in thymocytes stimulated by conconavalin A (33) and in
NIH/3T3 serum-starved cells stimulated by serum (41). Both
investigators attributed the rapid increase in T
4 to translational
control (33, 41). Zalvide et al. (41) found that T
4 mRNA had
pronounced stability and that protein biosynthesis was not necessary
for T
4 elevation in response to a stimulus. In the present study, we
show that T
4 staining is primarily located in the nucleus in
DFMO-treated cells and does not change significantly with EGF.
In summary, novel findings in our study are as follows.
1) In migrating control cells,
stimulation by EGF caused the redistribution of F-actin to favor strong
stress fibers, attachment sites at the migrating edge, and attenuation
of G-actin without changes in the quantity of total actin. If the
migrating cells were polyamine depleted, F-actin was redistributed
primarily to the cell cortex, and G-actin became scattered and
disorganized. 2) In migrating control cells, the extent and intensity of T4 immunostaining increased markedly within 2 min of exposure to EGF and then returned to
the original condition within 30 min. In migrating polyamine-depleted cells, immunostaining of T
4 was primarily located in the nucleus and
was not significantly changed by EGF. The quantity of T
4 mRNA was
not affected by DFMO. 3)
Supplementation of polyamine-depleted cells with putrescine maintained
both actin forms and T
4 in their normal patterns of localization and
did not affect T
4 mRNA.
We hypothesize that two consequences of polyamine deficiency contribute
to the reduced ability of polyamine-deficient cells to migrate. These
are, first, aberrant actin monomer sequestration due to changes in
T4 intracellular distribution and, second, increased cortical actin
polymerization impeding the localization of newly polymerized actin
filaments on activated EGF receptors in the plasma membrane.
The link between polyamines and T4 remains unclear. It is known that
T
4 binds to monomeric actin through its amino-terminal residues in
an
-helical conformation (7, 31, 37). It is tempting to speculate
that the structure of the polyamines, namely a flexible carbon chain
with positive charges distributed at fixed lengths, may assist T
4 in
maintaining the
-helical conformation required for binding to
G-actin. To our knowledge, this study is the first report that attempts
to relate F-actin, G-actin, and T
4 to an underlying role for the
polyamines in intestinal cell migration.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Danny Morse for graphic assistance and Easter Jenkins for
final manuscript preparation. For the confocal images, we used the
Confocal Laser Scanning Microscope Facility, University of Tennessee
(Memphis, TN; National Institutes of Health Grant CLSM 1 S10-RR-08385,
Dr. Andrea Elberger, principal investigator). We gratefully acknowledge
the gifts of DFMO from Marion Merrell Dow (Cincinnati, OH), T4
antibody from Dr. Gregory Evangelatos (Radioimmunochemistry Laboratory,
National Centre for Scientific Research "Demokritos," Athens,
Greece), and T
4 murine
-4 cDNA from Dr. Grazyna Bozek (Dept. of
Molecular Genetics and Cell Biology, University of Chicago, Chicago, IL).
![]() |
FOOTNOTES |
---|
This research was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-52784 (to L. R. Johnson).
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. §1734 solely to indicate this fact.
Address for reprint requests: S. A. McCormack, Dept. of Physiology and Biophysics, University of Tennessee, Memphis, 894 Union Ave., Memphis, TN 38163.
Received 20 July 1998; accepted in final form 12 November 1998.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Blikstad, I.,
and
L. Carlsson.
On the dynamics of the microfilament system in HeLa cells.
J. Cell Biol.
93:
122-128,
1982
2.
Bradford, M. M.
A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem.
72:
248-254,
1976[Medline].
3.
Cano, M. L.,
L. Cassimeris,
M. Fechheimer,
and
S. H. Zigmond.
Mechanisms responsible for F-actin stabilization after lysis of polymorphonuclear leukocytes.
J. Cell Biol.
116:
1123-1134,
1992[Abstract].
4.
Cao, L.,
D. J. Fishkind,
and
Y. Wang.
Localization and dynamics of nonfilamentous actin in cultured cells.
J. Cell Biol.
123:
173-181,
1993[Abstract].
5.
Carlier, M.-F.,
D. Didry,
I. Erk,
J. Lepault,
M. L. Van Troys,
J. Vandekerckhove,
I. Perelroizen,
H. Yin,
Y. Doe,
and
D. Pantaloni.
T4 is not a simple G-actin sequestering protein and interacts with F-actin at high concentration.
J. Biol. Chem.
271:
9231-9239,
1996
6.
