1 Department of Oral Cell
Biology, Bone adapts to mechanical stress, and bone
cell cultures from animal origin have been shown to be highly sensitive
to mechanical stress in vitro. In this study, we tested whether bone
cell cultures from human bone biopsies respond to stress in a similar
manner as animal bone cells and whether bone cells from osteoporotic patients respond similarly to nonosteoporotic donors. Bone cell cultures were obtained as outgrowth from collagenase-stripped trabecular bone fragments from 17 nonosteoporotic donors between 7 and
77 yr of age and from 6 osteoporotic donors between 42 and 72 yr of
age. After passage, the cells were mechanically stressed by treatment
with pulsating fluid flow (PFF; 0.7 ± 0.03 Pa at 5 Hz for 1 h) to
mimic the stress-driven flow of interstitial fluid through the bone
canaliculi, which is likely the stimulus for mechanosensation in bone
in vivo. Similar to earlier studies in rodent and chicken bone cells,
the bone cells from nonosteoporotic donors responded to PFF with
enhanced release of prostaglandin E2
(PGE2) and nitric oxide as well
as a reduced release of transforming growth factor-
prostaglandin E2; nitric
oxide; osteoporosis
THE INCREASING number of osteoporotic fractures makes
osteoporosis an important health risk of our time. The cause of bone loss in patients with osteoporosis is multifactorial, but in all cases
an imbalance between bone resorption and bone formation is the
underlying mechanism (7, 31). Bone mass and the mechanical performance
of the skeleton are affected by a variety of factors, both local and
systemic. Systemic control results from a number of calcium-regulating
hormones such as parathyroid hormone, calcitonin, and vitamin D as well
as growth hormone and sex hormones. Local control is exerted primarily
by mechanical demands resulting from gravity and the stressing of bone
by muscular contraction. Many studies have shown that bone as a tissue
adapts to these mechanical demands to produce an optimized structure in
terms of mass and geometry (10, 13, 16). Bone mass diminishes with
increasing age as a result of changes in circulating levels of
hormones, in particular decreased estrogen levels after menopause (30, 42, 47), but possibly also because of decreased anabolic effects of
mechanical loading as a result of declining levels of physical activity. Osteocytes, osteoblasts, and lining cells are anatomically in
an appropriate position to detect mechanical strain and are extremely
sensitive to mechanical stress in vitro (23). Therefore, these bone
cells are thought to play a role in the response to skeletal loading as
mechanosensors that can transduce the physical stimuli into biochemical
signals (23).
The pathogenesis of osteoporosis is complicated by the fact that it is
a multifactorial disorder. Diseases, drugs, and environmental influences such as chronic liver or kidney disease, an excess of
thyroid hormones, low calcium intake, alcoholism, and smoking appear to
exaggerate the normal bone loss that occurs with aging (34, 37).
Immobilization may also contribute to the loss of bone in osteoporosis
(46). However, the relationship of osteoporosis with an abnormal bone
cell mechanosensitivity has not been studied as far as we know.
Impaired responsiveness of bone cells to mechanical stimuli might lead
to less effective adaptation of the bone tissue and thereby to a
reduced ability to carry the prevalent loads resulting from normal
usage.
The intrinsic capacity of bone cells to respond to mechanical stress
may be studied with the use of cultures of isolated bone cells in
vitro. This approach also allows comparison of bone cells derived from
osteoporotic patients with control donors. Bone cells from younger
control and older osteoporotic donors were found to differ in their
response to cyclic strain, measured as enhanced cell proliferation and
release of transforming growth factor- The aim of this study was to test whether bone cells derived from
osteoporotic patients respond differently to mechanical stress than
cells derived from age-matched controls. To verify their bone cell
phenotype, bone-derived cell cultures were monitored for production of
osteocalcin, a bone-specific protein, and alkaline phosphatase (ALP)
activity under basal and 1,25-dihydroxyvitamin D3
[1,25(OH)2D3]-stimulated
conditions. We then tested the response of the bone cells to mechanical
stress by examining changes in the release of
PGE2, NO, and TGF- Donors.
Transiliac bone biopsies were obtained from 17 control donors (10 males, 7 females) between 7 and 77 yr of age with a mean age of 29 ± 7 yr. They were all without metabolic bone disease and entered the hospital for maxillofacial surgery (cleft palate or
mandibular reconstruction using iliac crest bone) or orthopedic surgery
(elective joint replacement or humerus reconstruction after trauma). In
addition, similar bone biopsies were obtained from six osteoporotic
(OP) donors (1 male, 5 females) with a mean age of 61 yr ranging
between 47 and 72 yr. They were diagnosed as OP by their low bone
mineral density in the lumbar spine
(t-score less than
ABSTRACT
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
(TGF-
). The
upregulation of PGE2 but not the other responses continued for 24 h after 1 h of PFF treatment. The bone
cells from osteoporotic donors responded in a similar manner as the
nonosteoporotic donors except for the long-term PGE2 release. The PFF-mediated
upregulation of PGE2 release
during 24 h of postincubation after 1 h of PFF was significantly
reduced in osteoporotic patients compared with six age-matched controls as well as with the whole nonosteoporotic group. These results indicate
that enhanced release of PGE2 and
nitric oxide, as well as reduced release of TGF-
, is a
characteristic response of human bone cells to fluid shear stress,
similar to animal bone cells. The results also suggest that bone cells
from osteoporotic patients may be impaired in their long-term response
to mechanical stress.
