From the Musculoskeletal Research Laboratory,
Department of Orthopaedics and Rehabilitation, The Pennsylvania State
University College of Medicine, Hershey, Pennsylvania 17033, § Department of Pharmacology and Physiology, MCP-Hahneman
School of Medicine, Drexel University, Philadelphia, Pennsylvania
19129, ¶ Biomechanical Engineering Division, Department of
Mechanical Engineering, Stanford University, Stanford, California
94305, and
Rehabilitation Research and Development
Center, Palo Alto Health Care System, Department of Veterans Affairs,
Palo Alto, California 94304
Received for publication, October 27, 2000, and in revised form, January 4, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Recently fluid flow has been shown to be a potent
physical stimulus in the regulation of bone cell metabolism. However,
most investigators have applied steady or pulsing flow profiles rather than oscillatory fluid flow, which occurs in vivo because
of mechanical loading. Here oscillatory fluid flow was demonstrated to
be a potentially important physical signal for loading-induced changes in bone cell metabolism. We selected three well known biological response variables including intracellular calcium
(Ca2+i), mitogen-activated protein
kinase (MAPK) activity, and osteopontin (OPN) mRNA levels to
examine the response of MC3T3-E1 osteoblastic cells to oscillatory
fluid flow with shear stresses ranging from 2 to Mechanical loading plays an important role in regulating bone
metabolism. Increased mechanical loading increases bone formation and
decreases bone resorption (1). The absence of mechanical stimulation
causes reduced bone matrix protein production, mineral content, and
bone formation, as well as an increase in bone resorption (2). However,
the mechanism by which bone cells sense and respond to their physical
environment is still poorly understood. In this study we examine a
novel physical stimulus and loading-induced oscillatory fluid flow and
demonstrate that when applied to cultured osteoblastic cells at levels
expected to occur in vivo it regulates mRNA levels for
an important bone matrix protein, osteopontin (OPN).1 Furthermore, this
regulation occurs via an increase in intracellular calcium
(Ca2+i) and mitogen-activated
protein kinases (MAPKs).
The sensitivity of bone tissue to mechanical loading has been proposed
to involve a variety of cellular biophysical signals including
loading-induced electric fields, matrix strain, and fluid flow. The
latter effect of loading, originally described by Piekarski et
al. (3), has recently been proposed to directly regulate bone cell
metabolism in vivo (4, 5). Furthermore, relative to other
loading-induced biophysical signals applied to cells in
vitro, fluid flow appears to be significantly more potent at
physiological levels (6-10). The origin of loading-induced fluid flow
is a consequence of the fact that a significant component of bone
tissue is unbound fluid. Bone tissue contains an extracellular fluid
compartment that has been demonstrated to communicate with the vascular
compartment, and mechanical loading has been shown to enhance fluid
exchange between the two spaces (11).
When bone is exposed to mechanical loading fluid in the matrix is
pressurized and tends to flow into haversian canals. As loading is
removed (e.g. during the gait cycle) the pressure
gradients, and consequently the direction of fluid flow, are reversed
resulting in a flow-time history experienced by the cells that is
oscillatory in nature. In vitro experiments have shown fluid
flow to have a number of effects on bone cells including
Ca2+i mobilization (12), production
of nitric oxide and prostaglandin E2 (8, 13), and
regulation of the expression of genes for OPN, Cyclooxygenase-2, and
c-Fos (9, 14, 15). However, it is important to note that only one study to date utilized a reversing flow profile and found significantly different results when contrasted with nonreversing flow
(16). Thus, the aim of this study is to detail important aspects of the
biochemical response pathway including immediate, intermediate, and
long term effects of oscillatory fluid flow on bone cells, as well as
on their inter-relationships.
To achieve this goal, we first investigated three well known biological
osteogenic response variables.
Ca2+i, a known second messenger
transducing extracellular signals to the cell interior, was our
immediate response variable. Activity of MAPKs is important for
regulating cell differentiation and apoptosis by transmitting
extracellular signals to the nucleus (17, 18) and was our intermediate
response variable. OPN is characterized as one of the predominant
noncollagenous proteins that accumulate in the extracellular matrix of
bone (19, 20) and is also believed to be an important factor associated with bone remodeling caused by mechanical stress in vivo
(21). Recently, strong evidence suggests that OPN is an important
factor in loading induced bone cell metabolism (22-24). Furthermore,
the role of osteopontin in extracellular matrix is more than
structural. It has been shown to be involved in regulating bone cell
attachment, osteoclast function, and mineralization, suggesting a
central role in both the initiation and regulation of bone remodeling (25, 26). Therefore, we quantified steady-state OPN mRNA levels as
a long term response to oscillatory flow.
Recently MAPK family members including extracellular signal-regulated
kinase (ERK), c-Jun N-terminal kinase (JNK), and p38 MAP kinase have
been shown to be important signaling components linking mechanical
stimuli to cellular responses, including cell growth, differentiation,
and metabolic regulation, in endothelial cells, smooth muscle cells,
and myocytes (27-30). However, the role of MAPKs in bone cell
mechanotransduction has not been determined. Moreover the role of
Ca2+i in osteogenic gene
transcription is unclear, especially in the case of oscillatory fluid
flow. Therefore, the second goal of this study is to elucidate the
roles of Ca2+i and the three major
MAPKs in bone cell osteopontin gene expression induced by oscillatory flow.
Finally, the mechanism responsible for fluid-flow-induced
Ca2+i mobilization has not been
fully established, particularly for the oscillatory flow profiles
expected to occur in vivo. Yellowley et al. (31)
demonstrated that the steady flow-induced
Ca2+i responses in bovine articular
chondrocytes involved both influx of external Ca2+ and
release of internal Ca2+ from IP3-sensitive
stores and that the mechanism is G-protein-activated. Similar results
were observed in bone cells stimulated by steady fluid flow (14, 32).
However there is evidence to suggest that steady and oscillatory fluid
flow may have different biophysical effects on bone cells (16).
Therefore, the third goal of this study is to elucidate the mechanism
contributing to oscillatory flow-induced
Ca2+i mobilization in bone cells.
Steady flow (32), substrate stretch (33), and whole bone loading experiments (34) suggest that either stretch-activated mechanosensitive channels and/or L-type voltage-operated calcium channels (L-type VOCCs)
may be involved. Additionally, it is not known whether the involvement
of the IP3-sensitive stores is as important in the response
to oscillatory fluid flow or whether other internal pathways (the
ryanodine-sensitive pathway) may be involved in Ca2+i mobilization.