Carpinterio, P.,
R. Anadon,
G. G. del Amo,
and
J. Gomez-Marques.
The thymosin 4 gene is strongly activated in neural tissues during early postimplantation mouse development.
Neurosci. Lett.
184:
63-66,
1995[Medline].
7.
Feinberg, J.,
F. Heitz,
Y. Benyamin,
and
C. Roustan.
The N-terminal sequence (5-20) of thymosin 4 binds to monomeric actin in an alpha-helical conformation.
Biochem. Biophys. Res. Commun.
222:
127-132,
1996[Medline].
8.
Goldschmidt-Clermont, P. J.,
M. I. Furman,
D. Wachsstock,
D. Safer,
V. T. Nachmias,
and
T. D. Pollard.
The control of actin nucleotide exchange by thymosin beta 4 and profilin. A potential regulatory mechanism for actin polymerization in cells.
Mol. Biol. Cell
3:
1015-1024,
1992[Abstract].
9.
Golla, R.,
N. Philp,
D. Safer,
J. Chintapalli,
R. Hoffman,
L. Collins,
and
V. T. Nachmias.
Co-ordinate regulation of the cytoskeleton in 3T3 cells overexpressing thymosin-4.
Cell Motil. Cytoskeleton
38:
187-200,
1997[Medline].
10.
Gomez-Marquez, J.,
F. F. del Amo,
P. Carpintero,
and
R. Anadon.
High levels of mouse thymosin 4 mRNA in differentiating P19 embryonic cells and during development of cardiovascular tissues.
Biochim. Biophys. Acta
1306:
1187-1193,
1996.
11.
Haugland, R. P.,
W. You,
V. B. Paragas,
K. S. Wells,
and
D. A. Dubose.
Simultaneous visualization of G- and F-actin in endothelial cells.
J. Histochem. Cytochem.
42:
345-350,
1994
12.
Heacock, C. S.,
and
J. R. Bamburg.
The quantitation of G- and F-actin in cultured cells.
Anal. Biochem.
135:
22-36,
1983[Medline].
13.
Heidemann, S. R.,
and
R. E. Buxbaum.
Cell crawling: first the motor, now the transmission.
J. Cell Biol.
141:
1-4,
1998
14.
Hoock, T. C.,
P. M. Newcomb,
and
I. M. Herman.
Beta actin and its mRNA are localized at the plasma membrane and the regions of moving cytoplasm during the cellular response to injury.
J. Cell Biol.
112:
653-664,
1991[Abstract].
15.
Janson, L. W.,
and
D. L. Taylor.
Actin-crosslinking protein regulation of filament movement in motility assays: a theoretical model.
Biophys. J.
67:
973-982,
1994[Abstract].
16.
Kaminska, B.,
L. Kaczmarek,
and
B. Grzelabowska-Sztaberrt.
Inhibitors of polyamine biosynthesis affect the expression of genes encoding cytoskeletal proteins.
FEBS Lett.
304:
198-200,
1992[Medline].
17.
McCormack, S. A.,
P. M. Blanner,
B. J. Zimmerman,
R. Ray,
H. M. Poppleton,
T. B. Patel,
and
L. R. Johnson.
Polyamine deficiency alters EGF receptor distribution and signaling effectiveness in IEC-6 cells.
Am. J. Physiol.
274 (Cell Physiol. 43):
C192-C205,
1998
18.
McCormack, S. A.,
M. J. Viar,
and
L. R. Johnson.
Polyamines are necessary for cell migration by a small intestinal crypt cell line.
Am. J. Physiol.
264 (Gastrointest. Liver Physiol. 27):
G367-G374,
1993
19.
Meijerman, I.,
W. M. Blom,
H. J. de Bont,
G. J. Mulder,
and
J. F. Nagelkerke.
Nuclear accumulation of G-actin in isolated rat hepatocytes by adenine nucleotides.
Biochem. Biophys. Res. Commun.
240:
697-700,
1997[Medline].
20.
Mitchison, T. J.,
and
L. P. Cramer.
Actin-based cell motility and cell locomotion.
Cell
84:
371-379,
1996[Medline].
21.
Mogilner, A.,
and
G. Oster.
Cell motility driven by actin polymerization.
Biophys. J.
71:
3030-3045,
1996[Abstract].
22.
Nachmias, V. T.
Small actin-binding proteins: the beta-thymosin family.
Curr. Opin. Cell Biol.
5:
56-62,
1993[Medline].
23.
Oetken, C.,
T. Pessa-Morikawa,
M. Autero,
L. C. Andersson,
and
T. Mustelin.
Reduced tyrosine phosphorylation in polyamine-starved cells.
Exp. Cell Res.
