Effects of microtubules and microfilaments on
[Ca2+]i and contractility
in a reconstituted fibroblast fiber
Kazuo
Obara1,
Koji
Nobe1,
Hiromi
Nobe1,
Michael S.
Kolodney2,
Primal
De
Lanerolle3, and
Richard J.
Paul1
1 Departments of Physiology and Biophysics, University of
Cincinnati College of Medicine, Cincinnati, Ohio 45267;
3 College of Medicine, University of Illinois at
Chicago, Chicago, Illinois 60612; and 2 Dermatology Division,
Department of Medicine, University of California at Los Angeles, Los
Angeles, California 90095
 |
ABSTRACT |
We used a reconstituted fiber formed when 3T3
fibroblasts are grown in collagen to characterize nonmuscle
contractility and Ca2+ signaling. Calf serum (CS) and
thrombin elicited reversible contractures repeatable for >8 h. CS
elicited dose-dependent increases in isometric force; 30% produced the
largest forces of 106 ± 12 µN (n = 30), which
is estimated to be 0.5 mN/mm2 cell cross-sectional
area. Half times for contraction and relaxation were 4.7 ± 0.3 and 3.1 ± 0.3 min at 37°C. With imposition of constant shortening velocities, force declined with time, yielding
time-dependent force-velocity relations. Forces at 5 s fit the
hyperbolic Hill equation; maximum velocity
(Vmax) was 0.035 ± 0.002 Lo/s.
Compliance averaged 0.0076 ± 0.0006 Lo/Fo. Disruption of microtubules with nocodazole in a CS-contracted fiber had no net effects on force, Vmax, or stiffness; force increased in 8, but
decreased in 13, fibers. Nocodazole did not affect baseline
intracellular Ca2+ concentration
([Ca2+]i) but reduced (~30%) the
[Ca2+]i response to CS. The force after
nocodazole treatment was the primary determinant of stiffness and
Vmax, suggesting that microtubules were not a
major component of fiber internal mechanical resistance. Cytochalasin D
had major inhibitory effects on all contractile parameters measured but
little effect on [Ca2+]i.
cytochalasin D; nocodazole; nonmuscle mechanics; Swiss 3T3; tensegrity; intracellular calcium concentration
 |
INTRODUCTION |
THE ROLE OF CYTOSKELETAL
FILAMENT networks in modulating nonmuscle contractility is
unclear. Giuliano and colleagues (7) have proposed that
nonmuscle contractility is regulated both by modulating the activity of
molecular motors, such as myosin II, and by altering the cytomatrix in
such a manner as to either resist or yield to the tension applied by
the motors. This hypothesis is supported by the observation that
tension increases in response to disruption of microtubules by
nocodazole (5) in cultured fibroblasts (4)
and reconstituted fibroblast fibers (21). These data are
also consistent with the view that microtubules provide some form of
internal resistance. In tensegrity models (6), a portion
of a cell's contractile force is borne by rigid internal structures,
reducing the force transmitted to external structures, as monitored by
force transducers. Alternatively, other mechanisms may underlie the
effects of nocodazole. For example, the increase in smooth muscle
contractility elicited by nocodazole has been correlated with an
increase in intracellular Ca2+ concentration
([Ca2+]i) (17).
If microtubules constitute an internal resistance in parallel with the
actin-myosin network and are capable of bearing compressive forces,
depolymerization of microtubules should result in a greater measured
force as the load shifts from internal microtubule structures to the
external force transducer. One would also expect that other mechanical
parameters, such as cell mechanical stiffness and shortening velocity,
would be sensitive to the presence of an intracellular mechanical
resistance as that postulated for microtubules. In fact, on theoretical
grounds, one might anticipate that the maximum shortening velocity
(Vmax) would be particularly sensitive to intracellular resistances.
With the use of similar arguments, the effects of microfilament
assembly in fibroblast contractility could also be deduced by studying
the mechanical effects of cytochalasin D, known to disrupt
microfilaments (3).
In this study, we developed a reconstituted fibroblast fiber,
based on previous models (19, 21), that
enables not only force to be quantitated but also velocity and
stiffness in nonmuscle cells. We report that disruption of
microfilaments has a profound effect on fibroblast contractility but
not [Ca2+]i. Our detailed
mechanical analysis, in contrast to hypotheses based on measurement of
isometric force alone, does not support a role for microtubule
resistance in the mechanical properties of reconstituted fibroblast fibers.
 |
MATERIALS AND METHODS |
Cell culture and reconstitution of fibroblast into fibers.
Swiss 3T3 fibroblast cells (passages 15-30) were
cultured in Dulbecco's modified Eagle's medium (DMEM) plus 10% fetal
calf serum (CS) and antibiotic-antimycotic (Ab/Am; 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B) at 37°C in
a humidified, 5% CO2-95% O2 atmosphere.
Fibroblast fibers were formed by growing 3T3 fibroblasts in rat tail
collagen (Upstate Biotechnology, Lake Placid, NY) gel matrix by a
modification of the method of Kolodney and Wysolmerski
(21). Dispersed cells were suspended in an ice-cold
collagen solution that contained 2 × 106 cells/ml and
0.5 mg/ml rat tail collagen in DMEM + 10% CS with added Ab/Am
(Sigma). An aliquot (2 ml) of the collagen/cell suspension was poured
into a well (0.8 × 5 cm × 0.5 cm deep) cut in a layer of
silicone rubber in 100-mm glass petri dishes and placed in a
CO2 incubator at 37°C. After 8 h, an additional 1 ml
of DMEM + 10% CS and Ab/Am were added to each well. The
preparations were incubated for 3-5 days or until the cells shrank
the gel and formed a fiber. Except where noted, cells were placed in
serum-free media the night preceding the day on which experiments were conducted.
