Regulation of cytoskeletal mechanics and cell growth by myosin
light chain phosphorylation
Shuang
Cai1,
Lidija
Pestic-Dragovich1,
Martha E.
O'Donnell2,
Ning
Wang3,
Donald
Ingber4,
Elliot
Elson5, and
Primal
De
Lanerolle1
1 Department of Physiology and
Biophysics, University of Illinois at Chicago, Chicago, Illinois
60612-7342; 2 Department of Human
Physiology, University of California, Davis, California 95616-8644;
3 Physiology Program, Harvard
University School of Public Health, Boston, 02115-6021;
4 Departments of Pathology and
Surgery, Children's Hospital and Harvard University Medical School,
Boston, Massachusetts 02116-5737; and
5 Department of Biophysics and
Molecular Biophysics, Washington University Medical School, St. Louis,
Missouri 63110-1093
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ABSTRACT |
The role of
myosin light chain phosphorylation in regulating the mechanical
properties of the cytoskeleton was studied in NIH/3T3 fibroblasts
expressing a truncated, constitutively active form of smooth muscle
myosin light chain kinase (tMK). Cytoskeletal stiffness determined by
quantifying the force required to indent the apical surface of adherent
cells showed that stiffness was increased twofold in tMK cells compared
with control cells expressing the empty plasmid (Neo cells).
Cytoskeletal stiffness quantified using magnetic twisting cytometry
showed an ~1.5-fold increase in stiffness in tMK cells compared with
Neo cells. Electronic volume measurements on cells in suspension
revealed that tMK cells had a smaller volume and are more resistant to
osmotic swelling than Neo cells. tMK cells also have smaller nuclei,
and activation of mitogen-activated protein kinase (MAP kinase) and
translocation of MAP kinase to the nucleus are slower in tMK cells than
in control cells. In tMK cells, there is also less
bromodeoxyuridine incorporation, and the doubling time is
increased. These data demonstrate that increased myosin light chain
phosphorylation correlates with increased cytoskeletal stiffness and
suggest that changing the mechanical characteristics of the
cytoskeleton alters the intracellular signaling pathways that regulate
cell growth and division.
cell stiffness; osmotic swelling; volume regulation; mitogen-activated protein kinase activation; cell division; myosin
light chain kinase
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INTRODUCTION |
THE ABILITY OF THE cytoskeleton to deform and reform is
a crucial aspect of many cellular responses, such as cell motility and
cell division (4). Migrating cells continuously change their
cytoskeleton, and preventing these cytoskeletal changes by affecting
actin polymerization or myosin II phosphorylation inhibits cell
motility (47). A plastic cytoskeleton is also essential for proper
progression through the cell cycle. Specific cytoskeletal alterations
are required for a cell to enter S phase (20), and disrupting actin
filaments prevents S phase entry and blocks mitosis (4, 32, 50). In
addition, exposure to growth factors results in the stimulation of the
mitogen-activated protein kinase (MAP kinase) pathway and cytoskeletal
changes (20, 25), both of which are needed for a cell to commit to
mitosis (25). Other experiments have shown that stabilizing
microtubules blocks mitosis (39) and that inhibiting myosin II activity
blocks cytokinesis (10, 26). Thus many cellular responses depend on the
ability of the cytoskeleton to restructure itself.
The organization and, hence, the plasticity of the cytoskeleton is
determined primarily by the forces or tension generated within the
cytoskeleton by actin and myosin II (12, 23, 29, 39). The actin-myosin
II interaction in smooth muscle and nonmuscle cells is regulated by the
phosphorylation of the 20-kDa light chain of myosin by the enzyme
myosin light chain kinase (MLCK) (1, 44). MLCK is naturally a
calcium/calmodulin-dependent enzyme (1, 44). Proteolytic digestion (22)
or insertion of a stop codon (19) between the catalytic and regulatory
domains results in a truncated, constitutively active enzyme.
Expression of the truncated catalytic domain of MLCK (tMK) in 3T3 cells
increases myosin light chain phosphorylation (33). Therefore, to
further characterize the relationship between cytoskeletal plasticity and cellular responses, we quantified the mechanical properties of the
cytoskeleton, MAP kinase activation, and cell growth in NIH/3T3 cells
expressing tMK or the empty plasmid.
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MATERIALS AND METHODS |
Cell culture.
