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Laboratory of Developmental Biology and Repair, Department of Surgery, New York University School of Medicine, New York, New York 10016; Department of Surgery, University of Connecticut (P.B.S.), Farmington, Connecticut 06032; Department of Surgery, New York University (B.J.M., D.S.S., J.A.S., J.A.G., G.S.C., G.K.G., M.T.L.), New York, New York 10016; and Department of Cardiology, Kyushu University School of Medicine (H.U.), Fukuoka 812, Japan
Address all correspondence and requests for reprints to: Michael T. Longaker, M.D., Laboratory of Developmental Biology and Repair, Room H-169, New York University Medical Center, 550 First Avenue, New York, New York 10016. E-mail: michael.longaker{at}med.nyu.edu
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Introduction |
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On a molecular level, cytokines that modulate bone development have been identified and include members of the fibroblast growth factor (FGF) and transforming growth factor-ß (TGFß) families (6, 7, 8, 9). FGF-2, a highly conserved heparin-binding growth factor, has been implicated in the control of skeletal and neural differentiation, angiogenesis, wound healing, and tissue repair (10). FGF-2 is involved in early limb development (11, 12, 13), and in the developing skeleton, FGF-2 and FGF receptor 1 colocalize to proliferating chondrocytes of the epiphyseal growth plate (14). In vitro, FGF-2, which is produced by osteoblastic cells and is stored in the extracellular matrix (15, 16), stimulates osteoblastic proliferation (17) and TGFß1 production (18).
Given the involvement of FGF-2 in differentiation, growth, and repair and the dependence of these related processes on an adequate blood supply, it is not surprising that FGF-2 is a potent modulator of angiogenesis. FGF-2 stimulates endothelial cell migration, proliferation, and angiogenesis in vitro (10) and has been implicated in embryonic vascular development (14, 19). Additionally, FGF-2 induces vascular endothelial growth factor (VEGF) in endothelial cells (20). Finally, in an in vitro angiogenic assay, VEGF-induced angiogenesis is dependent on FGF-2 (21).
VEGF, a dimeric heparin-binding glycoprotein, plays a central role in the development and modulation of angiogenesis. VEGF is expressed in highly vascular tissues and is an endothelial cell-specific mitogen (22). VEGF receptor knockout mice lack adequate blood vessel formation (23), whereas loss of a single VEGF allele is lethal in the mouse embryo (24). With respect to bone, VEGF is expressed in both the normal rat tibia (25) and mandible (26) and by unstimulated osteoblastic cells (27). Additionally, VEGF expression in osteoblastic cells is increased by several cytokines and growth factors, including TGFß1, PGE1, PGE2, insulin-like growth factor, platelet-derived growth factor, and 1,25-dihydroxyvitamin D3 (25, 26, 28, 29).
Although FGF-2 may exert its angiogenic effects both directly and through VEGF, an additional potential angiogenic mechanism may involve alterations in TGFß biological activity, as FGF-2 stimulates both increased TGFß1 production as well as latent TGFß activation. The three defined TGFß isoforms in mammals, TGFß1, TGFß2, and TGFß3, are ubiquitous cytokines with pleiotropic effects and have been implicated in osteoblastic proliferation and differentiation. TGFß1, the largest source of which is bone (30), is expressed in high levels during bone growth and development (27, 31), localized to cells within the developing skeleton (32), and increased in fracture healing (33). In vitro, TGFß1 stimulates osteoblastic migration, modulates osteoblastic proliferation, and increases VEGF expression in osteoblastic cells (26, 34, 35). Despite these findings, the effects of TGFß2 and TGFß3 on VEGF production remain unknown. Given the variable affinity of TGFß receptors for TGFß isoforms, in addition to the finding that individual TGFß subtypes have been implicated in the regulation of palatal and craniofacial development as well as scar formation, analysis of isoform-specific regulation of VEGF expression is warranted. (36, 37, 38, 39) Additionally, the interactions between the TGFß isoforms and FGF-2 in the control of VEGF remain undefined.
