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Institute of Molecular Medicine and Genetics, Department of Medicine, Medical College of Georgia (R.J.B., Q.Z., P.P., L.M., L.Z., R.C., H.R., F.Q., K.H.D.), and the Augusta Veterans Administration Medical Center (A.L.M., C.M.I.), Augusta, Georgia 30912
Address all correspondence and requests for reprints to: Carlos M. Isales, M.D., Medical College of Georgia, Institute of Molecular Medicine and Genetics, 1120 15th Street, Augusta, Georgia 30912. E-mail: cisales{at}mail.mcg.edu
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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A number of hormones whose levels depend on nutritional status are known to impact bone metabolism. For example, insulin, released from the ß-cells of the pancreatic islets in response to meal ingestion engenders bone matrix deposition and mineralization (1, 2). Another hormone secreted by ß-cells, amylin, has also been shown to impact calcium homeostasis by inhibiting bone resorption, thereby reducing serum calcium and increasing bone mass (3, 4, 5). The possible role of gut-derived peptide hormones in bone homeostasis, however, has not been addressed.
Glucose-dependent insulinotropic peptide (GIP) is one such hormone, initially identified as a duodenal hormone with potential to inhibit gastric acid secretion (6). Subsequently, the 42-amino acid active form of GIP was shown to inhibit amylase release from acinar cells of the pancreas and to modulate insulin secretion from ß-cells in response to elevations in ambient glucose (7). Based on its insulinotropic action, GIP is classified as one of the two main incretin hormones, the other being glucagon-like polypeptide-1 (GLP-1). GIP not only potentiates insulin release, but also stimulates amylin release from the ß-cells of the pancreas, and in this way may potentially modulate bone turnover (5). In addition to the well characterized incretin effect in the pancreas (7), GIP has been shown to activate adipocytes (8), to modulate hepatic blood flow (9), and to inhibit acid secretion in the stomach (10). The receptor for GIP falls into the subclass of seven-transmembrane domain-spanning G protein-coupled receptors that includes receptors for PTH, calcitonin, and several other peptide hormones (11). As a receptor in this class, the GIP receptor (GIPR) is able to activate both cAMP-activated pathways as well as the phosphoinositide/Ca2+-dependent signaling systems. The GIPR localizes not only to the exocrine pancreas, but also to a wide range of tissues and organs, including the distal small bowel, pituitary gland, adrenal cortex, heart, brain, adipose tissue, and endothelial cells in several vascular beds (11).
In the present study we show that GIPRs are present in bone cells, including osteoblasts and osteocytes in bone proper as well as in osteoblast-like osteosarcoma cell lines. We show that these cell lines respond to GIP at physiological levels with metabolic responses exhibited by differentiated bone cells, namely enhanced collagen synthesis and alkaline phosphatase (ALP) activity. Finally, we propose that GIP could serve to coordinate nutrient intake in the intestine with nutrient disposal in a variety of peripheral tissues including bone.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cell culture
The cell lines studied in this report included SaOS2, MG63, ROS
17/2.8, HeLa, and NIH-3T3 fibroblasts. Normal primary human osteoblasts
were also used (Clonetics, San Diego, CA). Cells were grown to
confluence in MEM, RPMI, or DMEM as appropriate (BioWhittaker, Inc., Walkersville, MD), supplemented with 10% FCS (vol/vol;
HyClone Laboratories, Inc., Logan, UT), penicillin (100
U/ml), streptomycin (100 mg/ml), and amphotericin (3
mg/ml), and were used 3–7 days postconfluence. For studies on collagen
type I expression, SaOS2 cells were grown in a glutamine-free medium,
because we found that glutamine increased constitutive collagen
expression levels.
Western analyses
A polyclonal antibody was generated in rabbits to a synthetic
oligopeptide, SKGQTAGELYQRWERYRREC, corresponding to an extracellular
region of the human GIPR protein sequence. The oligopeptide was
conjugated to keyhole limpet hemocyanin using an Imject
maleimide-activated immunogen conjugation kit (Pierce Chemical Co., Rockford, IL), and was inoculated into rabbits (Animal
Pharm Services, Inc., Healdsburg, CA). The serum was affinity purified
on oligopeptide-BSA cross-linked to cyanogen bromide-Sepharose, and the
antibody was assessed by Western blot analysis to bacterially expressed
GIPR protein. The specificity of the primary antibody was tested by
competitively displacing the GIPR primary antibody binding in SaOS2
cells with primary antibody in the presence of an excess (140 µg) of
the GIPR peptide antigen used to generate the primary antibody.