202:
370-375,
1992[Medline].
24.
Otey, C. A.,
M. H. Kalnoski,
J. L. Lessard,
and
J. C. Bulinski.
Immunolocalization of the gamma isoform of nonmuscle actin in cultured cells.
J. Cell Biol.
102:
1726-1737,
1986[Abstract].
25.
Pantaloni, D.,
and
M.-F. Carlier.
How profilin promotes actin filament assembly in the presence of thymosin beta-4.
Cell
75:
1007-1014,
1993[Medline].
26.
Pohjanpelto, P.,
I. Virtanen,
and
E. Holtta.
Polyamine starvation causes disappearance of actin filaments and microtubules in polyamine-auxotrophic CHO cells.
Nature
293:
475-477,
1981[Medline].
27.
Pollard, T. D.,
and
J. A. Cooper.
Actin and actin-binding proteins, a critical evaluation of mechanisms and functions.
Annu. Rev. Biochem.
55:
987-1035,
1986[Medline].
28.
Quaroni, A.,
J. Wands,
R. L. Trelstad,
and
K. J. Isselbacher.
Epithelioid cell cultures from rat small intestine.
J. Cell Biol.
80:
248-265,
1979[Abstract].
29.
Rijken, P. J.,
W. J. Hage,
P. M. van Bergen en Henegouwen,
A. J. Verkleij,
and
J. Boonstra.
Epidermal growth factor induces rapid reorganization of the actin microfilament system in human A431 cells.
J. Cell Sci.
100:
491-499,
1991[Abstract].
30.
Rijken, P. J.,
S. M. Post,
W. J. Hage,
P. M. van Bergen en Henegouwen,
A. J. Verkleij,
and
J. Boonstra.
Actin polymerization localizes to the activated epidermal growth factor receptor in the plasma membrane, independent of the cytosolic free calcium transient.
Exp. Cell Res.
218:
223-232,
1995[Medline].
31.
Safer, D.,
T. R. Sosnick,
and
M. Elzinga.
Thymosin 4 binds actin in an extended conformation and contacts both the barbed and pointed ends.
Biochemistry
36:
5806-5816,
1997[Medline].
32.
Santos, M. F.,
M. J. Viar,
S. A. McCormack,
and
L. R. Johnson.
Polyamines are important for attachment of IEC-6 cells to extracellular matrix.
Am. J. Physiol.
273 (Gastrointest. Liver Physiol. 36):
G175-G183,
1997
33.
Schobitz, B.,
R. Netzker,
E. Hannappel,
and
K. Brand.
Cell-cycle-regulated expression of thymosin 4 in thymocytes.
Eur. J. Biochem.
199:
257-262,
1991[Abstract].
34.
Snabes, M. C.,
A. E. Boyd,
R. L. Pardue,
and
J. Bryan.
A DNase I binding/immunoprecipitation assay for actin.
J. Biol. Chem.
256:
6291-6295,
1981
35.
Sun, H. Q.,
K. Kwiatkowska,
and
H. Yin.
-Thymosins are not simple actin monomer buffering proteins.
J. Biol. Chem.
271:
9223-9230,
1996
36.
Tabor, C. W.,
and
H. Tabor.
Polyamines.
Annu. Rev. Biochem.
53:
749-790,
1984[Medline].
37.
Van Troys, M.,
D. Dewitte,
M. Goethals,
M.-F. Carlier,
J. Vandekerckhove,
and
C. Ampe.
The actin binding site of thymosin 4 mapped by mutational analysis.
EMBO J.
15:
201-210,
1996[Abstract].
38.
Wang, J.-Y.,
and
L. R. Johnson.
Polyamines and ornithine decarboxylase during repair of duodenal mucosa after stress in rats.
Gastroenterology
100:
333-343,
1991[Medline].
39.
Watts, J.,
P. D. Cary,
P. Sautiere,
and
C. Crane-Robinson.
Thymosins: both nuclear and cytoplasmic proteins.
Eur. J. Biochem.
192:
643-651,
1990[Abstract].
40.
Yu, F.-X.,
S.-C. Lin,
M. Morrison-Bogorad,
and
H. L. Yin.
Effects of thymosin beta 4 and thymosin beta 10 on actin structures in living cells.
Cell Motil. Cytoskeleton
27:
13-25,
1994[Medline].
41.
Zalvide, J. B.,
C. V. Alvarez,
A. Vidal,
D. Dieguez,
F. V. Vega,
and
F. Dominguez.
Regulation of thymosin beta 4 mRNA levels during cell proliferation.
Cell Prolif.
28:
85-91,
1995[Medline].