Mechanics measurements.
The fibers made from 3T3 fibroblasts were cut into 5-mm pieces and
mounted between glass posts with a cyanoacrylate glue. One post was
fixed and the other connected to an AME 801 silicon strain gage
(SensoNor) force transducer. The fibers were bathed in physiological
salt solution (MOPS-PSS) that contained (in mM): 140 NaCl, 4.7 KCl, 1.2 NaH2PO4, 0.02 EDTA, 1.2 MgSO4, 2.5 CaCl2, 5.5 dextrose, and 20 MOPS, pH 7.4, at 37°C.
For measurement of shortening velocity and stiffness, fibers (5-mm)
were glued between the force transducer and a lever arm of which its
position, and therefore the length of fiber, was controlled by a
Cambridge Technology (Cambridge, MA) ergometer. Force-velocity
relations were measured by imposing constant shortening velocities on
the fiber, and measurement of the subsequent force. A series of eight
different velocities at 60-s intervals was used for measurement of each
individual force-velocity relation. Force-velocity data were fitted
with the Hill equation, (F + a) · (V + b) = b · (Fo + a), using a nonlinear
least-squares routine (Origin). Vmax was
taken as the velocity at zero force. Stiffness was measured by
imposition of rapid (<1 ms) shortening and stretching steps (
2.6 to
+0.6% Lo). A series of eight step changes at 60-s
intervals constituted an experimental set for the measurement of stiffness.
Measurement of the [Ca2+]i.
Measurements of [Ca2+]i levels were obtained
from cells loaded with the Ca2+-sensitive fluorescent dye,
fura 2, based on the techniques of Grynkiewicz et al.
(10). The fibroblast fibers were loaded with fura 2-AM.
Fura 2-AM was prepared as a stock solution of 1 mM dye in DMSO. The
fura 2 loading solution contained 3 µM fura 2-AM, 0.015% Pluronic
F-127, and 0.5% DMSO in MOPS-PSS buffer. To aid in dispersion of the
fura 2-AM, the loading solution was well sonicated. Fibroblast fibers
were placed in this solution at room temperature for 3 h. After
this loading period, fibroblasts were washed in MOPS-PSS buffer for 20 min to remove any unesterified dye.
A segment of fura 2-loaded fibroblast fiber was placed in a glass
bottom culture dish and covered with nylon mesh to maintain isometric
conditions. The fiber chamber had a total volume of 500 µl, which was
perfused (5 ml/min) with the MOPS-PSS, to maintain a temperature of
37°C. [Ca2+]i was measured with an
Intracellular Imaging (Cincinnati, OH) microscope-based system (InCa
system). This fibroblast chamber was placed on a Nikon Diaphot inverted
microscope with a fluorphase objective. Fluorescent images of cells
excited at 340 and 380 nm and emitted at 510 nm were obtained with a
Dage silicon-intensified target camera. After subtraction of background
fluorescence, the 340 and 380 nm images were ratioed on a pixel by
pixel basis, and the ratios converted to
[Ca2+]i using a standard curve. Solutions
containing known concentrations of free Ca2+ (Molecular
Probes) were used to generate this standard curve. Fluorescence
intensity was measured in 150 µl for each standard solution (0, 0.065, 0.100, 0.225, 0.351, and 0.602 µM free Ca2+
concentration) containing 13.3 µg/ml fura 2 pentapotassium salt. Quantitative analysis of the average subcellular Ca2+ was
performed by defining the outline of the cell, summing the Ca2+ in all the pixels within the defined area, and
dividing by the number of pixels.
Confocal fluorescence microscopy.
For phalloidin staining, fibers were fixed with 4% paraformaldehyde
and embedded in 17% gelatin. Sections were cut at 100 µm using a
Leica VT 1000 vibrating blade microtome and stained with
rhodamine-conjugated phalloidin (Sigma, St. Louis, MO). Images were
constructed from optical sections acquired by scanning confocal microscopy.
For microtubule staining, fibers were fixed with 4% paraformaldehyde,
and 100-µm sections were cut with a Leica VT 1000 vibrating blade
microtome. Sections were blocked with 5% goat serum and incubated
overnight in a 1:100 dilution of monoclonal antibody to
-tubulin
(Amersham) followed by FITC-conjugated goat anti-mouse (Zymed, San
Francisco, CA) at 1:50. Images were acquired with an Olympus
epifluorescence microscope equipped with a high-resolution charge-coupled device camera.
Data analysis.
All data are presented as means ± SE. Control and serum data were
pooled and include some data previously reported (23). To
assess the effects of cytochalasin D and nocodazole, a control contraction was elicited by 30% serum and the mechanical parameters were measured. After treatment with the drugs, the mechanical parameters were again measured. Paired comparisons, with each fiber
serving as its own control, were made to control for variability among
fibers. Differences were analyzed with paired t-tests;
differences with a P value <0.05 were accepted as
statistically significant.
 |
RESULTS |
Structure of artificial fibers.