INTRODUCTION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
(TGF-
) (29). Although the
exact mechanism whereby bone cells sense the effect of stress placed on
a bone organ is not known, recent theoretical (5, 48) as well as in
vivo (45) and in vitro experimental (20, 23) evidence agrees that flow
of interstitial fluid is likely involved. Deformation (strain) of bone
tissue as a result of load causes flow of interstitial fluid through
the lacunar-canalicular porosity of bone (32). It has been calculated
that this flow of fluid over the osteocyte surface may trigger a
cellular response (48). Indeed, several in vitro studies have shown
that fluid flow causing a fluid shear stress of 0.5-1 Pa rapidly
increases prostanoid production in animal bone cells (20, 23, 36). In
addition, human (22), mouse (20), and chicken (20) bone cells all
rapidly respond to fluid flow with an enhanced release of nitric oxide
(NO). NO has been implicated in a number of conditions of orthopedic
interest, including inflammation, arthritis, aseptic loosening of joint
prostheses, sepsis, ligament healing, and osteoporosis (8). The
modulation of NO production by cells within the bone microenvironment
may therefore be a sensitive mechanism for local control of bone
remodeling. Prostaglandins, particularly prostaglandin
E2
(PGE2), are important local
regulators of bone metabolism (35) and have been shown to play a
central role in the ability of the skeleton to respond to mechanical
stress (49). Other molecules likely involved in mediating local gain
and loss of bone tissue are locally produced growth factors such as
TGF-
. TGF-
is an abundant noncollagenous protein in bone (41), it
is produced by bone cells (11), and it can modulate the growth and
differentiation of bone cells in vitro. We have recently found that
mouse bone cells react to mechanical stress with changes in TGF-
production (19).
1 in response
to pulsating fluid flow (PFF). Both short-term and long-term responses
were studied, i.e., immediately after 1 h of treatment and 24 h later.
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
2.5) in
combination with at least one vertebral fracture (type I osteoporosis).
Histology showed no mineralization defect in these patients. They were
all treated with calcium (500 mg/day) and vitamin D (400 IU/day), and
one of them had been treated with the bisphosphonate pamidronate for 1 yr. For statistical analysis, the data of the OP group were contrasted
with the non-OP controls of the same age range consisting of six donors
(4 males, 2 females) between 44 and 77 yr of age with a mean age of 67 yr (Table 1). The protocols were approved
by the ethical review board of the Academic Hospital, Vrije
Universiteit, and all donors gave informed consent.
Table 1.
Sex, age, and treatment data of 6 osteoporotic donors and 6 nonosteoporotic donors that served as age-matched control group
Cell culture. Bone cell cultures were established according to earlier described methods (2, 38, 44). Briefly, the trabecular bone specimens were placed in cold sterile PBS and dissected within 1 h after removal. Bone specimens were minced into small fragments, washed extensively and repeatedly with PBS, and incubated with 2 mg/ml of collagenase (type II, Worthington) for 2 h at 37°C in a shaking water bath. The collagenase-treated bone fragments were washed once with medium containing 10% fetal bovine serum (FBS; GIBCO, Paisley, UK) to inhibit collagenase activity and were transferred to 25- or 75-cm2 flasks (Nunc, Roskilde, Denmark) depending on the amount of bone tissue obtained. Roughly 5-10 mg of bone fragments were added per square centimeter of flask surface. The bone fragments were cultured in DMEM (GIBCO) supplemented with 100 U/ml of penicillin (Sigma, St. Louis, MO), 50 µg/ml of streptomycin sulfate (Sigma), 50 µg/ml of gentamycin (GIBCO), 1.25 µg/ml of Fungizone (GIBCO), 100 µg/ml of ascorbate (Merck, Darmstadt, Germany), and 10% FBS. Culture medium was replaced 3 times/wk. Bone cells started to migrate from the bone chips after 4-14 days. When the cell monolayer growing from the bone fragments reached confluence, after 2-5 wk, cells were trypsinized using 0.05% trypsin (Difco Laboratories, Detroit, MI) and 0.02% EDTA (Sigma) in PBS for 5-10 min and plated at 25 × 103 cells/well in six-well culture dishes (Costar, Cambridge, MA) containing 3 ml of DMEM with 10% FBS. Cells were grown until subconfluence, when they were challenged with 1,25(OH)2D3 or PFF.
Characterization of bone cell cultures: response to
1,25(OH)2D3,
expression of von Willebrand factor.