NIH/3T3 fibroblasts were transduced with either the pLNCX Moloney
murine leukemia virus-based vector (a gift from A. Dusty Miller) or the
same vector engineered to contain tMK. In this vector, the neomycin
resistance gene is driven from the viral long terminal repeat, and the
cloned genes are driven from the cytomegalovirus promoter. The tMK gene
was constructed by inserting a stop codon following Lys-793 of the
chicken gizzard smooth muscle MLCK gene (19). The 1995-bp tMK gene was
passed through pBluescript and cloned into the pLNCX vector. The pLNCX
and pLNCX-tMK DNA were then used to transfect GP+envAM12 amphotropic
packaging cells. The packaging cells were then selected in G418 and
expanded in culture. Medium from packaging cells in log phase growth,
which contains infective viral particles, was then used to infect
NIH/3T3 cells. Infected cells were selected for 8 days in 0.9 mg/ml
G418 (effective concentration), expanded in culture, and used in the experiments described. Vector construction, generation of packaging cells, and infection and selection of NIH/3T3 cells are described in
detail in Ref. 33. Cells transduced with the neomycin resistance gene
alone (Neo cells) and tMK cells were grown in DMEM containing 10%
fetal bovine serum (FBS) and antibiotics at 37°C in an atmosphere containing 5% CO2.
Myosin light chain phosphorylation.
Neo or tMK cells grown in 60-mm dishes were washed in phosphate-free
DMEM and incubated with 50 µCi of
32P in 1 ml of phosphate-free DMEM
for 6 h at 37°C. The cells were then washed with ice-cold PBS,
frozen on dry ice, and extracted in 1% NP-40, 40 mM sodium
pyrophosphate, 100 mM NaF, 500 mM NaCl, 10 mM EGTA, 5 mM EDTA, 50 µg/ml leupeptin, 50 µg/ml pepstatin, and 25 mM Tris (pH 7.9). The
supernatant was collected by centrifugation (50,000 g, 20 min) and incubated with 10 µg
of affinity-purified antibody to macrophage myosin II for 2 h, followed
by incubation with protein A-Sepharose for 1.5 h. The beads were then
washed extensively, and the myosin II-antibody complexes were eluted by
boiling in 0.25% SDS, 5 mM dithiothreitol (DTT), and 5 mM Tris (pH
6.8). The immunoprecipitates were then analyzed by SDS-PAGE and
autoradiography.
Quantification of cytoskeletal stiffness.
Mechanical deformation assays were performed as described by Worthen et
al. (49). Cells were plated on coverslips coated with 15 µg/ml
poly(2-hydroxyethyl methacrylate) and allowed to attach for 4-6 h.
The coverslips were then mounted in the chamber of the cell poker in
the inverted position. Stiffness measurements were performed in
degassed medium containing FBS at 37°C. The surface of the cells
was indented near the center (depth of indentation was <2.6 µm and
velocity of indentation was 5.1 µm/s) with a glass microprobe (tip
diameter ~2 µm) attached to a flexible glass beam of known bending
constant. The degree of bending of the glass beam is used to calculate
cellular deformability (i.e., stiffness), which is the force resisting
indentation (in millidynes) per unit of indentation depth (in
micrometers), as described by Worthen et al. (49). Some cells were
pretreated with okadaic acid for 10 min at 37°C or with okadaic
acid for 10 min followed by cytochalasin D for 10 min at 37°C.
Deformation assays were then performed for a maximum of 30 min at
37°C in the continued presence of the drugs.
Magnetic twisting cytometry (21, 46) was performed on cells grown in
96-well dishes. The resistance to twisting of beads coated with RGD
peptides, in the presence or absence of 3 µM cytochalasin D, and of
beads coated with acetylated low-density lipoproteins (AcLDL) was
tested. Briefly, ~2 × 104
ferromagnetic beads coated with RGD peptide or AcLDL were added to each
well and incubated for 10-20 min at 37°C. Unbound beads were
removed by washing in serum-free medium, and the cells were placed
within the magnetometer and maintained at 37°C. The beads were then
magnetized with a 1,000-gauss pulse so that their magnetic moments were
aligned in one direction. A second magnetic field (26 gauss) that was
too weak to remagnetize the beads was applied orthogonally to the
original field. The beads rotated to reorient their magnetic moments
with the new field. Changes in the component of the remanent magnetic
field generated by the beads in the original direction were measured by
an in-line magnetometer and reflect bead rotation. The rotation of
RGD-coated beads is resisted by the cytoskeleton and is inversely
proportional to the stiffness of the cytoskeleton. The stiffness was
defined as the ratio of the applied stress to the measured bead
rotation (strain).