The formation of a fiber generally takes 3 days after the cell/collagen
suspension is poured into the mold. This transformation is shown in
Fig. 1. The final fiber has
dimensions of ~40 mm long and 1-2 mm diameter. For contractility
measurements, each fiber was cut into smaller segments (~5 mm). These
segments are about one-eighth of the whole fiber and would, if
proportional, contain ~5 × 105 of the original
cells. Histological analysis of sections indicated that the cells were
generally aligned along the long axis of the fiber. A more detailed
structural analysis of these types of fibers at both aggregate and
intracellular levels has been reported (21).

View larger version (175K):
[in this window]
[in a new window]
|
Fig. 1.
Artificial fibers reconstituted from Swiss 3T3
fibroblasts. Fibroblast cells reaggregate to form a fiberlike
preparation. Fibroblast cells (2 × 106 cells/ml) were
cultured in collagen (0.5 mg/ml) medium matrix, as described in
MATERIALS AND METHODS. Photograph shows a 100-mm petri dish
in which 3 wells (0.8 × 4 cm × 0.5 cm deep) were cut in a
layer of silicone rubber, previously cast in the dish. At 24-h
intervals, an aliquot (2 ml) of the collagen/cell suspension was poured
into 1 well and placed in a CO2 incubator at 37°C.
Photograph was taken at 48 h so that the total incubation time was
48 h for mold 1, 24 h for mold 2, and
30 min for mold 3.
|
|
Isometric contraction.
Fibers were mounted under isometric conditions, then the length was
increased to match the original fiber length in the mold. After stress
relaxation, force attained a baseline level of 53.0 ± 0.003 µN
(n = 30). Addition of serum increased isometric force that returned to prestimulus levels on washout (Fig.
2). These contraction/relaxation cycles
could be repeated for at least 8 h, though over prolonged periods
some stress relaxation in baseline force was noted. We could not
determine a length dependence of force generation as classically done
for striated muscle (9) because these preparations did not
sustain stable forces when lengthened beyond the length formed in the
mold. If untethered, the fibers shortened to approximately one-third
the original formed length. Thus, at shorter lengths, the force would
decrease. Serum induced dose-dependent contractions (Fig.
3), and a maximum isometric force of
106 ± 12 µN (n = 30) was developed in response
to 30% serum. The times to half-maximal contraction and relaxation
were 4.7 ± 0.3 and 3.1 ± 0.3 min (n = 22),
respectively. Thrombin (2 U/ml) also elicited an increase in force,
averaging 19.0 ± 2.0 µN (n = 13) above the
prestimulus baseline.

View larger version (10K):
[in this window]
[in a new window]
|
Fig. 2.
Serum-induced contraction of a reconstituted fibroblast fiber.
A: After mounting in the isometric apparatus, fiber length
was increased to its original length at formation in the mold (ST),
left scale. After stress relaxation, the force scale was
changed (right scale). Contraction was induced by 30% calf
serum (CS). PSS indicates a return to the original physiological salt
solution. Rapid spikes on solution changes are due to surface tension
changes. B: a continuation of A. The
serum-induced contraction was reversible, and the
contraction-relaxation cycle was reproducible for at least 8 h.
|
|

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 3.
Contractile response of a reconstructed fibroblast fiber
to CS. Mean dose-response curve for 10 fibers. Isometric force was
normalized to the maximum value for each fiber. Maximum force was
101.8 ± 15.1 µN (n = 10). Fibers were incubated
for short (30-60 min, ; n = 5) or long
terms (>15 h, ; n = 10) in serum-free
medium before force measurements. Inset: isometric force
developed in response to cumulative addition of CS to a reconstituted
fiber. PSS indicates a return to the original physiological salt
solution.
|
|
As noted in previous studies (21), cytochalasin D (1 µM)
reduced force in a serum-induced contracture to levels below the initial prestimulus levels (Fig. 4). The
force remaining after cytochalasin D treatment was ~20% of the total
force in a maximum serum-induced contracture. Because cytochalasin D
would disrupt actin-containing filament, forces due to both
actin-myosin motors, analogous to "active force" or "tone" in
smooth muscle, and cytoskeleton-related "passive" force, would be
reduced. Thus an upper bound for the active force under unstimulated
conditions would be 17.5% of the force developed on serum stimulation.

View larger version (9K):
[in this window]
[in a new window]
|
Fig. 4.
Effect of cytochalasin D (CytoD) on serum-induced
contraction of a reconstituted fibroblast fiber. Contraction was
induced by 30% CS and then 1 µM CytoD was added.
|
|
Our normal protocol was to place the fiber in serum-free medium the
night before mechanical measurements. We investigated whether this
serum-free period was required for the observed contractile responses.
Test fibers were kept in serum until they were mounted for force
measurements in serum-free media. After attainment of a steady
baseline, which required between 30 and 60 min in serum-free conditions, isometric force was measured as a function of serum concentration. Figure 3 shows that the cumulative dose-response curves
for fibers incubated for short (30-60 min) or long terms (>15 h)
in serum-free conditions were identical. These experiments indicate
that a prolonged serum-free period is not required by these fibers for
contractile responses.
Force-velocity relations.
These relations were studied by imposing a series of constant speed
decreases in length and measurement of the consequent force responses.
Figure 5A shows a typical
experimental record for a single fiber composed of individual force
responses to eight imposed shortenings of varying speeds. Force can be
seen to continuously decline with the duration of shortening, similar
to behavior reported for rat aorta (22) and hog coronary
artery (17). Thus there is a family of force-velocity
relations (Fig. 5B), each corresponding to the point in time
at which the force values were measured. In the inset to Fig.