Cell Culture--
The mouse osteoblastic cell line MC3T3-E1 was
cultured in minimal essential medium (MEM- Oscillatory Fluid Flow Device--
Two different parallel plate
flow chamber sizes were utilized. Larger chambers with a rectangular
fluid volume of 56 × 24 × 0.28 mm were employed for long
term flow to accommodate the larger glass slides. This size of slide
was necessary to obtain adequate amounts of cell protein and mRNA.
The smaller chamber design, fluid volume 38 × 10 × 0.28 mm,
was employed in the calcium imaging studies where total cell number is
not an issue. The oscillatory flow device was described in our previous
study (16). Briefly, a Hamilton glass syringe was mounted in a small
servopneumatic loading frame (EnduraTec, Eden Prairie, MN). The flow
rate was monitored with an ultrasonic flowmeter with a 100-Hz frequency response (Transonic Systems Inc., Ithaca, NY).
Calcium Imaging--
Intracellular calcium ion concentration
([Ca2+]i) was quantified with the
fluorescent dye fura-2. fura-2 exhibits a shift in absorption when
bound to Ca2+ such that the emission intensity when
illuminated with ultraviolet light increases with calcium concentration
at a wavelength of 340 nm and decreases with calcium concentration at
380 nm. The ratio of light intensity between the two wavelengths
corresponds to calcium concentration. A calibration curve of intensity
ratio and calcium concentration was obtained using fura-2 in buffered calcium standards supplied by the manufacturer (Molecular Probes, Inc.,
Eugene, OR).
Preconfluent (80%) cells were washed with MEM-
Cell ensembles were illuminated at wavelengths of 340 and 380 nm in
turn. Emitted light was passed through a 510-nm interference filter and
detected with an intensifier charge coupled device camera
(International Ltd., Sterling, VA). Images were recorded, one every
2 s, and analyzed using image analysis software (Metafluor; Universal Imaging, West Chester, PA). Basal
[Ca2+]i was sampled for 3 min and
followed by 3 min of oscillatory fluid flow (peak shear stress 2 N/m2, 1 Hz).
MAPK Activity Assay--
There are three major MAPKs, p38 MAPK,
ERK, and JNK. 100 µg of lysate protein from either control or
flowed cells was immunoprecipitated with anti-p38 MAPK, anti-ERK1/2, or
anti-JNK antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA)
overnight. Following addition of 15 µl of protein A/G for 2 h,
the immunocomplex was collected by centrifugation, and the kinase
reaction was then conducted in a kinase reaction buffer containing
substrates myelin basic protein (for p38 MAPK or ERK) or c-Jun
glutathione S-transferase (for JNK) in the presence of
[ Osteopontin mRNA Analysis--
The steady-state osteopontin
mRNA level was quantified by quantitative real time reverse
transcription polymerase chain reaction (QRT RT-PCR) (9). Briefly, this
technique is based on the detection of a fluorescent signal produced by
an OPN-specific oligonucleotide probe during PCR primer extension
(Prism 7700 sequence detection system; Applied Biosystems, Frost City,
CA). The RNeasy mini kit (Qiagen Inc., Valencia, CA) was used to
extract total RNA after lysis and homogenization with the QIAshredder
mini column system (Qiagen Inc., Valencia, CA). Mouse osteopontin
cDNA primers and probes were designed using sequence data from
Miyazaki et al. (37) (GenBankTM accession number
X51834) and the QRT RT-PCR probe/primer design software Primer
Express (version 1.0; Applied Biosystems, Frost City, CA). The
fluorogenic oligonucleotide probe for mouse osteopontin was 5'-CGG TGA
AAG TGA CTG ATT CTG GCA GCT C-3' (Synthetic Genetics, San Diego,
CA). The forward and reverse PCR primers were 5'-GGC ATT GCC TCC TCC
CTC-3', and 5'-GCA GGC TGT AAA GCT TCT CC-3', respectively. These
sequences were synthesized, and PCR conditions were optimized with
respect to concentrations of Mg2+, probe, and both primers.
Relative changes in the levels of OPN mRNA and 18 S rRNA were
quantified 24 h after mechanical stimulation.
Pharmacological Agents--
The following series of
pharmacological agents was used to examine the mechanism of calcium
mobilization: thapsigargin (50 nM), gadolinium chloride (10 µM), nifedipine (20 µM), ryanodine (1 and
20 µM), 2-aminoethoxydiphenyl borate (2APB; 100 mM), U73122 and U73343 (4 or 5 µM).
Thapsigargin is an inhibitor of the ATP-dependent
Ca2+ pump of intracellular Ca2+ stores that
causes Ca2+ discharge (38) and was used to empty the
intracellular calcium stores. Gadolinium chloride (10 µM)
(Aldrich) is a putative stretch-activated channel blocker (39).
Nifedipine is a blocker of the L-type VOCC (40). Ryanodine, which
affects ryanodine-sensitive channels in intracellular calcium stores,
was used in two concentrations, 1 µM, which is expected
to hold the channel open, and 20 µM, which is expected to
block the channel (41, 42). U73122 inhibits the action of phospholipase
C and possibly phospholipase A2 and thereby the production
of IP3. Thus, it inhibits the release of calcium through
IP3-sensitive intracellular calcium stores (14, 43).
U73343, an isoform of U73122 that does not inhibit IP3
production, was used as a control. 2APB is a specific inhibitor of the
IP3 receptor and does not affect ryanodine-sensitive or membrane calcium channels (44, 45). Cells were pretreated with medium
containing the required drug for 30-60 min prior to flow, and the drug
remained present during the flow experiments.
Nifedipine, ryanodine, and 2APB were dissolved in 100% ethanol to give
a final concentration of ethanol in the flow medium of 0.1% (v/v), and
vehicle controls were conducted with the same concentration of ethanol.
Thapsigargin, U73122, and U73343 were dissolved in
Me2SO to give a final concentration of
Me2SO in the flow medium of 0.0032, 0.17, and 0.17%
(v/v), respectively. Gadolinium chloride was directly dissolved in medium.
Thapsigargin and gadolinium chloride were also used in long term flow
experiments to examine the role of
[Ca2+]i in downstream responses.
For the MAPK investigations, cells were incubated with MAPK inhibitor
for 2 h before the fluid flow experiments were performed. The p38
inhibitor SB203580 (10 µM) or the ERK inhibitor PD98059
(10 µM; Calbiochem-Novabiochem) was also present in the
flow medium. All pharmacological agents were from Sigma unless indicated.