Quantification of responses to changes in osmolarity.
The responses of cells in monolayer to decreasing osmolarity was
determined using an ECIS Biosensor (17). Cells were plated in a
modified 96-well tissue culture plate containing a
10
3
cm2 gold electrode and a large
counter electrode. A 1-µA current was applied between the gold and
counter electrodes, and the voltage was monitored using a lock-in
amplifier. As cells attach, spread, and make cell-cell contacts, the
insulating cell bodies block the current path and the resistance
increases. The increase in resistance is a function of cell thickness
and the integrity of the cell contacts, and changes in resistance
reflect changes in cell morphology. The effect of changing osmolarity
was determined by first equilibrating the cells in HEPES-buffered
Hanks' salt solution (290 mosM; 20 mM HEPES). After baseline
resistance measurements, the osmolarity was adjusted by sequentially
replacing one-half of the medium bathing the cells with (in mM) 5.5 KCl, 1 CaCl2, 1 MgSO4, and 20 HEPES (pH 7.4) every
10 min, and the resistance was measured continuously.
The responses of cells in suspension to decreasing osmolarity were
determined by performing electronic cell sizing (34). Cells were
removed from the tissue culture plates by brief typsinization and
washed and resuspended at 5 × 104 cells/ml in HEPES-buffered
Hanks' salt solution (290 mosM; 20 mM HEPES) and were maintained at
37°C throughout the experiment. The osmolarity of the cells in
suspension was adjusted by sequentially diluting the isosmotic solution
with a hyposmotic solution consisting of (in mM) 5.5 KCl, 1 CaCl2, 1 MgSO4, and 20 HEPES (pH 7.4). Cell volumes were measured using a Coulter counter (model ZM) with a Coulter
channelizer (C-1000). Absolute cell volumes were calculated from
distribution curves of cell diameter, using a standard curve generated
by polystyrene latex beads of known diameter (9.97 and 14.51 µm).
Measurement of nuclear sizing.
Relative nuclear size was determined by performing flow cytometry on
cells synchronized by growing them in 0.5% FBS for 48 h. The cells
were then harvested, fixed, permeabilized, stained with propidium
iodide (45), and analyzed for their DNA content using a Coulter EPICS
Elite ESP flow cytometer. The pulse width of the fluorescence peak
(time of flight) was used to obtain relative nuclear size. Nuclear size
was measured three times, but only the data from a single experiment
are reported. The time of flight is a relative measurement, and there
is substantial interexperiment variability due to variability in the
binding of propidium iodide to the DNA. Therefore, in each experiment,
the Neo and tMK cells were incubated with propidium iodide for the same
amount of time, and the size of the nuclei in Neo cells averaged 89 relative units larger in all three experiments.
Growth characteristics.
To determine the doubling time, Neo and tMK cells were synchronized by
growing them in the presence of 0.5% FBS for 48 h. The cells were then
trypsinized and washed in medium containing 0.5% serum, and 50,000 Neo
and tMK cells were plated in triplicate in 0.5% serum. After
attachment, the cells were serum stimulated by raising the serum
concentration to 10%. Cells were trypsinized at 0, 24, 48, and 72 h
after serum stimulation and counted. A curve-fitting program was then
used to determine the doubling time. Only the mean doubling time is
reported in Table 1 because the
curve-fitting program used to calculate the doubling time only
considers the mean of data at each time point. Bromodeoxyuridine (BrDU)
incorporation was quantified in cells that were synchronized and serum
stimulated as described above. Cells were incubated with BrDU and
stained with an antibody to BrDU, and positive cells were expressed as
a percentage of total cells, as previously described (18).
MAP kinase assays.