To test their osteoblastic phenotype, cell cultures were incubated for
3 days in the presence or absence of
108 M
1,25(OH)2D3
as follows. The medium of the subconfluent six-well plate cultures (see
above) was replaced by fresh medium containing 2% FBS and
10
8 M vitamin
K1 (Hoffmann-La Roche, Basel,
Switzerland) (27, 33, 44) with or without
10
8 M
1,25(OH)2D3.
Subsequently, osteocalcin was measured in the conditioned medium by
radioimmunoassay (Incstar, Stillwater, MN) with the use of an antibody
raised against bovine osteocalcin. The detection limit amount was 0.2 ng/ml. All osteocalcin values were corrected for the amount of
osteocalcin in medium with 2% FBS. In addition, ALP activity was
determined in the cell lysate by using
p-nitrophenyl phosphate (Merck) as a
substrate at pH 10.3, according to the method as described by Lowry
(25). The assay was performed in 96-well microtiter plates, and the
absorbance was read at 410 nm using a Dynatech MR7000 microplate reader
(Dynatech, Billinghurst, UK).
PFF. PFF was generated with the use of a flow apparatus containing a parallel-plate flow chamber as described earlier (20, 23) (Fig. 1). The apparatus contained 13 ml of medium that was pumped over the cells in a pulsatile manner by a revolving pump. This resulted in a pulsating (5 Hz) fluid shear stress of 0.7 ± 0.03 Pa and an estimated peak stress rate of 12.2 Pa/s. During an experiment, the apparatus was placed in a 37°C incubator and connected to a gassing system that maintained a pH of 7.4 in the medium using 5% CO2 in air. For PFF experiments, cells were trypsinized and plated onto polylysine-coated (50 µg/ml; poly-L-lysine hydrobromide, mol wt 15-30 × 104; Sigma) glass slides, which served as the bottom of the flow chamber. Cells were plated at 5 × 105 cells/glass slide (size 2.5 × 6.5 cm) and preincubated overnight in DMEM with 10% FBS, resulting in a subconfluent monolayer. Then the medium was changed to DMEM supplemented with 2% FBS, antibiotics, and 100 µg/ml of ascorbate, and the cells were incubated for 1 h in the absence (static control) or presence of PFF. Subsequently, the glass slide with the cells was removed from the flow apparatus, a culture well was created around the cells by securing a rectangle silicone rubber ring (Dow Corning, Midland, MI) on the glass slide, 1 ml of fresh culture medium was added, and the cells were postincubated for 24 h in the absence of PFF (post-PFF). Static control cultures were cultured in 13 ml of static medium during the first "PFF" hour and in 1 ml of medium thereafter.
|
NO.
NO was measured as nitrite (NO2)
accumulation in the conditioned media.
NO
2 is the stable end product of NO,
and its concentration has been shown to be a good reflection of NO
production. The amount of NO
2 release
was determined using Griess reagent (12) consisting of 1%
sulfanylamide, 0.1% naphthylethylenediamine dihydrochloride, and 2.5 M
H3PO4.
Serial dilutions of NaNO2 were
used as standard curve. Briefly, 75 µl of conditioned medium were
mixed with 75 µl of Griess reagent and incubated for 15 min at room
temperature under continuous shaking. The assay was performed in
96-well microtiter plates, and the absorbance at 540 nm was determined
using a Dynatech MR 7000 microplate reader.
PGE2. PGE2 concentrations in the conditioned medium were measured by an enzymeimmunoassay system (Amersham, Buckinghamshire, UK) with the use of an antibody raised against mouse PGE2. The detection limit was 16 pg/ml. The absorbance was read at 450 nm using a Dynatech MR 7000 microplate reader.
TGF-.
TGF-
1 concentrations in the conditioned medium were determined by an
ELISA system using an antibody raised against porcine TGF-
1
(Promega, Madison, WI). Total (latent and biologically active) TGF-
1
was measured after thermal activation of the samples for 5 min at
75°C (3). Biologically active TGF-
1 was below the detection
limit of the assay (25 pg/ml). The absorbance was read at 450 nm using
a Dynatech MR7000 microplate reader.
Protein. After 24 h post-PFF, the protein content of the cell layer was measured using a BCA protein assay reagent kit (Pierce, Rockford, IL) (43). The assay was performed in 96-well microtiter plates, and the absorbance at 570 nm was determined using a Dynatech MR7000 microplate reader.
Statistical analysis. Results are expressed as means ± SE. The effects of treatment with 1,25(OH)2D3 were analyzed using a paired two-tailed Student's t-test. The effects of treatment with PFF were analyzed using Wilcoxon's signed rank test because the data did not meet the requirements for normal distribution. The response of the OP group was contrasted with the non-OP control group using the Mann-Whitney U test. Differences were considered significant at P < 0.05.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cells became visible as outgrowth of the collagenase-stripped bone chips after 1 or 2 wk in culture. They formed a subconfluent layer within 2-3 wk, when they were passaged. The speed of cell outgrowth (measured as time of first visible cell outgrowth, time of monolayer subconfluence) was somewhat faster in the younger donors below 44 yr of age than in the older donors. The cell growth characteristics of the OP group were not different from the non-OP control group (data not shown).