Confluent bone cells (500,000/pair of lanes) were scraped into ice-cold PBS (pH 7.4) and disrupted by sonication for 60 sec in ice-cold homogenization buffer [60 mM Tris buffer (pH 7.4), 0.25 M sucrose, 10 mM EGTA, 2 mM EDTA, 10 mM ß-mercaptoethanol, and protease inhibitors]. Proteins were placed in sample buffer [0.5 M Tris (pH 6.8), 4% SDS, 20% glycerol, and 0.1% bromophenol blue] and boiled.
The denatured proteins were separated by SDS-PAGE and incubated with affinity-purified GIPR antibody at a 1:250 dilution. Immunoreactive bands were visualized with a horseradish peroxidase-conjugated secondary goat antirabbit serum and developed with enhanced chemiluminescence (Pierce Chemical Co.).
Indirect immunofluorescence
For the animal studies, 3-month-old Sprague Dawley rats
(Charles River Laboratories, Inc., Wilmington, MA) were
killed, and tissue was removed. This protocol was approved by the
Medical College of Georgia animal care committee. Rats were
anesthetized and perfused transcardially with 4% paraformaldehyde and
0.2% glutaraldehyde. Tibiae and vertebrae were harvested and fixed in
4% paraformaldehyde overnight at 4 C. The bones were decalcified in
10% EDTA for 4–6 days, dehydrated in ethanol and xylene, and embedded
in paraffin. Seven-micron thick sections were cut, deparaffined, and
rehydrated.
SaOS2 and MG63 cells were plated on glass coverslips, grown in DMEM supplemented with 10% FCS for 48 h, and fixed in ice-cold 4% paraformaldehyde-PBS (137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2PO4, 1.15 mM KH2PO4, and 1 mM MgSO4, pH 7.4) for 30 min. Cells were then rinsed three times for 5 min each time in PBS and transferred into PBS-NH4Cl (50 mM) for 15 min. After rinsing, the fields were covered with PBS-BSA (100 mg/10 ml) for 15 min (blocking buffer). The cells were then incubated with the GIP antibody in 500 µl PBS-BSA for 45 min. The cells were rinsed and then covered with secondary antibody, Cy3 goat antimouse IgG (5 µl; Molecular Probes, Inc.), and visualized by epifluorescence (Axiophot microscope, Carl Zeiss, Inc., Thornwood, NY).
The immunohistochemical localization analyses were carried out using the Vectastain ABC peroxidase system with 3,3'-diaminobenzidine as the peroxidase substrate (Vector Laboratories, Inc., Burlingame, CA). Developed slides were dehydrated in ethanol, cleared in xylene, and counterstained in hematoxylin (Vector Laboratories, Inc.).
Receptor binding studies
Receptor binding studies were performed as previously
described (12). Briefly SaOS2, MG63, or NIH-3T3 fibroblasts were grown
in six-well plates and incubated with increasing concentrations of
[125I]GIP (Amersham Pharmacia Biotech) in the presence or absence of an excess of unlabeled
GIP (1 µM) for 2 h at room temperature. Cells were
then washed three times with 1 ml cold PBS (plus 0.05% BSA) and
solubilized with 0.3 M NaOH. The extract was counted in a
-counter, and background counts were subtracted. Experiments were
performed in triplicate.
Intracellular calcium measurements with fura-2
Intracellular calcium measurements were made as previously
described (13). Briefly, SaOS2 cells were grown on glass coverslips.
Cells were loaded with the calcium-sensitive dye fura-2/AM in KRB.