5B, Vmax taken from each Hill
equation was plotted as a function of the point in time at which the
forces were measured. Vmax decreases rapidly at
first, then is relatively stable between 3 and 5 s. The velocities
measured at early time points may in fact include discharge of series
mechanical compliances, before a quasisteady state is
achieved. In subsequent experiments, the force at 5 s was
taken for all force-velocity relations. This point in time was chosen
as it provided the widest range of measurable values over the differing
experimental conditions. This provides a means for determining an
operational, relative Vmax for comparison of the
different conditions.

View larger version (29K):
[in this window]
[in a new window]
|
Fig. 5.
Force-velocity relations of reconstituted fibroblast
fibers. A: original recording of force in response to the
imposition of constant shortening velocities in a serum-stimulated
contracture. Length step, L/Lo. B:
force-velocity curve. Force (P) at 1 ( ), 2 ( ),
3 ( ), 4 ( ), and 5 ( ) s
normalized to the isometric force (Po) was plotted against
the imposed shortening velocity. Inset: time dependency of
maximum shortening velocity (Vmax).
Vmax taken from each Hill equation was plotted
against time at which the forces were measured. Symbols in A
have same meaning as in B.
|
|
Force-velocity relations were fitted with the Hill equation, (F + a) · (V + b) = b · (Fo + a), using a nonlinear least-squares routine
(Origin). For maximum serum-induced contractures, Fo was taken as the initial total force, and Vmax, the
velocity at zero force, averaged 0.035 ± 0.002 Lo/s
(n = 30); in the example shown in Fig. 5B,
this was 0.030 Lo/s, where Lo was the initial
length of the fiber. This velocity is in the range reported for tonic smooth muscles (24) and is considerably slower than
striated muscle. a/Fo, a dimensionless curvature parameter,
averaged 1.11 ± 0.07. This parameter, often inversely related to
efficiency, is higher than generally observed for striated muscle
(0.25) but is similar to that reported for the phasic rat portal vein
(13). At the highest imposed shortening speeds, some
compressive forces (negative forces) were measured. There does not
appear to be any discontinuity in the force-velocity relations in this
range. For unstimulated fibers (n = 30), these
parameters averaged 0.015 ± 0.001 Lo/s and 1.96 ± 0.20, respectively.
Fiber stiffness.
Stiffness was measured by imposition of rapid (<1 ms) step changes in
length. Typical responses are shown in Fig.
6A. After imposition of the
step, the force response exhibited a peak value followed by stress
relaxation. A plot of the peak force responses against the imposed step
length change is shown in Fig. 6B. Typical of muscle
behavior, the fibroblast peak force responses were linear in the region
where stretches were imposed, and this linear range extended to short
step decreases. In the serum-stimulated fibers, the slope of the linear
relation between force and length (stiffness) was 23.4 ± 0.92 mN/Lo (n = 30). Extrapolation of the linear
portion gives an intercept on the length axis of
0.0076 ± 0.0006
L/Lo (n = 30); i.e., a step of
0.76% Lo is required to discharge the maximum isometric
force. This value is lower than isolated smooth muscle cells
(28) but comparable to that reported for skinned smooth
muscle fibers (2).

View larger version (19K):
[in this window]
[in a new window]
|
Fig. 6.
Stiffness measurements in reconstructed fibroblast fibers.
A: time course of changes in length and force in
nonstimulated fiber. B: the change in force plotted against
the magnitude of the imposed length step ( L/Lo) in
nonstimulated ( ) and CS-stimulated ( ) fibers.
The peak force reached during rapid length steps (completed within 1 ms) is plotted against the amplitude of the imposed length steps.
Regression lines (dashed lines) for force change vs. length change for
stretches (positive L/Lo values) are shown in the
figure.
|
|
Effects of nocodazole and cytochalasin D on ultrastructure and
mechanical parameters of reconstituted fibroblast fibers.
We used fluorescence microscopy to verify the ability of cytochalasin D
and nocodazole to disrupt microfilaments and microtubules in these
preparations, respectively. We selected concentrations based on the
reported disruption of microtubules and actin filaments in cultured
fibroblasts (21). As illustrated in Fig.
7A, intact microtubules were
clearly visible in cells populating control fibers, whereas cells from
fibers treated with 10 µM nocodazole for 10 min (Fig. 7B)
exhibited a diffuse distribution of tubulin, indicating
depolymerization of microtubules.

View larger version (53K):
[in this window]
[in a new window]
|
Fig. 7.
Immunofluorescent micrographs of fibroblast fibers
immunolabeled with anti-tubulin. Control (A) and fibers
treated with nocodazole (B; 10 µM) were sectioned and
labeled with a monoclonal antibody to -tubulin. Intact microtubules
were present in cells from control fibers but not in cells from
nocodazole-treated fibers.
|
|
Organization of polymerized actin was assessed by staining cells with
rhodamine-conjugated phalloidin. In control fibers (Fig. 8A), actin filaments were
predominantly distributed at the cell periphery. Cells treated with CS
(Fig. 8B) demonstrated increased staining for polymerized
actin. Cytochalasin D treatment (Fig. 8C) disrupted actin
morphology in CS-treated cells that resulted in clumping of polymerized
actin.

View larger version (120K):
[in this window]
[in a new window]
|
Fig. 8.