Cells synchronized and serum stimulated as described above were washed
quickly in ice-cold PBS at various times and frozen on dry ice. Cells
were extracted in ice-cold 1% Triton X-100, 5 mM EGTA, 5 mM
MgCl2, 1 mM benzamidine, 1 mM DTT,
1 mM sodium vanadate, 10 mM sodium pyrophosphate, 10 µg/ml aprotinin,
10 µg/ml leupeptin, 2 µg/ml pepstatin A, and 40 mM HEPES (pH 7.5),
as described (40). Thirty micrograms of each extract were boiled in SDS
sample buffer, and the proteins were separated by SDS-PAGE, transferred to nitrocellulose, and probed with an antibody to extracellular signal-regulated kinase (ERK) 1 (no. 06-182, Upstate Biotechnology, Lake Placid, NY). The primary antibodies were visualized by incubation with peroxidase-labeled secondary antibodies and development of the
color reaction. These primary antibodies primarily recognize ERK1, and
developing the color reaction to visualize ERK2 obscured ERK1.
Therefore, only ERK1 is shown (see Fig. 7). To quantify changes in MAP
kinase activity, cells were extracted in nondenaturing buffer, MAP
kinase was immunoprecipitated, and MAP kinase activity assays were
performed using
[
-32P]ATP and
myelin basic protein (MBP) as substrate, as described (40). Aliquots of
the reaction mixtures were analyzed by SDS-PAGE, and the bands
representing the MBP were excised and counted. The translocation of MAP
kinase was investigated by performing antibody immunofluorescence
studies on cells plated on fibronectin-coated coverslips. These cells
were synchronized and serum stimulated as described above. Cells were
then fixed at time zero (no serum stimulation) and at 60 min after serum stimulation in freshly made 3%
formaldehyde in PBS for 7 min and were permeabilized by incubation in
0.1% Triton X-100 and 0.1% deoxycholate in PBS for 7 min. MAP kinase
was visualized using the same antibodies to ERK1 and ERK2 described
above and a Texas red-labeled secondary antibody. The coverslips were
mounted on slides and photographed using a Zeiss IM 35 inverted
photomicroscope and a Planapo ×63, 1.4-numerical aperture
objective.
Statistical analyses.
The data were evaluated by Student's
t-test and considered statistically
significant when P < 0.05.
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RESULTS |
MLCK is a calcium/calmodulin-dependent enzyme that phosphorylates
Ser-19 of the 20-kDa light chain of vertebrate smooth muscle and
nonmuscle myosin II (1, 44). Myosin light chain phosphorylation stimulates the actin-activated ATPase activity of myosin II purified from smooth muscle and nonmuscle cells (1, 44). This reaction regulates
smooth muscle contraction (9) and promotes the association of myosin II
into filaments (43, 48). Proteolytic digestion of the purified enzyme
(22) or insertion of a stop codon following Lys-793 (19) results in a
truncated, constitutively active enzyme that maintains the specificity
of the parent enzyme for Ser-19. Therefore, NIH/3T3 fibroblasts were
transduced with retroviral vectors containing either the neomycin
resistance gene and tMK (tMK cells) or the neomycin resistance gene
alone (Neo cells) and selected in G418 (33). We previously reported
that the stoichiometry of myosin light chain phosphorylation is
increased in tMK cells compared with Neo cells (Table 1) and that this
phosphorylation is confined to Ser-19 (33). Immunoprecipitation of
myosin II from Neo and tMK cells labeled with
32P performed throughout these
experiments (Fig. 1) demonstrated increases
in myosin light chain phosphorylation qualitatively similar to those
previously reported (33).

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Fig. 1.
Myosin light chain phosphorylation in Neo and tMK cells. Myosin was
immunoprecipitated from Neo and tMK cells labeled with
32P and analyzed by SDS-PAGE and
autoradiography. Note marked increase in myosin light chain (MLC)
phosphorylation in tMK cells compared with Neo cells. There was
approximately the same amount of myosin heavy chain (MHC), as judged by
Coomassie blue staining, in both lanes.
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The effects of increasing myosin light chain phosphorylation on
cytoskeletal mechanics was studied using various assays. First, cortical tension was quantified directly by using a microprobe to
deform the free surface of adherent cells (49). Mean cytoskeletal stiffness was increased almost 100% in tMK cells compared with Neo
cells (Fig. 2). Okadaic acid (0.03-3.0
µM), a phosphatase inhibitor that increases myosin light chain
phosphorylation (48), increased stiffness in a dose-dependent fashion
in Neo cells (not shown), with the maximal effect observed at 3 µM.