After approximately 30 min at room temperature to allow esterase
cleavage of fura-2/AM to fura-2, the coverslips were placed in a
cuvette in a dual wavelength spectrophotometer (Photon Technologies
International, South Brunswick, NJ). Fluorescence was measured using
excitation wavelengths of 340 and 380 nm and an emission wavelength of
510 nm. After 200 sec to allow baseline stabilization, cells were
stimulated by appropriate concentrations of GIP as indicated in Fig. 4A
. Measurements were subsequently collected for an additional 800 sec,
and peak calcium levels were determined by convention as follows:
minimum emission (Rmin) was measured upon
addition of EGTA (4 mM) buffered with Tris (pH 8.0)
followed by addition of the calcium ionophore, ionomycin (50
µM). Maximum emission (Rmax) was
measured upon addition of 12 mM calcium chloride. The free
intracellular calcium concentration was then calculated using the
equation [Ca2+] = Kd x
b(R - Rmin)/(Rmax -
r) (14). Autofluorescence was measured in unloaded cells, and this
value was subtracted from all measurements.
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cAMP measurements
cAMP measurements were performed as previously described (15).
Briefly, SaOS2 cells were grown to confluence in
60-mm2 dishes and placed in KRB for 24 h
before use. To facilitate the measurement of cAMP production, 1
mM IBMX was added for 10 min before agonist addition,
followed by incubation with vehicle (control) or with GIP at
concentrations ranging from 0.1 nM to 1 µM
for 10 min. Incubations were stopped by addition of 5% trichloroacetic
acid and left on ice for 15 min, and the cell extract was collected.
The extracts were neutralized by the addition of a 1:1 solution of
ice-cold freon/tri-N-octylamine (4:1, vol/vol). Each sample
was vortexed for at least 30 sec to ensure adequate mixing. The mixture
was then centrifuged at 2500 rpm for 20 min at 4 C. The top aqueous
phase containing cAMP was collected. The pH of the upper phase was
checked to ensure adequate neutralization. The samples were stored at
-70 C until analysis. cAMP was measured using a commercially available
RIA (Biomedical Technologies). All incubations were
performed in triplicate, and each experiment was repeated three times
using different cell preparations.
Preparation of RNA and Northern blot analysis
Total RNA was extracted from cells using Trizol (Life Technologies, Inc.). RNA was stored at -70 C until use. RNA (20
µg/lane) was electrophoresed on a 1.2% agarose-formaldehyde gel and
transferred to a nylon filter. The blots were hybridized overnight at
65 C with a 32P-labeled probe
(106 cpm/ml), labeled by the random priming
method, and washed at maximum stringency. Hybridization was carried out
in a solution of 7% SDS, 1% BSA, 1 mM EDTA, and 250
mM Na2HPO4. The
hybridized filters were washed with four 5-min washes of 2 x
SSC/0.1% SDS at room temperature and twice in 0.1 x SSC/0.1%
SDS for 30 min at 65 C. The blots were then exposed to
Kodak XAR 5 film (Eastman Kodak Co.,
Rochester, NY). The probes used were glyceraldehyde-3-phosphate
dehydrogenase (American Type Culture Collection, Manassas,
VA; clone 57090), collagen I
(i.m.a.g.e. clone 308919), and a human
GIPR fragment derived by PCR corresponding to transmembrane domains
2–7.
ALP activity
ALP activity was measured using a commercially available assay
kit (ALP EC 3.1.1.1 colorimetric test, Sigma). This kit
measures the conversion of p-nitrophenyl phosphate to
p-nitrophenol and inorganic phosphate. The change in
absorbance at 405 nm is directly proportional to ALP activity. MG63
cells were grown in six-well plates and incubated with the indicated
agonist for the indicated times (medium was changed daily with
readdition of fresh agonist), and samples were collected. The kit
reagents were added to the sample cuvette in the spectrophotometer at
30 C, and absorbances at 405 nm were obtained at 1, 2, and 3 min. ALP
activity (units per liter) was determined using the change in
absorbance with time and a millimolar absorptivity of
p-nitrophenol at 405 nm of 18.45.
Statistics
Results are expressed as the mean ± SEM. Data
were analyzed using either ANOVA or unpaired t tests, where
appropriate, with a commercial statistical package (Instat,
GraphPad Software, Inc., San Diego, CA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). Protein extracts from several
tissues from a normal Sprague Dawley rat were prepared and probed with
the GIPR antibody by Western blot (Fig. 1B). The same 50-kDa
immunoreactive band was observed in normal rat bone. In addition, the
pancreas, brain, and heart, tissues previously reported to contain the
GIPR (11), were positive; whereas the spleen, which has been reported
not to contain the GIPR (11), did not contain any immunoreactive band,
thus suggesting antibody specificity.