Immunofluorescent micrographs of fibroblast fibers
labeled with rhodamine. Polymerized actin was visualized with
rhodamine-conjugated phalloidin control fibers (A), fibers
treated with CS (B), and fibers treated with CS followed by
cytochalasin D (C; 1 µM). Scale bar (A) applies
to all panels. Actin polymerization was enhanced by CS but disrupted by
cytochalasin D.
|
|
With the use of the same concentrations of nocodazole and cytochalasin
D, we assessed the effects of disruption of microtubules and actin
filament networks on the mechanics of the fibroblast fiber. Addition of
nocodazole to fibers precontracted with CS (30%) did not result in any
statistical differences in the mean values, as summarized in Table
1. This average can be somewhat misleading in that there were appreciable increases in force in 8 fibers and decreases in 13. Figure
9A shows the distribution of
the responses of isometric force.

View larger version (26K):
[in this window]
[in a new window]
|
Fig. 9.
Distribution of individual values for force
(A), Vmax (B), and
stiffness (C) under control conditions and in response to
serum, nocodazole (serum + Noco), and CytoD. The responses for
each individual fiber are designated by the same symbol and connected
by dotted lines.
|
|
Disruption of the microtubule network with nocodazole would be expected
to significantly enhance the maximum shortening velocity if intact
microtubules posed a significant internal resistance. The distribution
of Vmax shows similar variability as isometric force (Fig. 9B). Nocodazole had no significant effect on the
average Vmax (Table 1). As shown in Fig.
10, there is, however, a strong correlation between Vmax and force.

View larger version (14K):
[in this window]
[in a new window]
|
Fig. 10.
Effects of CS, nocodazole, and cytochalasin D on the
relation between maximum shortening velocity and force. Force and
Vmax were normalized to those in the resting
state. Nocodazole and cytochalasin D were applied after the
serum-induced contraction reached a plateau. , CS (Ser,
30%); , nocodazole (Noco, 10 µM); ,
cytochalasin D (CytoD, 1 µM). Regression lines for CS (dashed line),
nocodazole (broken line), and cytochalasin D (solid line) are shown.
The level of isometric force, independent of how it is achieved,
appears to be the major determinant of Vmax.
|
|
Fiber stiffness would also be expected to reflect any resistance
attributable to microtubule networks. The distribution of stiffness
values measured is given in Fig. 9C, and the dependence of
stiffness on force is given in Fig. 11.
Nocodazole did not alter the average stiffness (Table 1), but, like
Vmax, the level of force appeared to be the
major determinant of stiffness. Thus the increase in force seen in some
cases with nocodazole treatment was associated with an increase in
resistance. This is the opposite of that expected if reduction of
microtubular mechanical resistance was responsible for the increase in
force.

View larger version (15K):
[in this window]
[in a new window]
|
Fig. 11.
Effects of CS, nocodazole, and cytochalasin D on the
relation between stiffness and force. Force and stiffness were
normalized to those in the resting state. Nocodazole or cytochalasin D
were applied after the serum-induced contraction reached a plateau.
Symbols and regression lines have same meanings as in Fig. 10. Note
that nocodazole has little effect on the relation between stiffness and
force, whereas cytochalasin D elicits major changes.
|
|
In contrast to nocodazole, cytochalasin D had major and consistent
effects on the mechanical parameters studied (Fig. 10, Table 1).
Isometric force was reduced to below the intrinsic levels of force in
unstimulated fibroblasts. Velocity was proportionately reduced (Fig.
10). Stiffness was decreased to ~50% of the initial unstimulated
values; however, the precision of these measurements was reduced due to
the low force remaining after cytochalasin D treatment (Fig. 11).
Effects of nocodazole and cytochalasin D on
[Ca2+]i.
To further delineate the mechanisms underlying the effects of
microtubule or microfilament disruption on contractility, we measured
[Ca2+]i using ratiometric fluorescence
spectroscopy and the Ca2+-sensitive dye fura 2. Nocodazole
(10 µM) elicited a small increase in force with little effect on
[Ca2+]i (Fig.
12). Addition of serum produced a
transient increase in [Ca2+]i that was
smaller (72.1% ± 3.8, n = 8, P = 0.002) than that seen in the absence of nocodazole. The addition of
serum in the presence of nocodazole elicited a force response that was
nearly identical (97.4% ± 0.9, n = 8, P = 0.02) to that in its absence.

View larger version (37K):
[in this window]
[in a new window]
|
Fig. 12.
Effects of nocodazole on force (A) and intracellular
Ca2+ concentration ([Ca2+]i;
B and C) in fibroblast fibers. A: the
effects of nocodazole on basal force and on the force developed in
response to CS. B: fiber [Ca2+]i
measured with fura 2 fluorescence in 2 typical cells. C: the
average value for 9 cells in the fiber.
|
|
Cytochalasin D (1 µM) also had little effect on baseline
[Ca2+]i and reduced (89.2% ± 1.9, n = 7, P < 0.001), but did not
eliminate, the increase in [Ca2+]i in
response to serum (Fig. 13). In
contrast to nocodazole treatment, cytochalasin D treatment nearly
abolished (15.8% ± 0.8, n = 7, P = <0.001) the increase in force in response to addition of serum.

View larger version (39K):
[in this window]
[in a new window]
|
Fig. 13.