Three days of treatment with
108 M
1,25(OH)2D3
increased ALP activity and osteocalcin release in all non-OP control
donors (Table 2) as well as in the OP group
(Fig. 2). Basal as well as
1,25(OH)2D3-stimulated
ALP activity was similar in the non-OP control group [basal, 6 ± 3 nmol p-nitrophenol
(PNP) · h
1 · µg
protein
1;
1,25(OH)2D3-treated
cultures, 14 ± 6 nmol
PNP · h
1 · µg
protein
1] and in the
OP group [basal, 7 ± 3 nmol
PNP · h
1 · µg
protein
1;
1,25(OH)2D3-stimulated
conditions, 15 ± 6 nmol
PNP · h
1 · µg
protein
1]. The same
applied to osteocalcin release [non-OP control group: basal, 93 ± 41 ng/mg protein;
1,25(OH)2D3treated
cultures, 287 ± 81 ng/mg protein; OP group: basal, 145 ± 93 ng/mg protein;
1,25(OH)2D3-treated cultures, 284 ± 140 ng/mg protein]. Immunostaining for vWF
using the monoclonal vWF antibody did not show any positive cells in the same cell cultures (data not shown).
|
|
Application of mechanical stress by PFF for 1 h did not result in
permanent changes in cell shape or alignment of the cells in the
direction of the flow and did not affect cell viability as determined
by trypan blue exclusion (data not shown). The amount of protein
per cell culture was also not affected when measured 24 h after PFF
treatment (non-OP donors: static controls, 85 ± 11 µg;
PFF-treated cultures, 82 ± 8 µg; OP group: static controls, 91 ± 11 µg; PFF-treated cultures, 97 ± 12 µg). Treatment with PFF did modulate the release of
PGE2, NO, and TGF- in the
non-OP donors. After 1 h of PFF treatment, the release of
PGE2 and NO had roughly doubled,
whereas the release of TGF-
was reduced (Table
3). The release of
PGE2 was still upregulated during
24 h postincubation after PFF treatment, but the releases of NO and TGF-
were no longer different from nonstressed cell cultures (Table 3).
|
Last, we studied the responses to PFF by cells from OP donors and
compared them with six non-OP donors of the same age group. OP cells
responded to 1 h of PFF with an increased release of PGE2, the same as the non-OP
control group (Fig.
3A; non-OP
control group: PFF, 68 ± 33 ng
PGE2/mg protein; +PFF, 230 ± 152 ng PGE2/mg protein,
P < 0.05; OP group:
PFF, 14 ± 7 ng PGE2/mg protein; +PFF,
27 ± 17 ng PGE2/mg protein,
P < 0.05). However, during 24 h
postincubation after PFF treatment, no further upregulation of
PGE2 release occurred in any of
the five OP patients tested (Fig. 3B;
non-OP control group:
PFF, 73 ± 41 ng
PGE2/mg protein; +PFF, 155 ± 59 ng PGE2/mg protein,
P < 0.05; OP group:
PFF, 32 ± 25 ng PGE2/mg protein; +PFF,
33 ± 22 ng PGE2/mg protein,
P = 0.5). The difference in
PGE2 release during 24 h post-PFF
between cells that had and those that had not been treated with PFF was significantly lower in OP than in non-OP control cells, in both an
absolute sense and as the ratio of +PFF to
PFF (
+PFF,
PFF: non-OP control, 81 ± 22 PGE2
ng/mg protein; OP, 6 ± 2 PGE2 ng/mg protein,
P < 0.005, Mann-Whitney
U test; ratio of +PFF to
PFF: non-OP control, 5.1 ± 1.5; OP, 1.5 ± 0.2, P < 0.02, Mann-Whitney U test).
|
The pattern of NO release was similar in the OP group and the non-OP
control group, i.e., a twofold increase after 1 h of PFF
treatment that was not continued during post-PFF incubation (Fig.
4, A and
B). Also, the pattern of TGF-
release was similar. In both the non-OP control group and the OP group,
TGF-
release was diminished by 1 h of PFF treatment (non-OP control:
PFF, 14 ± 7 ng TGF-
/ng protein; +PFF, 11 ± 7 ng
TGF-
/ng protein, P < 0.05; OP:
PFF, 14 ± 5 ng TGF-
/ng protein; +PFF, 9 ± 3 ng TGF-
/ng protein, P < 0.05) but not during the 24 h postincubation after PFF treatment (Fig.
5, A and
B).
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present study shows that cell cultures derived from human trabecular bone biopsies respond to fluid shear stress in a manner that is largely similar to the response of embryonic chicken or neonatal mouse calvarial bone cells (20, 22, 23). Primary cultures of untransformed cells derived from human bone explants have been shown to express an osteoblast-like phenotype (1, 2, 44). In the present study, all cultures tested responded to 1,25(OH)2D3 treatment with enhanced release of osteocalcin and enhanced ALP activity, both markers of the osteoblastic phenotype (2, 38). In addition, staining for vFF, a characteristic of endothelial cells, was negative.