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). In normal rat bone both osteocytes
and osteoblasts demonstrated intense fluorescence (Fig. 2A). This
staining pattern was comparable to that seen by in situ
hybridization for GIPR mRNA (data not shown). In contrast, rat bone did
not demonstrate any cell labeling when incubated only with the
secondary antibody (Fig. 2B). SaOS2 cells also demonstrated intense
nonnuclear fluorescence (Fig. 2C), which could be blocked by the
presence of an excess of the antigen used to generate the antibody
(Fig. 2D). This result indicates the specific binding of the antibody
to the GIPR. MG63 cells also demonstrated intense fluorescence (Fig. 2E) with a pattern similar to that of SaOS2 cells. As an additional
control, MG63 cells were labeled only with the secondary antibody (Fig. 2F), demonstrating the specificity of the primary antibody. Although
MG63 and SaOS2 cells are transformed cells derived from human
osteosarcomas, we sought to determine whether, as expected from the
observations presented in Fig. 2A, normal human osteoblasts would also
stain positively with our GIPR antiserum. As shown in Fig. 2G, primary
cultures of normal human osteoblasts show positive signal with
anti-GIP-R antiserum, albeit with lower intensity than the transformed
cells. A control with secondary antibody alone (Fig. 2H) shows that the
signal is specific.
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). The osteoblast-like cell
lines SaOS2 and MG63 displayed high affinity binding sites, whereas the
3T3 fibroblasts showed minimal background binding effects. The
biological efficacy of the radiolabeled GIP was verified by monitoring
intracellular Ca2+ mobilization. Radiolabeled GIP
at 1 nM elicited an elevation in intracellular
Ca2+ in SaOS2 cells comparable to that observed
with unlabeled agonist (data not shown).
This is consistent with the idea that the binding sites identified in
Fig. 3 are physiologically relevant. Both MG63 and SaOS2 cells had
similar Kd values of approximately 0.3
nM, although the binding capacity in MG63 cells was greater
than that in SaOS2 cells.
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, GIP indeed increased both the cytosolic calcium concentration and the
cellular cAMP content in SaOS2 cells. The calcium response shows a
dose-dependent increase in Ca2+, with a
significant increase occurring at 0.1 nM GIP (Fig. 4A). GIP
at concentrations of 1 nM and above was able to
significantly stimulate elevations in cellular cAMP content (Fig. 4B).
These elevations reached 710% above the control level at a GIP
concentration of 1 µM.
GIP stimulates collagen (I) gene expression and activates ALP
activity in osteoblast-like cells lines
While the GIPR coupled to signaling events in bone cells, an issue
still unanswered was the possible role of GIP in normal bone cell
biology. To address this issue, we examined the effect of GIP on two
anabolic indexes of bone formation: new matrix synthesis and ALP
activity in osteoblast-like cells.
We initially determined whether GIP could stimulate collagen type I
expression in SaOS2 cells (Fig. 5). SaOS2
cells were stimulated with increasing concentrations of GIP, and
collagen type I expression was assessed by Northern blot and
quantitated by densitometry. GIP, at a concentration of 1
nM or above, stimulated the expression of type 1 collagen,
a marker for bone formation (Fig. 5, A and B). This GIP effect appeared
to show a threshold effect, as no further increases in collagen type I
expression were seen at the higher GIP concentrations. Since collagen
is the primary constituent of bone matrix, the ability to effect
collagen synthesis is consistent with an anabolic effect of GIP on
bone. To examine this issue further, we monitored the time course of
the GIP effect on collagen synthesis using the dose (1 nM)
determined to be maximally effective in the previous experiment. The
GIP effect on collagen mRNA could be observed after 6 h of
stimulation, with no further increases observed at the later time
points (Fig. 5C).