Effects of cytochalasin D on force (A) and
[Ca2+]i (B and C) in
fibroblast fibers. A: the effects of cytochalasin D on basal
force and on the force developed in response to CS. B: fiber
[Ca2+]i measured with fura 2 fluorescence in
2 typical cells. C: the average value for 4 cells in the
fiber.
|
|
 |
DISCUSSION |
We have shown that artificial fibers composed of fibroblasts grown
in a culture media including collagen provide a unique model for
measurement of mechanical parameters in nonmuscle cells. Because
cytochalasin D nearly abolishes mechanical responses, these mechanical
parameters reflect aggregate cellular properties rather than those of
the collagen matrix in which the cells are grown. In the absence of
cells, collagen alone does not form fibers or support mechanical loads.
The aggregated cells generate force under unstimulated conditions. This
is demonstrated by the ability of the fiber to shorten and by the
inhibition of force by cytochalasin D. Both serum and thrombin elicit
dose-dependent increases in isometric force. The response to serum of
these Swiss fibroblasts was approximately fivefold greater than
thrombin. The maximum isometric forces measured in this study were on
the order of 150 µN. If one assumes that the original 4 × 106 fibroblasts were evenly distributed throughout the
reconstituted fiber, and were 100 µm long with a 10-µm diameter,
then the cellular cross-sectional area of the fiber can be calculated
as ~0.8 mm2. The actual fiber cross-sectional area is
difficult to determine exactly but is roughly between 1 and 4 mm2. Thus this estimate is not unrealistic and is
comparable to some vascular smooth muscle tissue in which smooth muscle
cells account for 25% of the vessel wall. Thus force per cellular
cross-sectional area would be 0.2 mN/mm2. This is
considerably less than that for smooth muscle, in which tissue values
range from 10 to 200 mN/mm2, and potentially larger when
translated to smooth muscle cellular cross-sectional areas. However,
our calculated value of 0.2 mN/mm2 is of similar magnitude
to those estimated for fibroblasts from wound contraction models in
skin preparations (15).
Hyperbolic force-velocity relations, similar to tonic smooth muscle,
were observed in these reconstituted fibers. As force continuously
declined with time during the imposed constant shortening, assignment
of a maximum velocity depended on the time of measurement. This is
similar to that observed in studies of smooth muscle, in which, for
example, unloaded velocities showed a similar slowing with time.
Whether this is due to some internal resistance, as suggested for
smooth muscle (12), or a possible dependence on length
(11), is not known. However, the maximum velocities
deduced from these force-velocity relations were similar to those
visually measured in unloaded chicken embryo fibroblasts cells
on cutting from the naturally occurring constraints in the forming
molds (21). These maximum velocities are much slower than
striated muscle but similar to those reported for the slower, tonic
smooth muscles, such as the aorta (25). Velocity and
myosin ATPase activities are generally correlated. The low velocity of
these fibroblast fibers is consistent with the low-myosin
ATPase rates reported for nonmuscle myosin. The actin-activated
Mg2+-ATPase reported for clonal mouse
fibroblast myosin is 90 nmol · min
1 · mg
1
(1). This can be compared with 89 nmol · min
1 · mg
1 for
platelet myosin and 51 nmol · min
1 · mg
1 reported
for gizzard smooth muscle measured under similar conditions (26).
In assessment of the data as a whole, the level of isometric force
appeared to be the major determinant of the speed of shortening. In
Fig. 10, the maximum velocities under the various conditions studied
were plotted against the level of isometric force. Whether the
variations in force were due to the natural variations in response to
serum or modified by nocodazole or cytochalasin D, higher levels of
force were strongly correlated with higher velocities.
Fibroblast fiber stiffness was also assessed in this model and was
found to be similar to that reported for smooth muscle cells but
considerably higher than that for isolated smooth muscle tissue. This
may reflect the differences in structures through which force is
transmitted. Stiffness was related to the level of isometric force
(Fig. 11), though this dependence was not as highly correlated as
Vmax and was sharply reduced after cytochalasin D treatment. The relations between stiffness and force were similar for
serum contractures in the presence or absence of nocodazole, which
suggest that microtubules are not a major determinant of this
parameter. The linear dependence of stiffness on force of these cell
aggregates is similar to that observed for individual cells and
predicted by tensegrity models (27).
We tested whether microtubules constituted significant mechanical
resistance or internal load by measuring the effects of nocodazole, an
agent known to disrupt microtubules (16), on shortening
velocity and stiffness. Our immunomicroscopy confirmed that nocodazole
at 10 µM also disrupted microtubule structures in fibroblasts in our
reconstituted fibers (Fig. 7). Nocodazole has been shown in previous
studies to increase isometric force in stimulated fibroblast fibers
(21) that would be consistent with a reduction in internal
resistance opposing the force generated by actin-myosin motors. In
contrast to previous studies, we did not consistently observe an
increase in force in response to nocodazole (Fig. 9A).
However, in our studies, nocodazole was added to cells previously
contracted with CS. Contraction with CS sharply decreased the
additional response to nocodazole in chicken embryo fibroblasts (18). This finding suggests that, in contrast to the
predictions of the tensegrity model, nocodazole and CS may cause
contraction through a common signaling pathway that may become
saturated by CS, thereby diminishing the nocodazole response.
Vmax would be expected to be very sensitive to
such an internal resistance. Based on the measured force-velocity
relations (Fig. 5B), eliminating an internal load with a
magnitude of only 20% of the maximum isometric force would increase
fiber Vmax by 40%. The effects of nocodazole on
Vmax depended on its effects on force, which
were variable. However, there was no net effect of nocodazole on
Vmax of serum-stimulated fibers (Table 1).