Data Analysis--
We used a numerical procedure from mechanical
analysis, known as Rainflow cycle counting, to identify calcium
oscillations (46). Briefly, this technique identifies complete cycles
or oscillations in the time history data even when they are
superimposed upon each other and therefore can be used to distinguish
and quantify [Ca2+]i responses
from background noise. We defined a response as an oscillation in
[Ca2+]i at least 2-fold greater
than that of the average baseline level of nontreated cells. Baseline
[Ca2+]i data were recorded for
each slide for 3 min prior to the application of oscillatory fluid flow.
Data were expressed as mean ± S.E. To compare observations from
no flow and flow responses a two-sample Student's t test
was used in which sample variance was not assumed to be equal. To compare observations from more than two groups, a one-way analysis of
variance was employed followed by a Bonferroni selected pairs multiple
comparisons test (Instat; GraphPad Software Inc., San Diego, CA).
p < 0.05 was considered statistically significant. For
calcium experiments all controls were combined as no effect of vehicles
was found (one-way analysis of variance).
Ca2+i Responses to Oscillatory Fluid
Flow--
Typical cell Ca2+i
responses are shown in Fig.
1A. The fraction of MC3T3-E1
cells responding with an increase in
Ca2+i to oscillatory fluid flow
(peak shear stress 2 N/m2, 1 Hz) is shown in
Fig. 1B. The data were obtained from six individual experiments (slides) and a total of 334 cells. Within 30 s of starting oscillatory flow, 59.1 ± 4.6% of cells increased
[Ca2+]i, which was significantly
different from no flow periods (8.9 ± 1.6%). However the
responding cell [Ca2+]i amplitudes
(65.5 ± 17.5 nM) for flow periods were not
statistically different from those for no flow periods (86.5 ± 18.3 nM).
MAPK Responses to Oscillatory Fluid Flow--
The time courses of
activation of three major MAPKs in MC3T3-E1 cells are shown in Fig.
2. At each time point cells from two slides were combined to yield sufficient protein for the MAPK activity
assay. In the absence of flow there was minimal p38, ERK1/2, and JNK
activity. However, dramatic responses for p38 and ERK1/2 activities
were observed beginning 15 min after applying oscillatory flow. p38
activity reached a maximum at 30 min and returned to initial levels 90 min after the onset of oscillatory flow (Fig. 2). ERK1/2 activity
reached a maximum at 60 min and returned to its pre-flow value at 90 min. In contrast, there was no change in JNK activity during a 90-min
flow period, indicating a selective activation of p38/ERKs in response
to flow.
OPN Responses to Oscillatory Fluid Flow--
The long time frame
biological response, steady-state OPN mRNA level, was quantified in
response to oscillatory fluid flow at 1 Hz, resulting in a wall shear
stress of 2 N/m2, utilizing QRT RT-PCR. The
cells that experienced oscillatory flow or no flow for 2 h were
then incubated for an additional 24 h prior to collection. Our
results show oscillatory fluid flow increased steady-state osteopontin
mRNA levels by 3.96 ± 0.76-fold over no flow control (see
Fig. 3; p < 0.05).
Role of Ca2+i and MAPK Activities in the
OPN mRNA Response to Oscillatory Fluid Flow--
To assess the
role of Ca2+i in the OPN mRNA
response to oscillatory fluid flow, cells were subjected to oscillatory
fluid flow in the presence of 50 nM thapsigargin. Interestingly thapsigargin completely blocked the oscillatory flow
effect on steady-state OPN mRNA levels (0.93 ± 0.08 × no flow levels), which were not statistically different from those for
no flow period (Fig. 3). However, gadolinium chloride
(GdCl3; 10 µM) did not attenuate the flow
effect on steady-state OPN mRNA levels (4.19 ± 0.21 × no
flow levels), which were not statistically different from those for
flow control case.
Based on the MAPK activation results, two MAPK inhibitors were employed
to block the activity of ERK1/2 and p38. Cells were exposed to 10 µM of the p38 inhibitor SB203580 (SB) for 2 h prior to and for the duration of oscillatory fluid flow. SB reduced the
effect of fluid flow on steady-state OPN mRNA levels to 1.61 ± 0.50 × no flow levels. Similar results of ERK1/2 inhibitor PD98059 (PD; 10 µM) were obtained with a reduction of
steady-state OPN mRNA to 1.76 ± 0.21 × no flow levels.
Moreover the presence of both inhibitors (SB + PD) completely abolished
the effect (0.84 ± 0.05 × no flow levels; not statistically
different). Those results suggest that activation of p38 MAPK and ERKs
is synergistically involved in flow-mediated OPN expression.
Sources of Ca2+ Mobilization in Response to Oscillatory
Fluid Flow--
The number of cells that responded to oscillatory
fluid flow with a change in intracellular calcium in the presence of
GdCl3 was not significantly different from control
(62.2 ± 3.4%; see Fig.
4A). In contrast to the
results for GdCl3, nifedipine, an L-type VOCC blocker, did
reduce the number of cells responding to flow. The number of cells
responding in the presence of nifedipine (29.3 ± 13.6%) was as
low as in the no flow case, and the mean increase in
[Ca2+]i over baseline (35.6 ± 2.9 nM) was lower than in flow controls.
On application of thapsigargin, which emptied intracellular stores,
there was a significant (p < 0.05) decrease in the
percentage of cells responding to oscillatory flow. U73122, which
inhibits production of IP3 via the phospholipase C pathway
and does not affect membrane channels, reduced the number of cells
responding to 18.0 ± 9.0%, compared with 48.0 ± 9.5% in
the control group. 2APB, which acts directly on IP3
receptors (45) rather than on IP3 catalysis, blocked the
response completely. Ryanodine at 1 µM had a small but
statistically significant effect on the number of cells responding,
which was reduced to 41.6 ± 9.9%. It also had a small effect on
the mean increase in [Ca2+]i over
baseline, which was reduced to 37.4 ± 5.5 nM. At 20 µM, at which concentration the ryanodine-sensitive
channel should have been blocked, there was a very small and not
statistically significant reduction in the number of cells responding,
to 55.1 ± 10.4%. Although there were some differences in the
mean response amplitudes (Fig. 4B), these were not found to
be statistically significant. Some drug-treated cells (nifedipine and
ryanodine) showed higher responses in the no flow period compared with
controls (data not shown), because the drugs caused increased
spontaneous calcium oscillations and increased drift in the baseline levels.