Okadaic acid had no effect on the stiffness of tMK cells, suggesting
that cytoskeletal stiffness is near maximal in tMK cells. The
combination of 3 µM okadaic acid and 3 µM cytochalasin D, which
disrupts actin filaments (7), decreased stiffness in Neo cells to a
very low level (Fig. 2). In contrast, stiffness remained elevated in
tMK cells under similar conditions.

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Fig. 2.
Quantification of cytoskeletal stiffness by mechanical deformation.
Mechanical deformation assays were performed on untreated cells, cells
treated with 3 µM okadaic acid (OA), and cells treated with 3 µM
okadaic acid and 3 µM cytochalasin D (Cyto D), as described in
MATERIALS AND METHODS. Data are means ± SE; n > 35 cells in each
group. * P 0.05 compared with
respective control.
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Cytoskeletal stiffness was also quantified by measuring the ability of
cell surface receptor-bound magnetic beads to resist shear stresses
applied using magnetic twisting cytometry (46). Beads coated with RGD
peptide, which is a ligand for integrin receptors (46), showed ~50%
increase in stiffness in tMK cells compared with Neo cells (Fig.
3). Okadaic acid also increased the
twisting stiffness in Neo cells in a dose-dependent fashion (data not
shown). Cytochalasin D treatment (3 µM, final) decreased twisting
stiffness in both cell types (Fig. 3), indicating that the tMK effects
on stiffness are mediated, at least in part, through an interaction
with microfilaments. This conclusion is supported by the observation
that the twisting of beads coated with AcLDL, a ligand for a
transmembrane receptor that is not efficiently coupled to the
cytoskeleton (46), was equally low in both cell types (Fig. 3).

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Fig. 3.
Quantification of cytoskeletal stiffness by magnetic twisting
cytometry. Magnetic twisting cytometry using RGD-coated beads in
presence or absence of 3 µM cytochalasin D or beads coated with
acetylated low-density lipoproteins (AcLDL) was performed as described
in MATERIALS AND METHODS. Data are
means ± SE; n 3. * P 0.05 compared with
respective control.
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We then investigated the ability of Neo and tMK cells to resist
internal deformation by examining the responses of plated cells and
cells in suspension to decreasing osmolarity. The responses of cells in
monolayer were determined using an ECIS Biosensor (17) in which cells
are plated in a well of a modified tissue culture dish that contains a
small gold electrode and a counter electrode. A current passed across
the gold electrode is restricted and eventually plateaus as the cells
attach and form a confluent monolayer on the electrode. Cell
contraction increases the dimensions of the paracellular spaces,
thereby increasing current flow and decreasing resistance. Decreasing
osmolarity is predicted to increase resistance by
1) swelling the cells and thus
increasing the thickness of the monolayer and
2) decreasing the dimensions of the
paracellular spaces, both of which impede current flow. Figure
4 shows that the peak resistance is greater
in Neo cells at all osmolarities, as predicted for cells with a softer
cytoskeleton. Moreover, at all osmolarities, there is a greater
increase in peak resistance in Neo cells than in tMK cells (Fig. 4).

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Fig. 4.
Changes in electrical resistance in response to decreasing osmolarity
in tMK and Neo cells grown in monolayers. Osmolarity was decreased
sequentially, and peak electrical resistance was measured using an ECIS
Biosensor. Note that resistance is higher and rises more rapidly in Neo
cells than in tMK cells (means ± SE;
n 4 separate sets of cells). All
values for tMK cells are significantly different
(P < 0.05) from those for Neo cells
at all osmolarities except 290 mosM.
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The responses of cells in suspension to decreasing osmolarity were
determined by performing electronic cell sizing (34). Figure
5 shows that tMK cells in suspension are
smaller in isosmolar medium than Neo cells, an observation that is
consistent with an internal contraction of the cytoskeleton in tMK
cells. Figure 5 also shows that tMK cells are much more resistant to
osmotic swelling than are Neo cells and that Neo cells appear to lyse at a higher osmolarity than tMK cells. Thus tMK cells have a stiffer, more stable cytoskeleton than Neo cells (Figs. 2 and 3); this increase
in cytoskeletal stiffness correlates with increased resistance to
osmotic swelling (Figs. 4 and 5).

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Fig. 5.
Responses of Neo and tMK cells in suspension to decreasing osmolarity.