One-hour treatment with PFF increased PGE2 release by the bone cell cultures, an effect that was also found in chicken osteocytes and in mouse bone cells (20, 22, 23). This response continued after stopping PFF application, leading to a twofold increase of PGE2 concentration in the culture medium after 24 h postincubation. Such a sustained upregulation of PGE2 release has also been observed in chicken and mouse bone cells (22, 23). In mouse bone cell cultures, Northern blot analysis showed that PFF treatment induced the expression of prostaglandin G/H synthase (PGHS)-2 [or cyclooxygenase (COX)-2] (17). PGHS, or COX, is a key enzyme regulating prostaglandin synthesis and is present as two isoforms, COX-1, the constitutive isoform, and COX-2, the inducible enzyme (24, 39, 50). Induction of COX-2 provides a cellular mechanism for continued upregulation of prostaglandin release, even when the cell stress itself is removed. Because expression of COX-2 but not COX-1 seems to be required for inducing an adaptive response to mechanical stress in bone in vivo (9), the in vitro data by Klein-Nulend et al. (17) suggest that continued upregulation of PGE2 release after a stress experience is part of the signal transduction pathway involved in mechanical adaptation of bone. In this respect, the reduced long-term response of the OP group is remarkable. In none of the five OP bone cell cultures tested did PFF treatment induce a continued upregulation of PGE2 release. This was surprising, considering that a continued response was always found in the non-OP controls, both in the age-matched control group and in the younger donors. We have recently found that 1 h of PFF treatment induces expression of COX-2 in (non-OP) human bone cells the same as in mouse bone cells (M. Joldersma, J. Klein-Nulend, and E. H. Burger, unpublished results). It will be of interest to study the prostaglandin response to stress by OP bone cells in more detail, including expression of COX-2. Impaired capacity of bone cells to respond to mechanical stress might be a cause of abnormal bone metabolism in OP patients (10, 29). It remains unclear whether the abnormal response of the OP cells was the cause of the OP status of the donor or whether it resulted from the OP status. All OP donors were treated with vitamin D, and we cannot exclude that this treatment was (part of) the underlying cause of the abnormal response to loading. On the other hand, the dose of vitamin D supplement was low (400 IU/day), and the control group did not show any evidence of vitamin D deficiency. In addition, both the OP and the non-OP control cells had been growing for 4-5 wk in standard tissue culture medium before they were subjected to PFF. It therefore seems unlikely that the abnormal PFF response of the OP cells was caused by extrinsic factors such as hormonal or nutritional abnormality of the donor. Rather, these in vitro results suggest an intrinsic impairment of the bone cells in their response to stress. Such an intrinsic abnormality could have a negative effect on bone adaptation and therefore on the mechanical quality of the skeleton and, as such, be a risk factor for osteoporosis.
Application of PFF also increased the release of NO, in line with earlier observations in mouse and chicken bone cell cultures (20). The kinetics of this response are consistent with activation of a constitutive NO synthase (NOS) enzyme. Of the two constitutively expressed isoforms, endothelial NOS (eNOS) was recently demonstrated in cultured osteoblasts as well as in trabecular lining cells in sections of rat bone, but neuronal NOS was not found (15). Therefore, it is likely that activation of eNOS was involved in the response to PFF. Interestingly, we have recently found, using semi-quantitative PCR, that mRNA levels for eNOS in human bone cell cultures increased approximately twofold after 1 h of PFF (18). eNOS, or NOS III, is located in the plasma membrane of endothelial cells and is involved in the adaptive response of the vasculature to changes in blood fluid flow (14, 28, 33, 40). The presence of this enzyme in bone cells and its activation by PFF suggest a similar role in the adaptive response of bone to mechanical stress via a similar mechanism of fluid shear stress-activated cells.
The release of TGF- by the bone cell cultures was inhibited by PFF
during but not after fluid flow treatment. In another study, using
osteoblastic cells from neonatal mouse calvariae, we also found a
reduction of TGF-
release after 1 h of PFF treatment (21). The
biological meaning of this observation is unsure because TGF-
may
have both anabolic and catabolic effects on bone (3, 6, 26). In
addition, the kinetics of the TGF-
response to stress seems to be
time variable because long-term (3 days) treatment with cyclic stress
has been reported to increase TGF-
production by bone cells (19,
29). This increase was reduced in bone cells from OP patients (29). The
short-term inhibitory effect of stress as studied in the present paper
was not altered in OP patients. Although it is clear that TGF-
release by bone cells may be modulated by mechanical stress, the
details of this effect and its relation with osteoporosis need further
study.