Although a large number of in vitro studies have been
aimed at discovering the regulatory effect of mechanical loading in bone adaptation, little consensus can be found in the literature regarding the appropriate biophysical signals. For example, bone cells
have been shown to respond with metabolic changes to deformation induced by stretching of the substrate to which they are attached (22,
47-49). However, these studies employed either hyperphysiologic levels
of strain or systems known to induce mechanical effects other than pure
strain (50). More recent studies have suggested that bone cells are
more responsive to the fluid flow induced by mechanical strain than
directly to the strain in the tissue (7-9). However, loading-induced
fluid flow in vivo involves a reversal of flow direction
associated with the cyclical unloading that occurs in the vast majority
of physical activities. To date, the ability of the resulting
oscillatory flow profiles to regulate bone cell behavior in
vitro has not been investigated beyond its ability to mobilize
cytosolic calcium (16). In this study a novel oscillatory fluid flow
system was designed to demonstrate that oscillatory fluid flow is
capable of regulating bone cell gene expression via ERK and p38 MAPK
activity and intracellular calcium signaling involving
IP3-mediated calcium release. Additionally, we were able to
demonstrate some potentially important differences in the
characteristics of the response of bone cells to oscillatory flow when
contrasted with published experiments on steady/pulsatile flow. The
study of the effects of oscillatory fluid flow on bone cells will allow
us to more accurately understand the mechanism of mechanotransduction
in bone cells in vivo, for which the other in
vitro systems may not be as suitable.
Our experimental data have shown that oscillatory fluid flow induced
three biological responses that are believed to be important in the
response of bone tissue to mechanical load. In the short term, within 2 min of the start of oscillatory flow, 59.1 ± 4.6% of cells
increased [Ca2+]i, which was
significantly different from the no flow period. This is consistent
with prior observations of the Ca2+i
response of bovine aortic endothelial cells (51), articular
chondrocytes (52), and bone cells (12, 16) to steady/pulsatile fluid
flow. However, oscillatory flow appears to be significantly less
stimulatory than steady/pulsatile flow for bone cells in terms of the
Ca2+i response (16). This suggests
that the mechanotransduction pathways induced by oscillatory flow could be different in part or in whole from those activated by
steady/pulsatile flow.
Recently MAPK activity has been shown to be modulated by various
external stimuli such as growth factors, cytokines, and physical stresses (ultraviolet radiation, hyperosmolarity, hypoxia, and fluid
flow shear stress) (17, 18, 53) and is known to play a pivotal role in
a variety of cell functions. Our results are the first to examine the
regulation of MAPK activity in response to biophysical stimulation in
bone cells. We show that fluid flow induces an increase in the activity
of two of three major MAPKs (ERKs and p38) over a period of 2 h
for bone cells. p38 activity started to increase at 15 min and reached
a maximum at 30 min and then returned to initial levels 90 min after
the onset of oscillatory flow. A similar pattern was observed for
ERK1/2 activity with some delay, at 60 min it reached a peak and
returned to its pre-flow value at 90 min. JNK activity was unchanged
during 90 min of oscillatory fluid flow stimulus. Our biphasic time
course ERK1/2 and p38 activity results are consistent with previous
studies in endothelial cells and smooth muscle cells (27, 28). However, our time to peak ERK1/2 activity (60 min) is slower than observed for
steady fluid flow (5 min), possibly because of the different mechanical
stimuli. Another difference is that oscillatory fluid flow did not
induce JNK activity in bone cells; however steady fluid flow is capable
of activating JNK in endothelial cells within 60 min (27). Although
possibly because of differences between the cell types, it may be a
result of differences in the effects of the physical signals applied.
This is consistent with the possibility that oscillatory fluid shear
stress may stimulate different mechanotransduction pathways from
steady/pulsatile fluid shear stress.
It was suggested previously that the ERK pathway is involved in the
regulation of cell proliferation and differentiation whereas p38 and
JNK are important signaling pathways in the regulation of cell
apoptosis (54). However, recent information demonstrated that p38 MAPK
may also play a critical role in the regulation of differentiation (55,
56). In this study, both the p38 inhibitor SB and the ERK1/2 inhibitor
PD were applied to determine whether the increased MAPK activity we
observed was required for the effect of oscillating flow on
steady-state OPN mRNA levels. Either MAPK inhibitor alone was found
to greatly attenuate (80%) the flow effect on steady-state OPN
mRNA, whereas the presence of both inhibitors (SB + PD) completely
abolished the effect of flow on steady-state OPN mRNA levels. This
indicates that oscillatory flow-induced OPN expression involves both
ERK and p38 MAPK activity with mild redundancy but does not require JNK
activity. It is interesting to note that JNK activity has been observed
in endothelial cells in response to the steady flow associated with
apoptosis (27). In contrast, bone cells experiencing more moderate
oscillatory shear stress exhibit increased ERK1/2 activity associated
with proliferation and differentiation but no change in JNK activity. These findings support the view that oscillatory fluid flow may be a
potent cellular physical signal in bone remodeling in
vivo.
Our results also suggest that the biochemical mechanism of
Ca2+i mobilization is different
between nonreversing steady/pulsatile fluid flow and oscillatory flow. The results of the calcium experiment using nifedipine show that the
L-type VOCC membrane channel is involved in the calcium response to
oscillatory flow in contrast to steady flow experiments in primary bone
cells (32) and in the same cell line (14). However our data are in
agreement with substrate stretch experiments on primary osteoblasts in
which the calcium response was inhibited by nifedipine
(33). In those experiments fluid flow may have been induced in the
system, as well as substrate stretch (50). Thus, it is possible
that the Ca2+i response that the
investigators observed was because of the pathway we describe here in
response to oscillatory flow. Furthermore, the nitric oxide and
prostaglandin E2 response of loaded whole rat bones in an
in vivo model has been shown to be eliminated by nifedipine
(34). This is again consistent with the view that oscillatory flow,
rather than steady flow, is the cellular physical signal that regulates
the adaption of bone to mechanical load in vivo.
Our data also suggest that the stretch-activated membrane
channel, blocked using gadolinium chloride, is not important for response to oscillatory fluid flow. This is in contrast to data for
steady flow where calcium responses were inhibited by blocking this
channel (14, 32). However, our finding that GdCl3 did not
influence steady-state OPN gene mRNA is consistent with the results
of Chen et al. (14) that showed that the effect of steady flow on cytoskeletal reorganization and Cyclooxygenase-2
mRNA involved IP3-mediated intracellular calcium
release but not extracellular calcium. One interpretation is that both
oscillatory and steady flow activate an IP3 cascade that is
important in bone adaptation; however steady flow also stimulates an
GdCl3-sensitive calcium influx whereas oscillatory flow
does not.