Osmolarity of cells in suspension was decreased sequentially, and cell
volume was quantified electronically (MATERIALS AND
METHODS). No Neo cells were detected below 145 mosM,
suggesting that cells had ruptured. Data are means ± SE;
n 3. All values for tMK cells are
significantly different (P < 0.05)
from those for Neo cells at all osmolarities.
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The effects of increasing myosin light chain phosphorylation on cell
growth and division were studied next. Analysis of the growth rate
showed that the doubling time is significantly increased in tMK cells
compared with Neo cells (Fig. 6 and Table
1). In addition, more Neo cells than tMK cells were in S phase at 24 h
following serum stimulation of synchronized cells as judged by BrDU
incorporation (Table 1). Because cell size has been correlated with S
phase entry and division (see
DISCUSSION), we performed assays to
quantify cell and nuclear size. Electronic cell sizing demonstrated
that tMK cells in isotonic solution are smaller than Neo cells (Fig. 5
and Table 1). Time of flight measurements were also performed to
compare the sizes of the nuclei in Neo and tMK cells. Although three
such experiments were performed, Table 1 shows the data from a single
experiment because the time of flight gives a relative measure of
nuclear size due to variability in the binding of propidium iodide to
the DNA. Nevertheless, the nuclei in Neo cells averaged 89 relative
units larger than those in tMK cells in all three experiments.

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Fig. 6.
Doubling time of Neo and tMK cells. Synchronized cells were trypsinized
and counted, and equal numbers of cells were plated in triplicate as
described in MATERIALS AND METHODS.
Cells were then serum stimulated and counted at times shown. Data are
means for triplicates from a single experiment, and tMK values at 24, 48, and 72 h are significantly different from those for Neo cells at
same times.
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We then investigated whether the decrease of growth kinetics in tMK
cells was associated with alterations in the activation of MAP kinase.
Gel shift assays that are designed to detect the phosphorylated,
activated forms of ERK1 and ERK2 showed that the activation of ERK1
(Fig. 7) and ERK2 (not shown) is slower in tMK cells than in Neo cells. The data also suggest that the rate of ERK
dephosphorylation is also faster in tMK cells (Fig. 7). Phosphorylation
assays (Fig. 8) performed on
immunoprecipitated enzyme using MBP as a substrate confirmed that MAP
kinase activation is retarded in tMK cells compared with Neo cells.
Immunofluorescence assays were also performed because MAP kinase is
known to translocate to the nucleus following mitogenic stimulation (5,
32, 40, 50). Figure 9 shows that the
translocation of MAP kinase to the nucleus is retarded in tMK cells
compared with Neo cells at 60 min after serum stimulation.

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Fig. 7.
Gel shift assay of mitogen-activated protein kinase (MAP kinase)
activation in Neo (top) and tMK (bottom)
cells. Synchronized Neo and tMK cells were stimulated with serum and
analyzed as described in MATERIALS AND METHODS. Active,
phosphorylated, and slower migrating form of MAP kinase, extracellular
signal-regulated kinase (ERK) 1 (+PO4), appears by 5 min in Neo cells, whereas it does not appear until 10 min after serum
stimulation in tMK cells. Although only ERK1 is shown, developing color
reaction longer showed a similar relationship for ERK2 in Neo and tMK
cells. This experiment was repeated 3 times.
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Fig. 8.
MAP kinase activity assays. Synchronized Neo and tMK cells were
stimulated with serum, MAP kinase was immunoprecipitated, and kinase
activity was analyzed as described in MATERIALS AND METHODS
using myelin basic protein (MBP) as substrate. A: Coomassie
blue staining (SDS-PAGE) and autoradiogram (AUTORAD) of region of gel
containing MBP. B: experiment in A was repeated 3 times, and aggregate data (means ± SE) from these experiments are
shown. Data are expressed as increase in 32P
incorporation into MBP in terms of multiples of the level in Neo cells
at time 0, which is set to 1. * Difference in MBP
phosphorylation by Neo and tMK cells at 5 min after serum stimulation
is statistically significant (P < 0.05).
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Fig. 9.
Immunofluorescence localization of MAP kinase. Synchronized Neo
(A-C)
and tMK
(D-F)
cells before (A and
D) and after 60 min
(B and
E) and 120 min
(C and
F) of serum stimulation were fixed
and stained with antibodies to MAP kinase. Note nuclear staining in Neo
cells (B) and absence of nuclear
staining in tMK cells (E) at 60 min.