In sum, the present study shows that bone cells cultured from human
trabecular bone fragments respond to fluid shear stress with a
transient modulation of NO and TGF- release and a more sustained
upregulation of PGE2 release. The
sustained response was reduced in cells from OP patients, an
observation that warrants further study.
![]() |
ACKNOWLEDGEMENTS |
---|
We express gratitude to Drs. L. Smeele, J. C. Netelenbos, J. Baart, and J. van Boven for assistance in obtaining the bone biopsies. We gratefully acknowledge the staff of the Endocrine Laboratory (Dr. C. Popp-Snijders, Head) for performing the osteocalcin assays.
![]() |
FOOTNOTES |
---|
The research of J. Klein-Nulend was made possible by a fellowship of the Royal Netherlands Academy of Arts and Sciences, and that of J. G. H. Sterck was made possible by Vrije Universiteit-Universitair Stimulerings Fonds Grant 91/23.
This work was presented in part at the 1996 World Congress on Osteoporosis, Amsterdam, The Netherlands, May 1996.
Address for reprint requests: E. H. Burger, ACTA-Vrije Universiteit, Dept. of Oral Cell Biology, Van der Boechorststraat 7, 1081 BT Amsterdam, The Netherlands.
Received 25 September 1997; accepted in final form 26 February 1998.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Auf'mkolk, B.,
P. V. Hauschka,
and
E. R. Schwartz.
Characterization of human bone cells in culture.
Calcif. Tissue Int.
37:
228-235,
1985[Medline].
2.
Beresford, J. N.,
J. A. Gallagher,
M. Gowen,
M. K. B. McGuire,
J. Poser,
and
R. G. G. Russell.
Human bone cells in culture. A novel system for the investigation of bone cell metabolism.
Clin. Sci. (Colch.)
64:
38-39,
1983.
3.
Bonewald, L. F.,
and
G. R. Mundy.
Role of transforming growth factor- in bone remodeling.
Clin. Orthop.
250:
261-276,
1990[Medline].
4.
Brown, P. D.,
L. M. Wakefield,
A. D. Levinson,
and
M. B. Sporn.
Physiochemical activation of recombinant latent growth factor-'s 1, 2 and 3.
Growth Factors
3:
35-43,
1990[Medline].
5.
Cowin, S. C.,
L. Moss-Salentijn,
and
M. L. Moss.
Candidates for the mechanosensory system in bone.
J. Biomech. Eng.
113:
191-197,
1991[Medline].
6.
Dieudonné, S. C.,
P. Foo,
E. J. J. van Zoelen,
and
E. H. Burger.
Inhibiting and stimulating effects of TGF-1 on osteoclastic bone resorption in fetal mouse bone organ cultures.
J. Bone Miner. Res.
6:
479-487,
1991[Medline].
7.
Eriksen, E. F.,
S. F. Hodgson,
R. Eastell,
S. L. Cedel,
W. M. O'Fallon,
and
L. Riggs.
Cancellous bone remodeling in type I (postmenopausal) osteoporosis: quantitative assessment of rates of formation, resorption, and bone loss at tissue and cellular levels.
J. Bone Miner. Res.
5:
311-319,
1990[Medline].
8.
Evans, C. H.,
M. Stefanovic-Racic,
and
J. Lancaster.
Nitric oxide and its role in orthopaedic disease.
Clin. Orthop.
312:
275-294,
1995[Medline].
9.
Forwood, M. R.
Inducible cyclo-oxygenase (COX-2) mediates the induction of bone formation by mechanical loading in vivo.
J. Bone Miner. Res.
11:
1688-1693,
1996[Medline].
10.
Frost, H. M.
The role of changes in mechanical usage set points in the pathogenesis of osteoporosis.
J. Bone Miner. Res.
7:
253-261,
1992[Medline].
12.
Green, L. C.,
D. A. Wagner,
J. Glogowski,
P. L. Skipper,
J. S. Wishnok,
and
S. R. Tannenbaum.
Analysis of nitrate, nitrite, and [15N]nitrate in biological fluids.
Anal. Biochem.
126:
131-138,
1982[Medline].
13.
Gross, T. S.,
J. L. Edwards,
K. J. McLeod,
and
C. T. Rubin.
Strain gradients correlate with sites of periosteal bone formation.
J. Bone Miner. Res.
12:
982-988,
1997[Medline].
14.
Hecker, M.,
A. Mülsch,
E. Bassenge,
U. Förstermann,
and
R. Busse.
Subcellular localization and characterization of nitric oxide synthase(s) in endothelial cells: physiological implications.
Biochem. J.
299:
247-252,
1994[Medline].
15.
Helfrich, M. H.,
D. E. Evans,
P. S. Grabowski,
J. S. Pollock,
H. Ohshima,
and
S. H. Ralston.
Expression of nitric oxide synthase isoforms in bone and bone cell cultures.
J. Bone Miner. Res.
12:
1108-1115,
1997[Medline].
16.
Jee, W. S. S.,
and
X. J. Li.
Adaptation of cancellous bone to overloading in the adult rat: a single photon absorptiometry and histomorphometry study.