Given the importance of FGF-2 and VEGF in the related processes of angiogenesis and bone development, we proposed that FGF-2 may regulate VEGF expression in osteoblastic cells. We found that FGF-2 increased VEGF mRNA in both nonimmortalized and clonal osteoblastic cells. These data suggest a transcriptional mechanism of FGF-2-regulated VEGF expression. Additionally, FGF-2 increased VEGF protein production by osteoblastic cells. Both TGFß2 and TGFß3 increased VEGF mRNA and protein in osteoblastic cells. To dissect the effect of FGF-2 on VEGF from possible FGF-2-induced/activated TGFß (all isoforms of which also increased VEGF), we employed a recombinant adenovirus to mediate the transfer of a dominant negative truncated TGFß receptor II gene, thereby disrupting TGFß signal transduction. Whereas osteoblastic cells transfected with the dominant negative truncated TGFß receptor II adenovirus demonstrated significantly decreased induction of VEGF mRNA by exogenous TGFß1, FGF-2 induction of VEGF mRNA remained similar to control cells.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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MEM,
0.05% trypsin-EDTA, PBS, FBS, and cell culture reagents were purchased
from Life Technologies, Inc. (Gaithersburg, MD).
Recombinant human FGF-2, TGFß2, and TGFß3 were obtained from
Life Technologies, Inc.. Actinomycin D and cycloheximide
were purchased from Sigma (St. Louis, MO). Collagenase A
and dispase II were obtained from Roche Molecular Biochemicals (Mannheim, Germany).
Animals
Pregnant Sprague Dawley rats were purchased from Taconic Farms, Inc. (Germantown, NY) and housed in separate cages.
Animals were kept under a constant 12-h light, 12-h dark schedule and
fed Purina rodent chow (Ralston Purina Co., St. Louis, MO)
ad libitum. All procedures were approved by the
institutional care and use committee at New York University Medical
Center. On gestational day 21, the pregnant mothers were killed with
carbon dioxide, and the pups were harvested.
Cell culture
Fetal rat calvarial (FRC) cells were cultured from FRC explants
according to a modification of the methods described by Centrella
et al. (40). Frontal and parietal bones from gestational
21-day-old Sprague Dawley fetal rats were sterilely stripped of all
surrounding soft tissue. Calvaria were washed with sterile PBS
containing antibiotic/antimycotic and underwent serial digestions
(0.1% collagenase A/0.2% dispase II) in a shaking incubator at 180
rpm for 10 min at 37 C. Fractions 2–5 were pooled, centrifuged, and
resuspended in medium (DMEM supplemented with 10% FBS, 100 µg/ml
penicillin G, 50 µg/ml streptomycin, and 0.25 µg/ml
amphotericin). Cells (1.5 x
106 cells/flask) were plated in
75-cm2 flasks and reached confluence (2.5 x
106 cells/flask) in 3 days, after which they were
trypsinzed with 0.05% trypsin and replated at a subconfluence
(2.0 x 106 cells/flask). The next day,
confluent passage 2 cultures were obtained and used for all
experiments. Medium was changed every 2 days and after replating.
Verification of osteoblastic lineage was performed by mineralized bone
nodule formation assay, alkaline phosphatase staining, and Northern
blot analysis for osteocalcin (data not shown).
MC3T3-E1 cells, a mouse clonal osteoblastic cell line (gift from Dr. A. Gosain, Medical College of Wisconsin, Milwaukee, WI), were grown in DMEM supplemented with 10% FBS, 100 µg/ml penicillin G, 50 µg/ml streptomycin, and 0.25 µg/ml amphotericin. Medium was changed every 2–3 days. Confluent MC3T3-E1 cell cultures (2.5 x 106 cells/T75 flask) were trypsinized with 0.05% trypsin and replated in a 1:2 ratio. All cultures were maintained in a humidified atmosphere consisting of 95% air-5% CO2 at 37 C.
Probe preparation
Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was a 1-kb
probe from CLONTECH Laboratories, Inc. (Palo Alto, CA). A
410-bp probe against mouse VEGF was PCR generated from whole mouse
embryo cDNA using the following PCR primer sequences:
5'-CGAGACCCTGGTGGACATCT-3' and 5'-CACCGCCTCGGCTTGTCAC-3'. Resultant
bands were cloned into PCR.1 plasmids (Invitrogen,
Carlsbad, CA) and sequenced to confirm sequence identity (GenBank
accession no. NM 009505). The resultant probe was extensively tested
using known controls and was found to be capable of recognizing all
murine VEGF isoforms (not shown). Probe was generated after
EcoRI digestion and gel purification. One hundred nanograms
of each probe were labeled with
[-32P]deoxy-CTP using random oligonucleotide
primers and Klenow fragment (Ready To Go labeling beads,
Pharmacia Biotech, Cambridge, UK). Unincorporated
nucleotides were removed using Sephadex G-50 DNA grade nick columns
(Pharmacia Biotech). All probes had a specific activity of
more than 105 cpm/ml hybridization solution.