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). As shown in
Fig. 6, GIP at a concentration of 0.1
nM significantly increased ALP activity as early as 2 days
after stimulation and continued to increase ALP activity after 6 days
of GIP exposure. Higher doses of GIP (1–10 nM) did not
increase ALP activity any further than 0.1 nM GIP. The GIP
effect on ALP was larger than that observed with 1,25-dihydroxyvitamin
D3 (10 ng/ml) and transforming growth factor-ß (10
ng/ml), which were used as a positive control. Hence, GIP is a potent
inducer of ALP activity.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The functionality of the GIPRs in osteoblast-derived cells was also demonstrated by their ability to couple to signal transduction pathways. Like other related seven-transmembrane receptors, the GIPR in osteoblast-like cells appears to couple to both cAMP and phosphoinositide signaling pathways. In fact, the phosphoinositide response demonstrated greater potency, in that a significant effect was observed at 0.1 nM GIP vs. 1 nM for changes in cAMP content. The large increases in intracellular calcium concentration seen with the lower doses of GIP in osteoblast-like cells were unexpected based on previous studies with pancreatic ß-cell lines (16). Although GIP has been reported to increase extracellular calcium influx and to induce calcium mobilization from intracellular stores in pancreatic islets and cell lines, elevations in cAMP content have been considered the main intermediary in the incretin effect of GIP on glucose-induced insulin secretion (16, 20). One caveat, however, is that the cAMP measurements in the present study were of total cellular content; it is possible that localized changes in cAMP may occur to trigger cAMP-dependent protein kinase activation at lower concentrations of GIP.
The GIPRs were also functional, in that ligand binding elicited
cellular responses. Thus, treatment with GIP resulted in increased
collagen type I mRNA expression and ALP activity. One area of concern
is whether the concentrations of GIP used for the present study are
physiologically relevant. The reported basal GIP concentration varies
between 0.06–0.1 nM. After a meal, GIP levels increase
rapidly to reach levels of between 0.2–0.5 nM (21, 22, 23, 24, 25, 26).
GIP-induced changes in intracellular calcium and effects on ALP
activity could be observed with GIP concentrations as low as 0.1
nM, well within the physiological range. However, changes
in collagen type I expression required higher concentrations of GIP
(
1 nM), slightly above the normal physiological range.
This discrepancy may be related to limitations in the sensitivity of
this assay system itself and/or to the use of an established cell
line.
In view of the present data demonstrating functional GIPRs on bone-derived cells, the question then becomes: does GIP play a role in normal bone physiology? It is known that changes in the nutritional state of the organism influence skeletal homeostasis. A reduction in caloric intake rapidly inhibits linear growth (27). If the reduced caloric intake is accompanied by a reduced calcium intake, a shift in the balance between bone formation and resorption occurs, such that bone mass decreases over time (28). Nutrient-induced elevations of both insulin and amylin have been implicated in normal bone metabolism (3, 29, 30), and the ability of GIP to enhance the secretion of both hormones may contribute to an anabolic action of GIP on bone. However, based on our demonstration of GIPRs in bone cells and direct effects of GIP on bone-derived cells, we propose that GIP may serve directly to coordinate nutrient utilization by bone in addition to its incretin effects on insulin secretion. Thus, we speculate that the role of GIP in bone is analogous to that of insulin in other tissues in the body; when food is consumed, the rise in nutrient concentrations in the blood/gut stimulates insulin/GIP release, which then permits nutrient utilization by various insulin/GIP-sensitive tissues. Specifically, GIP may signal bone cells that nutrients are available for matrix deposition, thus allowing new bone formation.
In summary, GIPRs are present in bone and bone-derived cells, and stimulation of these cells with GIP results in increases in intracellular calcium levels, cellular cAMP content, type 1 collagen expression, and ALP activity. The physiological significance of these findings remains to be clarified, but we propose that GIP may be involved in an entero-osseous axis, in which GIP coordinates nutrient utilization for bone formation through direct effects of the hormone on osteoblasts.
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Acknowledgments |
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Footnotes |
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Received September 17, 1999.
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| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
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) demonstrated
specific high affinity binding, whereas NIH-3T3 (
) cells
demonstrated no specific binding. SaOS2 (
) demonstrated GIP binding
with an affinity similar to that of MG63, but total binding was lower.
Shown in the insets are the Scatchard plots for binding
in MG63 (A) and SaOS2 (B).