Our finding that thapsigargin completely blocked the calcium response
to oscillatory flow demonstrated that the source of Ca2+ is
release from intracellular stores. The next series of experiments were
designed to further elucidate the mechanism of this release. The
combination of the U73122 and the 2APB data strongly suggest that the
IP3 pathway is involved. We achieved a partial block of the
calcium response using U73122. This may be because there are other
pathways to the formation of IP3 besides the phospholipase C and phospholipase A2 pathways blocked by U73122. However the effect of U73122 is shown to be maximal at 10 µM, and
we used only 4-5 µM, because we found that the
concentration of solvent (Me2SO) necessary to achieve the
higher concentration of the drug induced cell toxicity. 2APB, a novel
IP3 blocker that is specific to the IP3 channel
(44), resulted in a total block of the Ca2+ response. This
finding is consistent with our previous observations using neomycin
sulfate (57) and published results from other laboratories supporting
the involvement of this second messenger in fluid flow responses in
various cell types and flow regimes (13, 14, 31, 32).
The role of ryanodine-sensitive internal stores in mechanotransduction
has received less attention, although they have been shown to be
present in osteoblasts (58, 59). Our finding that low concentration
ryanodine inhibited the response of MC3T3-E1 cells to oscillatory flow
confirms the presence of ryanodine-sensitive calcium channels. In our
experiments the opening of the ryanodine-sensitive stores, which would
cause calcium to leave the stores before the flow was applied, in a
similar way to thapsigargin, did have an inhibitory effect on the
response, though less than that of thapsigargin. However the blocking
of the ryanodine-sensitive channel with high concentration ryanodine
had no significant effect on the calcium response. This suggests that
ryanodine-sensitive Ca2+ stores, which can also be
mobilized by the IP3 pathway (60), were partially depleted
by the low concentration ryanodine but that ryanodine-sensitive
channels were not affected by oscillatory fluid flow.
Interestingly, our finding that the source of the calcium response is
IP3-mediated release from intracellular stores seems to be
contradicted by our finding that the L-type VOCC is also involved. If
the VOCC is important to a calcium response mechanism one might expect
to observe some residual calcium increase of extracellular origin, even
in the presence of blockers of intracellular stored calcium. However,
in our study both 2APB and thapsigargin totally abolished the calcium
response to oscillatory fluid flow. Similar results were found in the
Walker et al. substrate stretch study (33) in which
thapsigargin inhibited the calcium response more than would be expected
if only the residual calcium released after nifedipine treatment was
from intracellular stores sensitive to thapsigargin. An explanation for
these results may be that the IP3 receptor on the
endoplasmic reticulum has been shown to be coregulated by cytosolic
calcium concentration. Thus, the VOCC could potentate the fluid flow
calcium response by regulating the local calcium concentration
surrounding the endoplasmic reticulum IP3 receptor but at
levels that are not detectable with our imaging system. This mechanism
would require that the VOCC and endoplasmic reticulum
IP3 receptor are in close association. Such an arrangement has previously been described in muscle cells between the VOCC and
ryanodine-sensitive channels (61).
In the final phase of our investigation we related these intracellular
signaling pathways to the regulation of gene expression. OPN has been
implicated as an important factor in triggering bone remodeling caused
by mechanical stress in vivo (21). Our OPN data are
consistent with the in vitro results of Owan et
al. (7). Steady-state OPN mRNA levels increased almost 4-fold
within 24 h after 2-h oscillatory fluid flow. To elucidate the
role of Ca2+i in bone cell
mechanotransduction and OPN gene regulation, thapsigargin was employed
to empty Ca2+i stores, which
prevents Ca2+i from being available
to the cells during the oscillatory flow period. Thapsigargin
completely abolished the increase in steady-state OPN mRNA levels
that occurred on application of fluid flow. This finding combined with
the role of OPN in mechanically mediated remodeling suggests a
prominent role of cytosolic calcium mobilization in the adaptation of
bone to mechanical loading.
Although our results suggest that both
Ca2+i and MAPK are involved in the
mechanical stress-induced OPN expression in bone cells via oscillatory
flow, the relationships between
Ca2+i and MAPK are still unclear.
Our results show that Ca2+i is
required for OPN expression induced by oscillatory flow. However some
investigators demonstrated that steady flow in chondrocytes activated
ERK1/2 in a way that did not require
Ca2+i, and
Ca2+i alone was not sufficient for
MAPK activation by steady flow (62). Therefore the role of
Ca2+i in MAPK activation under
oscillatory flow remains to be determined. However, both
IP3 and Ca2+i have been
shown to be a necessary step in G-protein-mediated MAPK activation in
smooth muscle cells (63). Little is known about the signaling pathways
between MAPK and target genes, although some investigations have shown
that MAPK phosphatase-1 may act as a mediator to regulate target gene expression in vascular smooth muscle cells (29). Further investigation of the whole cascade of mechanotransduction in bone cells is necessary.
In summary, our study demonstrates that oscillatory fluid flow is a
potent physiological stimulator that induces
Ca2+i release and OPN gene
expression via ERK1/2 and p38 activation but not JNK. OPN gene
expression required Ca2+i
mobilization. Ca2+i is mobilized
using primarily the IP3 pathway, with the L-type VOCC
membrane channel also playing a role. Although we did not compare
oscillatory fluid flow directly to steady/pulsatile flow in this study,
when compared with previously published studies on steady/pulsatile
flow, our findings suggest that there are some potentially important
differences in the response of bone cells to these two stimuli. This
contrast indicates that there may exist multiple mechanotransduction
pathways in bone cells that are activated depending on stimulus type
and that determining an appropriate cellular mechanical stimulus is
critical in understanding the role of mechanical loading in the
regulation of bone.