The relation between the level of isometric force and
Vmax seen after nocodazole treatment was also
similar to that observed for serum stimulation alone (Fig. 10). This
provides evidence against the presence of a specific mechanical
resistance that can be attributed to microtubules. This is further
supported by the effects of nocodazole on fiber stiffness. When
nocodazole treatment was associated with increased force, stiffness
increased. This is the opposite to expectation if disruption of
microtubules removed an internal resistance. The increase may be
attributed to more actin-myosin cross bridges under these conditions.
This is consistent with recent studies suggesting that nocodazole
treatment increases myosin light chain phosphorylation in fibroblasts
(18).
The effects of nocodazole on both stiffness and
Vmax appear to be mediated through its effects
on force. Larger forces, independent of how they were generated, were
associated with greater Vmax. This would be
consistent with some type of nonmicrotubule internal resistance in the
aggregated cell fiber. If such a resistance was constant, higher forces
would lead to relatively less loading and faster velocities. This would
also be consistent with the decrease in shortening velocity with time,
as has been proposed for smooth muscle cells (12).
In contrast to nocodazole, cytochalasin D, which disrupts actin
filaments (3), had dramatic inhibitory effects on the
mechanical process(es) underlying all measured parameters. The effects
of cytochalasin D on these mechanical parameters was paralleled by its
depolymerization of actin filaments in the fibroblast fiber (Fig. 8).
Thus the ability to develop force and to shorten is highly dependent on
the integrity of the actin filament network. The decreases in both
stiffness and Vmax by cytochalasin D were parallel to that of its inhibition of force and consistent with force
being the primary determinant of these parameters. This would be
consistent with actin-myosin interaction being the primary locus of
force generation in these fibers.
It is also of interest to consider the possibility of linkage between
actin microfilaments and microtubules, as suggested by actin binding
properties of some microtubule-associated proteins (8,
14). If, by some manner, actin and microtubule networks are linked, and nocodazole destroys the linkage decreasing parallel cross bridges, then force, as well as stiffness, would be expected to
decrease. This is what is observed. If these cross bridges are
"lost" to force generation, then this may underlie the loss of
force with nocodazole observed in some cases. On the other hand, if the
loss of parallel cross bridges is translated into more series cross
bridges, then one might anticipate Vmax to
increase, which was not observed. Further speculation in lieu of
high-resolution ultrastructure studies is not warranted. However, these
mechanical data provide the first insight into the relations between
force, stiffness, and shortening velocity in a nonmuscle contractile system.
Because cytochalasin D and nocodazole appear to be mediated via their
effect on force, it was of interest to assess their effects on
[Ca2+]i. The mechanism of activation of
nonmuscle myosin is controversial (20, 23),
but [Ca2+]i is likely a key second messenger.
Treatment with nocodazole reduced the increase in
[Ca2+]i in response to serum by ~30%, but
had little effect on the developed force. Cytochalasin D, which
dramatically reduced force, had an even lesser effect than nocodazole
on the Ca2+ response to serum. Thus it is unlikely that the
effects of either were mediated primarily through effects on
[Ca2+]i.
In summary, reconstituted fibers formed by fibroblasts cultured in a
collagen gel can be used for precise mechanical measurements in
aggregated nonmuscle cells. Although the isometric force generated by
these reconstituted fibers is significantly lower, maximum velocity and
compliance are similar to that of smooth muscle fibers. In terms of the
contractile response to serum, actin microfilaments play a major role
in the mechanical properties of the reconstituted fibers, as might be
expected for a myosin motor-driven system. Microtubules do not make a
major contribution to an internal mechanical resistance or load against
which the myosin system has been proposed to operate (7).
Because the composition of the major components in these cultured cells
can be readily manipulated by genetic techniques (23),
this model system may provide a unique approach to the understanding of
contractile mechanisms in nonmuscle as well as cultured muscle cells.
 |
FOOTNOTES |
Address for reprint requests and other correspondence: R. J. Paul, Dept. of Molecular and Cellular Physiology, Univ. of
Cincinnati College of Medicine, Cincinnati, OH 45267-0576 (E-mail:
Richard.Paul{at}uc.edu).
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.
Received 6 December 1999; accepted in final form 23 March 2000.
 |
REFERENCES |
1.
Adelstein, RS,
Conti MA,
Johnson GS,
Pastan I,
and
Pollard TD.
Isolation and characterization of myosin from cloned mouse fibroblasts.
Proc Natl Acad Sci USA
69:
3693-3697,
1972[Abstract].
2.
Arheden, H,
and
Hellstrand P.
Force response to rapid length change during contraction and rigor in skinned smooth muscle of guinea-pig taenia coli.
J Physiol (Lond)
442:
601-630,
1991[Abstract].
3.
Cooper, JA.
Effects of cytochalasin and phalloidin on actin.
J Cell Biol
105:
1473-1478,
1987[ISI][Medline].
4.
Danowski, BA.
Fibroblast contractility and actin organization are stimulated by microtubule inhibitors.
J Cell Sci
93:
255-266,
1989[Abstract].
5.
DeBrabander, M,
and
DeMey J
(Editors).
Microtubules and Microtubule Inhibitors. Amsterdam: Elsevier, 1985.
6.
Dennerll, TJ,
Joshi HC,
Steel VL,
Buxbaum RE,
and
Heidemann SR.
Tension and compression in the cytoskeleton of PC-12 neurites. II. Quantitative measurements.
J Cell Biol
107:
665-674,
1988[Abstract].
7.