All cells (100%) of both cell types contained nuclear staining at 120 min.
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DISCUSSION |
Inhibiting actin-myosin II interactions is known to decrease
cytoskeletal stiffness. Experiments have shown that disrupting actin
filaments with cytochalasin D decreases cytoskeletal stiffness as
quantified by both mechanical deformation (49) and magnetic twisting
cytometry (46). Similarly, cortical tension is lower in
Dictyostelium amoebas lacking myosin
II heavy chain (35). The implication of these data is that decreasing
actin-myosin II interactions releases the internal tension generated by
these proteins and makes the cytoskeleton less stiff. They also predict that the converse, namely increasing actin-myosin II interactions, should increase stiffness by generating a contractile tension within
cells. This, in turn, leads to the hypothesis that myosin light chain
phosphorylation, which regulates the actin-myosin II interaction in
smooth muscle and nonmuscle cells (1, 9, 44), determines at least in
part the stiffness of the cytoskeleton. This hypothesis is indirectly
supported by the demonstration that vasoconstrictors and vasodilators
that are known to modulate myosin light chain phosphorylation increase
and decrease cytoskeletal stiffness, respectively, in smooth muscle
cells (21). We tested this hypothesis by studying the mechanical
properties of cells expressing the catalytic domain of MLCK. The data
presented in Figs. 2-5 provide direct support for it.
The data in Figs. 4 and 5 also implicate myosin light chain
phosphorylation in regulating cell swelling and support the hypothesis that the cytoskeleton acts as a resistive element to volume changes (3,
8, 31). Although volume regulation is a complex process, the
cytoskeleton contributes to the cellular response to changes in
osmolarity (3, 8, 31), and
Dictyostelium resist volume changes in
response to increasing osmolarity by redistributing their actin and
myosin to the cortex (27). Other experiments have shown that muscle
cells from MDX mice lacking dystrophin have a decreased stability in
response to osmotic shock (30) compared with normal cells containing
dystrophin. Mechanical deformation assays have also shown that muscle
cells from MDX mice have a substantially lower stiffness than matched
normal cells (30, 36). These studies demonstrate that decreasing
stiffness by depleting the cortical cytoskeleton of specific structural
proteins also results in a decrease in osmotic stability. Our data
further support the hypothesis that the cytoskeleton resists volume
changes (3, 8, 31), by demonstrating for the first time that cells with
increased myosin light chain phosphorylation and a stiffer cytoskeletal
are more resistant to osmotic swelling.
Our data also provide insights into how changes in the physical
characteristics of the cytoskeleton affect cell growth and division. It
is well known that actin and microfilaments are crucial for cell growth
and cell division. Actin is considered an early response gene (11, 24),
and actin-dependent morphological changes are among the earliest seen
following growth factor or serum stimulation (20). In addition, growth
factor stimulation or ligation of integrin receptors by extracellular
matrix activates signaling pathways that result in MAP kinase
activation and cytoskeletal changes, both of which are essential for
G0-to-G1
transition (25). It is also known that cytochalasin D blocks MAP kinase
activation and progression through the cell cycle (20). Thus decreasing cytoskeletal stiffness by disrupting the actin cytoskeleton blocks cell
cycle progression. Similarly, our data show that increasing cytoskeletal stiffness (Figs. 2-5) also retards cell growth and cell division (Table 1 and Fig. 6). Moreover, the data suggest that
both decreasing and increasing cytoskeletal tension alter the efficient
transduction of signals that regulate cell growth, as indicated by
delays in MAP kinase activation and extended doubling time (Figs.
7-9).
How changes in the mechanical properties of the cytoskeleton affect
cell division is currently unclear. It is clear that the transition
from interphase into M phase requires almost complete dissolution of
all components of the cytoskeleton (2). Tubulin must subsequently
reassemble into the spindle while actin and myosin II must congregate
in the cleavage furrow before cytokinesis (2). This is a precise,
highly regulated process. Disrupting this process by destabilizing
actin filaments (20) or stabilizing microtubules (41) blocks mitosis.