2
Newtons/m2 at 1 Hz, which is in the range expected
to occur during routine physical activities. Our results showed that
within 1 min, oscillatory flow induced cell
Ca2+i mobilization, whereas two
MAPKs (ERK and p38) were activated over a 2-h time frame. However,
there was no activation of JNK. Furthermore 2 h of oscillatory
fluid flow increased steady-state OPN mRNA expression levels by
approximately 4-fold, 24 h after exposure to fluid flow. The
presence of both ERK and p38 inhibitors and thapsigargin completely
abolished the effect of oscillatory flow on steady-state OPN mRNA
levels. In addition, experiments using a variety of pharmacological
agents suggest that oscillatory flow induces
Ca2+i mobilization via the L-type
voltage-operated calcium channel and the inositol
1,4,5-trisphosphate pathway.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
; Life Technologies,
Inc.) containing 10% fetal bovine serum (FBS; Hyclone, Logan,
UT), 1% penicillin and streptomycin (Life Technologies, Inc.) and
maintained in a humidified incubator at 37 °C with 5%
CO2. All cells were subcultured on glass slides for 2 days
prior to experiments, with the exception of cells cultured for
Ca2+i studies, for which quartz
slides were used, for UV transparency. 3 × 105 cells
were seeded on the glass slides (75 × 38 × 1.0 mm), and 0.85 × 105 cells were seeded on the quartz slides
(76 × 26 × 1.6 mm). There are no significant differences
observed in the behavior of MC3T3-E1 cells grown on normal glass
slides versus quartz
slides.2 It is
important to note that under these conditions the cells had not reached
confluency nor did the medium (which did not include ascorbic
acid or
-glycerophosphate) or time in culture (2 days) support
differentiation or mineralization (35). Cells were exposed to
oscillatory fluid flow in MEM-
and 2% FBS for calcium imaging experiments, and in MEM-
and 10% FBS for long term 2-h experiments.
and 2% FBS at
37 °C, incubated with 10 µM fura-2-acetoxymethyl ester
(Molecular Probes, Inc., Eugene, OR) solution for 30 min at 37 °C,
then washed again with fresh MEM-
and 2% FBS prior to experiments.
-32P]ATP as described before (36). The reaction mix
was subjected to SDS polyacrylamide gel electrophoresis, and
phosphorylation of substrates was determined by autoradiography.
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
View larger version (19K):
[in a new window]
Fig. 1.
A, an example of the MC3T3-E1 cell
[Ca2+]i response traces obtained
for oscillatory flow (2 N/m2, 1 Hz). The
arrow depicts the onset of flow, and each line represents an
individual cell response. B, fraction of MC3T3-E1 cells
responding with an increase in
[Ca2+]i to oscillatory flow.
59.1 ± 4.6% of cells increased
[Ca2+]i for the flow period and
8.9 ± 1.6% of cells increased
[Ca2+]i for the no flow period.
The data were obtained from six individual experiments and a total of
334 cells. *, p < 0.001 versus no flow
control.
View larger version (63K):
[in a new window]
Fig. 2.
Time courses for p38, ERK1/2, and JNK
activation during oscillatory flow (2 N/m2, 1 Hz). At each time point
the cells from two slides were combined to yield sufficient protein for
the MAPK activity assay. Kinase activity was assayed by incubating
lysates with [ -32P]ATP and myelin basic protein (for
p38 MAPK and ERK1/2) or c-Jun glutathione S-transferase (for
JNK). The reaction mix was subjected to SDS polyacrylamide gel
electrophoresis, and phosphorylation of substrates was determined by
autoradiography. A representative autoradiograph is shown. The
experiments were repeated with similar results. The appearance of
double bands is an accepted occurrence with this assay (64, 65) and is
because of impurity, phosphorylation, or degradation of the myelin
basic protein (MBP) substrate but cannot be ascribed to
differential ERK1/2 activity.
View larger version (27K):
[in a new window]
Fig. 3.
The percent changes of cell osteopontin
mRNA levels in response to oscillatory flow (2 N/m2, 1 Hz) in the presence of
different drugs compared with no flow control. Each bar
represents the mean ± S.E., and each experiment was repeated on 3 slides (n = 3). *, p < 0.05 versus no flow control.
View larger version (22K):
[in a new window]
Fig. 4.
Effect of intracellular Ca2+
store modulators on the Ca2+i response to
oscillatory flow (2 N/m2, 1 Hz) in
MC3T3-E1 cells. A, mean percentage of cells showing a
spontaneous Ca2+ transient in absence of flow (No
Flow Control), a response to an oscillatory flow (Flow
Control), and the presence of 50 nM thapsigargin, 10 µM GdCl3, 20 µM nifedipine, 1 µM ryanodine, 20 µM ryanodine, 100 mM 2APB, 4-5 µM U73343 and U73122 (no
statistically significant difference was noticed between 4 and 5 µM). Each bar represents the mean ± S.E., and each experiment was repeated on 40, 40, 6, 6, 8, 10, 10, 7, 8, and 9 slides, respectively. #, p < 0.001 versus flow control. **, p < 0.01 versus U73343. B, mean increase in
[Ca2+]i in cells showing a
spontaneous Ca2+ transient in response to an oscillatory
flow (Flow Control) and the presence of 50 nM
thapsigargin, 10 µM GdCl3, 20 µM nifedipine, 1 µM ryanodine, 20 µM ryanodine, 4-5 µM U73343 and U73122.
Each bar represents the mean ± S.E., and each
experiment was repeated on 40, 6, 6, 8, 10, 10, 8, and 9 slides,
respectively. *, p < 0.05 versus flow
control.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Deborah Grove for designing primers and completing the QRT RT-PCR protocols.
![]() |
FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grants AR45989, AG13087, AG00811, and AG17021, by the Whitaker Foundation, Arthritis Foundation, and the United States Army Medical Research and Materiel Command Award DAMD 17-98-1-8509.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. Section 1734 solely to indicate this fact.
** To whom correspondence should be addressed: Biomechanical Engineering Division, Durand 211, Stanford University, Stanford, CA 94305-3030. Tel.: 650-723-3610; Fax: 650-725-1587; E-mail: christopher.jacobs@stanford.edu.
Published, JBC Papers in Press, January 26, 2001, DOI 10.1074/jbc.M009846200
2 Unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: OPN, osteopontin; Ca2+i, intracellular calcium; MAPK(s), mitogen-activated protein kinase(s); ERK(s), extracellular signal-regulated kinase(s); JNK, c-Jun N-terminal kinase; IP3, inositol 1,4,5-trisphosphate; VOCC(s), voltage-operated calcium channels; MEM, minimal essential medium; FBS, fetal bovine serum; [Ca2+]i, Intracellular calcium ion concentration; QRT, quantitative real time; RT, reverse transcription; PCR, polymerase chain reaction; 2APB, 2-aminoethoxydiphenyl borate; SB, SB203580; PD, PD98059.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Morey, E. R., and Baylink, D. J. (1978) Science 201, 1138-1141[Medline] [Order article via Infotrieve] |
2. |
Sessions, N. D.,
Halloran, B. P.,
Bikle, D. D.,
Wronski, T. J.,
Cone, C. M.,
and Morey-Holton, E.