Anat. Rec.
227:
418-426,
1990[Medline].
17.
Klein-Nulend, J.,
E. H. Burger,
C. M. Semeins,
L. G. Raisz,
and
C. C. Pilbeam.
Pulsating fluid flow stimulates prostaglandin release and inducible prostaglandin G/H synthase mRNA in primary mouse bone cells.
J. Bone Miner. Res.
12:
45-51,
1997[Medline].
18.
Klein-Nulend, J.,
M. H. Helfrich,
J. G. H. Sterck,
C. M. Semeins,
and
E. H. Burger.
Human primary bone cells respond to fluid flow with rapid production of nitric oxide by endothelial nitric oxide synthase (Abstract).
J. Bone Miner. Res.
12:
S191,
1997.
19.
Klein-Nulend, J.,
J. Roelofsen,
J. G. H. Sterck,
C. M. Semeins,
and
E. H. Burger.
Mechanical loading stimulates the release of transforming growth factor- activity by cultured mouse calvariae and periosteal cells.
J. Cell. Physiol.
163:
115-119,
1995[Medline].
20.
Klein-Nulend, J.,
C. M. Semeins,
N. E. Ajubi,
P. J. Nijweide,
and
E. H. Burger.
Pulsating fluid flow increases nitric oxide (NO) synthesis by osteocytes but not periosteal fibroblastscorrelation with prostaglandin upregulation.
Biochem. Biophys. Res. Commun.
217:
640-648,
1995[Medline].
21.
Klein-Nulend, J.,
C. M. Semeins,
and
E. H. Burger.
Prostaglandin-mediated modulation of transforming growth factor- metabolism in primary mouse osteoblastic cells in vitro.
J. Cell. Physiol.
168:
1-7,
1996[Medline].
22.
Klein-Nulend, J.,
J. G. H. Sterck,
C. M. Semeins,
P. Lips,
and
E. H. Burger.
Aging and mechanosensitivity of human bone cells (Abstract).
J. Bone Miner. Res.
11:
S266,
1996.
23.
Klein-Nulend, J.,
A. van der Plas,
C. M. Semeins,
N. E. Ajubi,
J. A. Frangos,
P. J. Nijweide,
and
E. H. Burger.
Sensitivity of osteocytes to biomechanical stress in vitro.
FASEB J.
9:
441-445,
1995
24.
Kujubu, D. A.,
B. S. Fletcher,
B. C. Varnum,
R. W. Lim,
and
H. R. Herschman.
TIS 10, a phorbol ester tumor promoter-inducible mRNA from Swiss 3T3 cells, encodes a novel prostaglandin synthase/cyclooxygenase homologue.
J. Biol. Chem.
266:
12866-12872,
1991
25.
Lowry, O. H.
Micromethods for the assay of enzyme. II. Specific procedures. Alkaline phosphatase.
Methods Enzymol.
4:
371,
1955.
26.
Marcelli, C.,
A. J. Yates,
and
G. R. Mundy.
In vivo effects of human recombinant transforming growth factor- on bone turnover in normal mice.
J. Bone Miner. Res.
5:
1087-1096,
1990[Medline].
27.
Marie, P. J.,
A. Lomri,
A. Sabbagh,
and
M. Basle.
Culture and behavior of osteoblastic cells isolated from normale trabecular bone surfaces.
In Vitro Cell. Dev. Biol.
25:
373-380,
1989[Medline].
28.
Marsden, D. A.,
K. T. Schappert,
H. S. Chen,
M. Flowers,
C. L. Sundell,
J. N. Wilcox,
S. Lamas,
and
T. Michel.
Molecular cloning and characterization of human endothelial nitric oxide synthase.
FEBS Lett.
307:
287-293,
1992[Medline].
29.
Neidlinger-Wilke, C.,
I. Stall,
L. Claes,
R. Brand,
I. Hoellen,
S. Rhbenacker,
M. Arand,
and
L. Kinzl.
Human osteoblasts from younger normal and osteoporotic donors show differences in proliferation and TGF- release in response to cyclic strain.
J. Biomech.
28:
1411-1418,
1995[Medline].
30.
Nordin, B. E. C.,
A. G. Need,
B. E. Chatterton,
M. Horowitz,
and
H. A. Morris.
The relative contributions of age and years since menopause to postmenopausal bone loss.
J. Clin. Endocrinol. Metab.
70:
83-88,
1990[Abstract].
31.
Parfitt, A. M.
Bone remodeling: relationship to the amount and structure of bone, and the pathogenesis and prevention of fractures.
In: Osteoporosis: Etiology, Diagnosis, and Management, edited by B. L. Riggs,
and L. J. Melton. New York: Raven, 1988, p. 45-93.
32.
Piekarski, K.,
and
M. Munro.
Transport mechanism operating between blood supply and osteocytes in long bones.
Nature
269:
80-82,
1997.
33.