The implication is that disrupting the timing of these cytoskeletal
changes can either block or retard mitosis. Our data demonstrate that
tMK cells enter S phase more slowly (Table 1) and have longer doubling
times than Neo cells (Fig. 6). While not excluding other mechanisms,
our data suggest that increasing myosin light chain phosphorylation
prevents the timely dissolution of the cytoskeleton, perhaps in a
manner that is analogous to the effect of taxol on cell division.
Another possibility is that cytoskeletal stiffness affects cell growth.
Cells must grow by doubling their DNA and protein contents and
duplicating their organelles before they can divide (2). Cell growth is
profoundly affected by cell shape (37). Highly spread cells exhibit
higher rates of DNA synthesis and cell division (14, 42). Cell
spreading, in turn, appears to be controlled by the mechanical tension
generated within the cytoskeleton by actin-myosin II interactions (23,
24, 41). The effect of spreading on DNA synthesis and cell division may
be due to a direct effect of cytoskeletal mechanics on the nucleus.
Maniotis et al. (29) demonstrated a direct connection between the
cytoskeleton and the nucleus by showing that exerting mechanical force
on cell surface integrin receptors changes the shape of the nucleus
through the cytoskeleton. tMK cells have a stiffer cytoskeleton, are
smaller, have smaller nuclei, enter S phase more slowly, and have more extended doubling times than Neo cells. When the data are considered in
the context of the work described above (4, 14, 23, 24, 29, 41, 42), it
seems possible that tMK cells take longer to enter the cell cycle and
divide because tension generated by more actin-myosin II cross bridges
retards both spreading and cell growth.
Our data also suggest a more complex role for myosin II in cell
division than previously suspected. Myosin II has been localized in the
cleavage furrow (15, 16) and shown to be essential for cytokinesis in
metazoan cells. Classic experiments have shown that injecting an
antibody to myosin II into starfish blastomeres blocks cytokinesis
(28), whereas genetic manipulation of myosin II heavy chain expression
in Dictyostelium
discoideum results in multinucleated
cells (10, 26). Myosin light chain phosphorylation has also been
implicated in regulating the timing of cytokinesis (13, 38). It is
possible that tMK cells divide more slowly because increasing myosin
light chain phosphorylation retards the redistribution of myosin II to
the cleavage furrow before cytokinesis. At the same time, the
observations that MAP kinase activation, S phase entry, and doubling
time are altered in tMK cells suggest that myosin II is involved in
cell growth and division in multiple, complex ways in addition to
mediating cytokinesis.
In conclusion, we used multiple approaches to obtain the most
compelling data to date that myosin light chain phosphorylation regulates the mechanical characteristics of the cytoskeleton. Our data
also provide important support for the idea that the cortical
cytoskeleton is the major barrier to osmotic swelling in mammalian
cells. In addition, the data demonstrate that changes in myosin light
chain phosphorylation affect the kinetics of MAP kinase activation and
progression through the cell cycle. Our current working hypothesis is
that the plasticity of the cytoskeleton is regulated in great part by
changes in the level of myosin light chain phosphorylation and that
changes in myosin light chain phosphorylation affect signal
transduction pathways by altering the mechanical properties of the
cytoskeleton. This hypothesis predicts that decreasing myosin light
chain phosphorylation will soften the cytoskeleton and make the cells
more susceptible to osmotic swelling and other types of physical
deformation. We also anticipate that decreasing myosin light chain
phosphorylation will alter cell cycle progression. These possibilities
require investigation.
 |
ACKNOWLEDGEMENTS |
We thank Asra Malik for the use of his ECIS Biosensor.
 |
FOOTNOTES |
This research was supported by National Institutes of Health (NIH)
Grants HL-45674 (to M. E. O'Donnell), HL-33009 (to N. Wang), CA-45548
(to D. Ingber), GM-38838 (to E. Elson), and HL-02411 and HL-59618 (to
P. de Lanerolle) and by the Harriet Brooks Fund of the University of
Illinois at Chicago. S. Cai and L. Pestic-Dragovich were supported by
NIH Training Grant HL-076922.
This work was done during P. de Lanerolle's tenure as the Florence and
Arthur Brock Established Investigator of the Chicago Lung Association.
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: P. de Lanerolle, Dept. of Physiology and
Biophysics, University of Illinois at Chicago, 835 S. Wolcott, Chicago,
IL 60612.
Received 26 March 1998; accepted in final form 21 July 1998.
 |
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