(1989)
Am. J. Physiol.
257,
E606-E610 |
3. | Piekarski, K., and Munro, M. (1977) Nature 269, 80-82[Medline] [Order article via Infotrieve] |
4. | Weinbaum, S., Cowin, S. C., and Zeng, Y. A. (1994) J. Biomech. 27, 339-360[Medline] [Order article via Infotrieve] |
5. | Cowin, S. C., Weinbaum, S., and Zeng, Y. (1995) J. Biomech. 28, 1281-1297[CrossRef][Medline] [Order article via Infotrieve] |
6. | McLeod, K. J., Donahue, H. J., Levin, P. E., and Rubin, C. T. (1991) in Electromagnetics in Biology and Medicine (Brighton, C. T. , and Pollack, S. R., eds) , pp. 111-115, San Francisco Press, San Francisco |
7. | Owan, I., Burr, D. B., Turner, C. H., Qiu, J., Tu, Y., Onyia, J. E., and Duncan, R. L. (1997) Am. J. Physiol. 42, C810-C815 |
8. |
Smalt, R.,
Mitchell, T.,
Howard, R. L.,
and Chambers, T. J.
(1997)
Am. J. Physiol.
273,
E751-E758 |
9. | You, J., Yellowley, C. E., Donahue, H. J., Zhang, Y., Chen, Q., and Jacobs, C. R. (2000) J. Biomech. Eng. 122, 387-393[CrossRef][Medline] [Order article via Infotrieve] |
10. | Hung, C. T., Allen, F. D., Pollack, S. R., and Brighton, C. T. (1996) J. Biomech. 29, 1403-1409[CrossRef][Medline] [Order article via Infotrieve] |
11. | Knothe Tate, M. L., Niederer, P., and Knothe, U. (1998) Bone 22, 107-117[CrossRef][Medline] [Order article via Infotrieve] |
12. | Hung, C. T., Pollack, S. R., Reilly, T. M., and Brighton, C. T. (1995) Clin. Orthop. 313, 256-269[Medline] [Order article via Infotrieve] |
13. | Ajubi, N. E., Klein-Nulend, J., Alblas, M. J., Burger, E. H., and Nijweide, P. J. (1999) Am. J. Physiol. 276, E171-E178[Medline] [Order article via Infotrieve] |
14. |
Chen, N. X.,
Ryder, K. D.,
Pavalko, F. M.,
Turner, C. H.,
Burr, D. B.,
Qiu, J.,
and Duncan, R. L.
(2000)
Am. J. Physiol.
278,
C989-C997 |
15. |
Pavalko, F. M.,
Chen, N. X.,
Turner, C. H.,
Burr, D. B.,
Atkinson, S.,
Hsieh, Y. F.,
Qiu, J.,
and Duncan, R. L.
(1998)
Am. J. Physiol.
275,
C1591-C1601 |
16. | Jacobs, C. R., Yellowley, C. E., Davis, B. R., Zhou, Z., Cimbala, J. M., and Donahue, H. J. (1998) J. Biomech. 31, 969-976[CrossRef][Medline] [Order article via Infotrieve] |
17. |
Seger, R.,
and Krebs, E. G.
(1995)
FASEB J.
9,
726-735 |
18. | Kyriakis, J. M., Banerjee, P., Nikolakaki, E., Dai, T., Rubie, E. A., Ahmad, M. F., Avruch, J., and Woodgett, J. R. (1994) Nature 369, 156-160[CrossRef][Medline] [Order article via Infotrieve] |
19. |
Denhardt, D. T.,
and Guo, X.
(1993)
FASEB J.
7,
1475-1482 |
20. | Gerstenfeld, L. C., Uporova, T., Ashkar, S., Salih, E., Gotoh, Y., McKee, M. D., Nanci, A., and Glimcher, M. J. (1995) Ann. N. Y. Acad. Sci. 760, 67-82[Medline] [Order article via Infotrieve] |
21. | Terai, K., Takano-Yamamoto, T., Ohba, Y., Hiura, K., Sugimoto, M., Sato, M., Kawahata, H., Inaguma, N., Kitamura, Y., and Nomura, S. (1999) J. Bone Miner. Res. 14, 839-849[Medline] [Order article via Infotrieve] |
22. | Toma, C. D., Ashkar, S., Gray, M. L., Schaffer, J. L., and Gerstenfeld, L. C. (1997) J. Bone Miner. Res. 12, 1626-1636[Medline] [Order article via Infotrieve] |
23. | Harter, L. V., Hruska, K. A., and Duncan, R. L. (1995) Endocrinology 136, 528-535[Abstract] |
24. | Kubota, T., Yamauchi, M., Onozaki, J., Sato, S., Suzuki, Y., and Sodek, J. (1993) Arch. Oral Biol. 38, 23-30[Medline] [Order article via Infotrieve] |
25. | Reinholt, F. P., Hultenby, K., Oldberg, A., and Heinegard, D. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 4473-4475[Abstract] |
26. | Giachelli, C. M., and Steitz, S. (2000) Matrix Biol. 19, 615-622[CrossRef][Medline] [Order article via Infotrieve] |
27. |
Jo, H.,
Sipos, K.,
Go, Y. M.,
Law, R.,
Rong, J.,
and McDonald, J. M.
(1997)
J. Biol. Chem.
272,
1395-1401 |
28. |
Yan, C.,
Takahashi, M.,
Okuda, M.,
Lee, J. D.,
and Berk, B. C.
(1999)
J. Biol. Chem.
274,
143-150 |
29. |
Li, C.,
Hu, Y.,
Mayr, M.,
and Xu, Q.
(1999)
J. Biol. Chem.
274,
25273-25280 |
30. |
Liang, F.,
and Gardner, D. G.