RNA extraction and Northern analysis
Confluent FRC cells (passage 2) and MC3T3-E1 cells in T75 flasks
were stimulated with 12.5 ng/ml FGF-2 in antibiotic-containing
serum-free medium for 0, 3, 6, or 24 h. Confluent MC3T3-E1 cells
in T75 flasks were stimulated with 2.5 ng/ml TGFß2 or 2.5 ng/ml
TGFß3 in antibiotic-containing serum-free medium for 0, 3, 6, or
24 h. In experiments designed to investigate the effect of FGF-2
on VEGF mRNA stability, transcription was interrupted with actinomycin
D (5 µg/ml) after 5 h of stimulation with FGF-2 (12.5 ng/ml) in
serum-free medium. To investigate disruption of protein synthesis,
confluent MC3T3-E1 cells in T75 flasks were treated with cyclohexamide
(10 µg/ml) with or without FGF-2 (12.5 ng/ml) In addition, to
investigate disruption of gene transcription, confluent MC3T3-E1 cells
in T75 flasks underwent 1 h of pretreatment, followed by 6 h
of exposure to actinomycin D (5 µg/ml) with or without FGF-2 (12.5
ng/ml) (28).
Experiments were designed to disrupt TGFß signal transduction. A dominant negative truncated TGFß receptor II adenovirus and a ß-galactosidase adenovirus have been previously described and characterized in endothelial cells (41) and in MC3T3-E1 clonal osteoblastic cells (42). Transgene expression is regulated via the chicken actin promoter, and at a multiplicity of infection (moi) of 100 (100 plaque-forming units/cell), cells are efficiently transfected and strongly overexpress the truncated TGFß receptor II on their cell surface, which acts in a dominant negative fashion by competing with endogenously expressed receptors. After binding TGFß (all isoforms), the truncated receptor disrupts TGFß biological activity because it fails to phosphorylate TGFß receptor I, thereby interrupting intracellular signaling. An adenovirus containing the Escherichia coli ß-galactosidase gene expressed using the same promoter served as a control for nonspecific viral effects. Confluent MC3T3-E1 cells in T75 flasks were infected with vehicle (PBS with 10% glycerol), the dominant negative truncated TGFß receptor II adenovirus, or the ß-galactosidase adenoviral control (moi = 100). After 60 h of incubation, cells were stimulated with 12.5 ng/ml FGF-2 in antibiotic-containing serum-free medium for 0, 3, 6, or 24 h, after which total cellular RNA was harvested, and VEGF mRNA levels were analyzed using Northern blot analysis. To verify TGFß blockade, identically treated MC3T3-E1 cells were stimulated with 2.5 ng/ml TGFß1 in antibiotic-containing serum-free medium for 0, 3, 6, or 24 h, after which total cellular RNA was harvested, and VEGF mRNA levels were analyzed using Northern blot analysis.
Northern blot analysis
Total cellular RNA was extracted using Trizol solution
(Life Technologies, Inc.) according to the manufacturer’s
specifications, and were quantified using an Ultraspec2000
spectrophotometer (Pharmacia Biotech). RNA integrity was
assessed by ethidium bromide staining of 18S and 28S ribosomal bands.
Twenty micrograms of total cellular RNA were loaded onto a 1.0%
denaturing formaldehyde gel and resolved using electrophoresis. RNA was
transferred to positively charged 0.45-µm pore size nylon membranes
(Schleicher & Schuell, Inc., Keene, NH) and UV
cross-linked for 2 min (Stratagene, La Jolla, CA) to link
the RNA to the membranes. Membranes were prehybridized for 1–2 h at 68
C in ExpressHyb hybridization solution (CLONTECH Laboratories, Inc.), followed by hybridization with
[-P32]deoxy-CTP-labeled cDNA probes against
VEGF, TGFß1, or GAPDH in fresh rapid hybridization solution
(CLONTECH Laboratories, Inc.) for 2 h at 68 C.