Pollock, J. S.,
U. Forstermann,
J. A. Mitchell,
T. D. Warner,
H. H. Schmidt,
M. Nakane,
and
F. Murad.
Purification and characterization of particulate endothelium-derived relaxing factor synthase from cultured and native bovine aortic endothelial cells.
Proc. Natl. Acad. Sci. USA
88:
10480-10484,
1991[Abstract].
34.
Raisz, L. G.
Osteoporosis.
J. Am. Geriatr. Soc.
30:
127-138,
1982[Medline].
35.
Raisz, L. G.,
and
T. J. Martin.
Prostaglandins in bone and mineral metabolism.
In: Bone and Mineral Research, Annual 2, edited by W. A. Peck. Amsterdam: Elsevier, 1983, p. 286-310.
36.
Reich, K. M.,
and
J. A. Frangos.
Protein kinase C mediates flow-induced prostaglandin E2 production in osteoblasts.
Calcif. Tissue Int.
52:
62-66,
1993[Medline].
37.
Riggs, B. L.,
and
L. J. Melton III.
Involutional osteoporosis.
N. Engl. J. Med.
314:
1676-1686,
1986[Medline].
38.
Robey, P. G.,
and
J. D. Termine.
Human bone cells in vitro.
Calcif. Tissue Int.
37:
453-460,
1985[Medline].
38a.
Robey, P. G.,
M. F. Young,
K. C. Flanders,
N. S. Roche,
P. Kondaiah,
A. H. Reddi,
J. D. Termine,
M. B. Sporn,
and
A. B. Roberts.
Osteoblasts synthesize and respond to transforming growth factor type (TGF-
) in vitro.
J. Cell Biol.
105:
115-119,
1987.
39.
Rosen, G. D.,
T. M. Birkenmeier,
A. Raz,
and
M. J. Holtzman.
Identification of a cyclooxygenase-related gene and its potential role in prostaglandin formation.
Biochem. Biophys. Res. Commun.
164:
1358-1365,
1989[Medline].
40.
Schmidt, H. H. H. W.,
J. S. Pollock,
M. Nakane,
L. D. Gorski,
U. Förstermann,
and
F. Murad.
Purification of a soluble isoform of guanylyl cyclase-activating-factor synthase.
Proc. Natl. Acad. Sci. USA
88:
365-369,
1991[Abstract].
41.
Seyedin, S. M.,
A. Y. Thompson,
D. M. Rosen,
and
K. A. Piez.
Purification and characterization of two cartilage inducing factors from bovine mineralized bone.
Proc. Natl. Acad. Sci. USA
82:
2267-2271,
1985[Abstract].
42.
Slemenda, C.,
S. L. Hui,
C. Longcope,
and
C. C. Johnston.
Sex steroids and bone mass.
J. Clin. Invest.
80:
1261-1269,
1987[Medline].
43.
Smith, P. K.,
R. I. Krohn,
G. T. Hermansom,
A. K. Mallia,
F. H. Gartner,
M. D. Provenzano,
E. K. Fujimoto,
N. M. Goeke,
B. J. Olson,
and
D. C. Klenk.
Measurement of protein using bicinchoninic acid.
Anal. Biochem.
150:
76-85,
1985[Medline].
44.
Sterck, J. G. H.,
J. Klein-Nulend,
E. H. Burger,
and
P. Lips.
1,25-Dihydroxyvitamin D3-mediated transforming growth factor- release is impaired in cultured osteoclasts from patients with multiple hormone deficiencies.
J. Bone Miner. Res.
11:
367-376,
1996[Medline].
45.
Turner, C. H.,
M. R. Forwood,
and
M. W. Otter.
Mechanotransduction in bone: do bone cells act as sensors of fluid flow?
FASEB J.
8:
875-878,
1994
46.
Van der Wiel, H. E.,
P. Lips,
J. Nauta,
J. C. Netelenbos,
and
G. J. Hazenberg.
Biochemical parameters of bone turnover during ten days bed rest and subsequent mobilization.
Bone Miner.
13:
123-129,
1991[Medline].
47.
Villareal, D. T.,
and
J. E. Morley.
Trophic factors in aging: should older people receive hormonal replacement therapy?
Drugs Aging
4:
492-509,
1994[Medline].
48.
Weinbaum, S.,
S. C. Cowin,
and
Y. Zeng.
A model for the excitation of osteocytes by mechanical loading-induced bone fluid shear stress.
J. Biomech.
27:
339-360,
1991.
49.
Weinreb, M.,
G. A. Rodan,
and
D. D. Thompson.
Osteopenia in the immobilized rat hind limb is associated with increased bone resorption and decreased bone formation.
Bone
10:
187-194,
1989[Medline].
50.
Xie, W.,
J. G. Chipman,
D. L. Robertson,
R. L. Erikson,
and
D. L. Simmons.
Expression of a mitogen-responsive gene encoding prostaglandin synthase is regulated by mRNA splicing.
Proc. Natl. Acad. Sci. USA
88:
2692-2696,
1991[Abstract].