(1999)
J. Clin. Invest.
104,
1603-1612 |
31. | Yellowley, C. E., Jacobs, C. R., and Donahue, H. J. (1999) J. Cell. Physiol. 180, 402-408[CrossRef][Medline] [Order article via Infotrieve] |
32. | Hung, C. T., Allen, F. D., Pollack, S. R., and Brighton, C. T. (1996) J. Biomech. 29, 1411-1417[CrossRef][Medline] [Order article via Infotrieve] |
33. | Walker, L. M., Publicover, S. J., Preston, M. R., Said Ahmed, M. A., and El Haj, A. J. (2000) J. Cell. Biochem. 79, 648-661[CrossRef][Medline] [Order article via Infotrieve] |
34. | Rawlinson, S. C., Pitsillides, A. A., and Lanyon, L. E. (1996) Bone 19, 609-614[CrossRef][Medline] [Order article via Infotrieve] |
35. | McCauley, L. K., Koh, A. J., Beecher, C. A., Cui, Y., Decker, J. D., and Franceschi, R. T. (1995) J. Bone Miner. Res. 10, 1243-1255[Medline] [Order article via Infotrieve] |
36. |
Zhen, X.,
Uryu, K.,
Wang, H. Y.,
and Friedman, E.
(1998)
Mol. Pharmacol.
54,
453-458 |
37. |
Miyazaki, Y.,
Setoguchi, M.,
Yoshida, S.,
Higuchi, Y.,
Akizuki, S.,
and Yamamoto, S.
(1990)
J. Biol. Chem.
265,
14432-14438 |
38. | Thastrup, O., Cullen, P. J., Drobak, B. K., Hanley, M. R., and Dawson, A. P. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 2466-2470[Abstract] |
39. | Hamill, O. P., and McBride, D. W., Jr. (1996) Pharmacol. Rev. 48, 231-252[Abstract] |
40. | Ferrante, J., and Triggle, D. J. (1990) Biochem. Pharmacol. 39, 1267-1270[Medline] [Order article via Infotrieve] |
41. |
Meissner, G.
(1986)
J. Biol. Chem.
261,
6300-6306 |
42. | Hasselbach, W., and Migala, A. (1987) FEBS Lett. 221, 119-123[CrossRef][Medline] [Order article via Infotrieve] |
43. | Bleasdale, J. E., Thakur, N. R., Gremban, R. S., Bundy, G. L., Fitzpatrick, F. A., Smith, R. J., and Bunting, S. (1990) J. Pharmacol. Exp. Ther. 255, 756-768[Abstract] |
44. | Maruyama, T., Kanaji, T., Nakade, S., Kanno, T., and Mikoshiba, K. (1997) J. Biochem. (Tokyo) 122, 498-505[Abstract] |
45. | Hamada, T., Liou, S. Y., Fukushima, T., Maruyama, T., Watanabe, S., Mikoshiba, K., and Ishida, N. (1999) Neurosci. Lett. 263, 125-128[CrossRef][Medline] [Order article via Infotrieve] |
46. | Jacobs, C. R., Yellowley, C. E., Nelson, D. V., and Donahue, H. J. (2000) Comput. Methods Biomech. Biomed. Eng. 3, 31-40[Medline] [Order article via Infotrieve] |
47. | Rodan, G. A., Bourret, L. A., Harvey, A., and Mensi, T. (1975) Science 189, 467-469[Medline] [Order article via Infotrieve] |
48. | Buckley, M. J., Banes, A. J., Levin, L. G., Sumpio, B. E., Sato, M., Jordan, R., Gilbert, J., Link, G. W., and Tran Son Tay, R. (1988) Bone Miner. 4, 225-236[Medline] [Order article via Infotrieve] |
49. | Brighton, C. T., Strafford, B., Gross, S. B., Leatherwood, D. F., Williams, J. L., and Pollack, S. R. (1991) J. Bone Jt. Surg. Am. 73, 320-331[Abstract] |
50. | Brown, T. D. (2000) J. Biomech. 33, 3-14[CrossRef][Medline] [Order article via Infotrieve] |
51. |
Geiger, R. V.,
Berk, B. C.,
Alexander, R. W.,
and Nerem, R. M.
(1992)
Am. J. Physiol.
262,
C1411-C1417 |
52. |
Yellowley, C. E.,
Jacobs, C. R.,
Li, Z.,
Zhou, Z.,
and Donahue, H. J.
(1997)
Am. J. Physiol.
273,
C30-C36 |
53. |
Davis, R. J.
(1993)
J. Biol. Chem.
268,
14553-14556 |
54. | Johnson, G. L., and Vaillancourt, R. R. (1994) Curr. Opin. Cell Biol. 6, 230-238[Medline] [Order article via Infotrieve] |
55. |
Morooka, T.,
and Nishida, E.
(1998)
J. Biol. Chem.
273,
24285-24288 |
56. | Nebreda, A. R., and Porras, A. (2000) Trends Biochem. Sci. 25, 257-260[CrossRef][Medline] [Order article via Infotrieve] |
57. | Reilly, G. C., Yellowley, C. E., Donahue, H. J., and Jacobs, C. R. (2000) J. Bone Miner. Res. 15 Suppl. 1, 508 |
58. | McDonald, F., Somasundaram, B., McCann, T. J., Mason, W. T., and Meikle, M. C. (1996) Arch. Biochem. Biophys. 326, 31-38[CrossRef][Medline] [Order article via Infotrieve] |
59. |
Adebanjo, O. A.,
Biswas, G.,
Moonga, B. S.,
Anandatheerthavarada, H. K.,
Sun, L.,
Bevis, P. J.,
Sodam, B. R.,
Lai, F. A.,
Avadhani, N. G.,
and Zaidi, M.
(2000)
Am. J. Physiol.
278,
F784-F791 |
60. | Khodakhah, K., and Armstrong, C. M. (1997) Biophys J. 73, 3349-3357[Abstract] |
61. |
Weigl, L. G.,
Hohenegger, M.,
and Kress, H. G.
(2000)
J. Physiol.
525,
461-469 |
62. | Hung, C. T., Henshaw, D. R., Wang, C. C., Mauck, R. L., Raia, F., Palmer, G., Chao, P. H., Mow, V. C., Ratcliffe, A., and Valhmu, W. B. (2000) J. Biomech. 33, 73-80[CrossRef][Medline] [Order article via Infotrieve] |
63. |
Chin, T. Y.,
and Chueh, S. H.
(1998)
Am. J. Physiol.
275,
C1255-C1263 |
64. |
Rao, G. N.,
and Runge, M. S.
(1996)
J. Biol. Chem.
271,
20805-20810 |
65. |
Guyton, K. Z.,
Liu, Y.,
Gorospe, M.,
Xu, Q.,
and Holbrook, N. J.
(1996)
J. Biol. Chem.
271,
4138-4142 |