Stringency washes were performed twice at room temperature with 2
x SSC-0.1% SDS (1 x SSC = 0.15 M NaCl-15
mM sodium citrate) for 10 min each, followed by two washes
in 0.1 x SSC-0.1% SDS at 50 C for 15 min each time. Membrane
signal intensity was quantified with a PhosphorImager (Molecular Dynamics, Inc., Sunnyvale, CA), and the resulting images were
analyzed using ImageQuant (Molecular Dynamics, Inc.) image
analysis software. All experiments were repeated in triplicate.
VEGF concentration in conditioned medium
A mouse VEGF quantitative sandwich enzyme immunoassay was used
to quantify VEGF production (R&D Systems, Minneapolis, MN). This assay
measures primarily the quantity of the 165-amino acid isoform of VEGF,
because it represents the main soluble isoform of VEGF. Assay and
controls were performed according to the manufacturer’s
recommendation. Briefly, 2 x 104 MC3T3-E1
cells were plated in each well of 3 24-well plates and allowed to reach
confluence over a period of 2–3 days in DMEM supplemented with 10%
FBS as described above. Once at confluence, medium was removed, and
cells were washed with PBS. Serum-free medium (400 µl) containing
antibiotic/antimycotic and recombinant human FGF-2 in concentrations of
0, 1.5, 3.1, 12.5, 25, and 50 ng/ml or either TGFß2 or TGFß3 in
concentrations of 0, 0.25, 0.5, 1, 5, and 50 ng/ml was then added to
the cultures. Additionally, after stimulating cultures with TGFß1 as
previously described (26), costimulatory experiments were performed in
which cultures were stimulated with both TGFß1 (0.31 and 0.63 ng/ml)
and FGF-2 (12.5 ng/ml). Each cytokine dose was repeated 4
times/experiment. After 24 h, the medium was removed and
centrifuged to remove particulate matter. A crystal violet colorimetric
assay was used to normalize the data for cell number (see below). All
experiments were repeated in triplicate.
Crystal violet staining
To minimize the effect of alterations in cellular proliferation
or plating, the number of plated cells was estimated using crystal
violet staining as described by Kueng et al. (43). Briefly,
cells were washed in PBS and fixed in ice-cold 3.7% paraformaldehyde
(Sigma) for 20 min. Cells were washed with PBS,
permeabilized with 20% methanol for 20 min, and stained with 0.5%
crystal violet (Sigma) in 20% methanol for 30 min. Excess
stain was removed after washes in deionized water, followed by elution
with 10% acetic acid for 30 min. The OD of the dye was measured at 650
nm using a SPECTRAmax 250 spectrophotometer (Molecular Devices, Menlo Park, CA).
Statistical analysis
All data from quantitative VEGF sandwich enzyme immunoassays are
expressed as the mean ± SD. Additionally, the
quantitative sandwich enzyme immunoassay and the crystal violet assay
underwent statistical significance testing with ANOVA (one-way ANOVA
comparing VEGF protein production by each dose of FGF-2, TGFß2, and
TGFß3). Post-hoc tests were performed using the
Tukey-Kramer multiple comparison test, with P < 0.05
considered significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). Initially,
FGF-2 stimulation yielded a 3.4-fold increase in VEGF mRNA by 3 h.
Increased VEGF mRNA expression was primarily noted in the 3.8-kb
isoform of VEGF mRNA. Maximal VEGF mRNA occurred at 6 h, with a
6-fold increase in VEGF mRNA followed by a decline to a 1.7-fold
increase in VEGF mRNA by 24 h. Similarly, when FRC cells were
stimulated with 12.5 ng/ml FGF-2, VEGF mRNA was increased at all time
points compared with unstimulated cells, with a 6-fold peak increase
occurring at 6 h (Fig. 2).
Importantly, the concentration of FGF-2 added to the cell cultures
falls within the range of previously reported physiologically relevant
levels (
10 ng/ml) (10, 20).
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). Blockade of transcription with
actinomycin D (5 µg/ml) decreased baseline and FGF-2-stimulated VEGF
mRNA expression. In contrast, translational blockade using
cycloheximide (10 µg/ml) did not alter baseline or FGF-2-stimulated
VEGF mRNA expression. Taken together, these findings suggest that the
regulation of VEGF expression by FGF-2 is primarily transcriptionally
mediated.