Giuliano, KA,
Kolega J,
DeBiasio RL,
and
Taylor DL.
Myosin II phosphorylation and the dynamics of stress fibers in serum-deprived and stimulated fibroblasts.
Mol Biol Cell
3:
1037-1048,
1992[Abstract].
8.
Gonzalez, M,
Cambiazo V,
and
Maccioni RB.
The interaction of Mip-90 with microtubules and actin filaments in human fibroblasts.
Exp Cell Res
239:
243-253,
1998[ISI][Medline].
9.
Gordon, AM,
Huxley AF,
and
Julian FJ.
The variation in isometric tension with sarcomere length in vertebrate muscle fibres.
J Physiol (Lond)
184:
170-192,
1966[ISI][Medline].
10.
Grynkiewicz, G,
Poenie M,
and
Tsien RY.
A new generation of Ca2+ indicators with greatly improved fluorescence properties.
J Biol Chem
260:
3440-3450,
1985[Abstract].
11.
Gunst, SJ.
Effect of length history on contractile behavior of canine tracheal smooth muscle.
Am J Physiol Cell Physiol
250:
C146-C154,
1986[Abstract/Free Full Text].
12.
Harris, DE,
and
Warshaw DM.
Slowing of velocity during isotonic shortening in single isolated smooth muscle cells. Evidence for an internal load.
J Gen Physiol
96:
581-601,
1990[Abstract].
13.
Hellstrand, P,
and
Johansson B.
Analysis of the length response to a force step in smooth muscle from rabbit urinary bladder.
Acta Physiol Scand
106:
221-238,
1979[ISI][Medline].
14.
Henrquez, JP,
Cross D,
Vial C,
and
Maccioni RB.
Subpopulations of
interact with microtubules and actin filaments in various cell types.
Cell Biochem Funct
13:
239-250,
1995[ISI][Medline].
15.
Higton, DIR,
and
James DW.
The force of contraction of full-thickness wounds of rabbit skin.
Br J Surg
51:
462-466,
1964[ISI].
16.
Jordan, MA,
and
Wilson L.
Use of drugs to study role of microtubule assembly dynamics in living cells.
Methods Enzymol
298:
252-276,
1998[ISI][Medline].
18.
Kolodney, MS,
and
Elson EL.
Contraction due to microtubule disruption is associated with increased phosphorylation of myosin regulatory light chain.
Proc Natl Acad Sci USA
92:
10252-10256,
1995[Abstract].
19.
Kolodney, MS,
and
Elson EL.
Correlation of myosin light chain phosphorylation with isometric contraction of fibroblasts.
J Biol Chem
268:
23850-23855,
1993[Abstract/Free Full Text].
20.
Kolodney, MS,
Thimgan MS,
Honda HM,
Tsai G,
and
Yee HF, Jr.
Ca2+-independent myosin II phosphorylation and contraction in chicken embryo fibroblasts.
J Physiol (Lond)
515:
87-92,
1999[Abstract/Free Full Text].
21.
Kolodney, MS,
and
Wysolmerski RB.
Isometric contraction by fibroblasts and endothelial cells in tissue culture: a quantitative study.
J Cell Biol
117:
73-82,
1992[Abstract].
22.
McMahon, EG,
Martin AF,
and
Paul RJ.
Isomyosins and mechanics in aortas from aldosterone salt hypertensive rats (Abstract).
Fed Proc
46:
1098,
1987[ISI].
23.
Obara, K,
Nikcevic G,
Pestic L,
Nowak G,
Lorimer DD,
Guerriero V, Jr,
Elson EL,
Paul RJ,
and
de Lanerolle P.
Fibroblast contractility without an increase in basal myosin light chain phosphorylation in wild type cells and cells expressing the catalytic domain of myosin light chain kinase.
J Biol Chem
270:
18734-18737,
1995[Abstract/Free Full Text].
24.
Paul, RJ.
The chemical energetics of vascular smooth muscle. Intermediary metabolism and its relation to contractility.
In: Handbook of Physiology. The Cardiovascular System. Vascular Smooth Muscle. Bethesda, MD: Am. Physiol. Soc, 1980, sect. 2, vol. II, chapt. 9, p. 201-235.
25.
Paul, RJ.
Smooth muscle: Mechanochemical Energy Conversion, Relations Between Metabolism and Contractility.
In: Physiology of the Gastrointestinal Tract (2nd ed.), edited by Johnson LR.. New York: Raven, 1987, p. 483-506.
25a.
Paul RJ, Bowman PS, and Kolodney MS. Effects of microtubule
disruption on force, velocity, stiffness, and
[Ca2+]i in porcine coronary artery. Am
J Physiol Heart Circ Physiol. In press.
26.
Sellers, JR,
Pato MD,
and
Adelstein RS.
Reversible phosphorylation of smooth muscle myosin, heavy meromyosin, and platelet myosin.
J Biol Chem
256:
13137-13142,
1981[Abstract/Free Full Text].
27.
Wang, N,
Butler JP,
and
Ingber DE.
Mechanotransduction across the cell surface and through the cytoskeleton.
Science
260:
1124-1127,
1993[ISI][Medline].
28.
Warshaw, DM,
and
Fay FS.
Cross-bridge elasticity in single smooth muscle cells.
J Gen Physiol
82:
157-199,
1983[Abstract].
Am J Physiol Cell Physiol 279(3):C785-C796
0363-6143/00 $5.00
Copyright © 2000 the American Physiological Society