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). Five hours after treatment with
either vehicle or FGF-2, transcription was inhibited by the addition of
actinomycin D. The VEGF mRNA half-life of cells treated with FGF-2 was
not significantly different from the half-life of vehicle-treated
cellular mRNA, indicating that the rapid up-regulation of VEGF mRNA by
FGF-2 is largely transcriptionally (and not mRNA stabilization)
mediated.
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). To control for the effect that FGF-2
may have had on cell proliferation, only identically seeded, confluent
wells were stimulated with FGF-2. Additionally, a crystal violet assay,
performed at the time of medium collection, was used to standardize
relative cell number. FGF-2 produced dose-dependent increases in VEGF
production, with a maximal increase to 650 pg/ml after stimulation with
50 ng/ml FGF-2. Stimulation with 12.5 ng/ml FGF-2 increased VEGF
production to 450 pg/ml. With higher FGF-2 doses, the slope of VEGF
production decreased to a plateau, suggesting receptor saturation.
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). Unlike TGFß1, which in MC3T3-E1
cells induces a peak increase in VEGF mRNA after 3 h (26), neither
TGFß2 nor TGFß3 yielded early increases in VEGF mRNA. By 6 h,
however, VEGF mRNA was increased 3- and 2-fold in cells stimulated by
TGFß2 and TGFß3, respectively. By 24 h, VEGF mRNA levels
returned to baseline.
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). MC3T3-E1
stimulated cells yielded similar patterns of VEGF protein increases,
with maximal VEGF stimulation occurring at 5 ng/ml of either cytokine
followed by a sharp drop at a supraphysiological dose (50 ng/ml).
Consistent with the VEGF mRNA findings, TGFß2-stimulated cells
yielded higher absolute increases in VEGF protein than
TGFß3-stimulated cells (1900 vs. 1200; P =
0.005).
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).
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When control (uninfected or ß-galactosidase adenovirus-infected) or
dominant negative TGFß-truncated receptor II-transfected cells were
stimulated with FGF-2, VEGF mRNA levels remained similar between the
groups, with peak increases in VEGF mRNA at 6 h after stimulation
(Fig. 9). In contrast, VEGF mRNA levels
were significantly curtailed in TGFß1-stimulated cells transfected
with dominant negative TGFß-truncated receptor II at all time points
(Fig. 9). TGFß1-stimulated control cells demonstrated the expected
3 h peak increases in VEGF mRNA (data not shown). These data
present evidence that FGF-2 may modulate the up-regulation of VEGF mRNA
independently of TGFß.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Although the roles of FGF-2 in gene patterning and, more specifically, skeletal development are areas of intensive investigation (10, 11, 12, 13, 50, 51), much in vitro research on the effects of FGF-2 on osteoblastic cells has focused on differentiation, proliferation, and extracellular matrix elaboration (17, 52, 53, 54). The significance of angiogenesis during development is well documented and by nature is coincident with skeletal development; however, bone remodeling and repair are life-long processes. Indeed, the fracture milieu contains many of the conditions and factors that have, in other tissues, been found to promote VEGF expression [hypoxia (55, 56), elevated FGF-2 (21), TGFß1 (57), and interleukin 1 (56)]. It is important, therefore, to identify potential mechanisms of FGF-2-induced angiogenesis by osteoblastic cells. We found that physiologically relevant FGF-2 doses promoted VEGF mRNA and protein expression in clonal and nonimmortalized osteoblastic cells. Although static mRNA expression as observed in Northern analysis is difficult to correlate directly with cumulative protein production measured by immunoassay, the finding that both mRNA and protein expression of VEGF are increased by FGF-2 stimulation supports the hypothesis that FGF-2 is a regulator of VEGF synthesis. Similar to the actions of TGFß1, insulin-like growth factor I, PGE1, and PGE2 (25, 26, 28), VEGF mRNA stability was unaffected by FGF-2, whereas actinomycin D, but not cycloheximide, strongly decreased FGF-2 induction of VEGF mRNA, suggesting transcriptional control of FGF-2-mediated increases in VEGF mRNA.
The ability of FGF-2 to induce angiogenesis directly and through VEGF is complicated by the fact that FGF-2 also activates and induces TGFß1, which, itself, modulates angiogenesis and stimulates VEGF production. For example, TGFß1 promotes angiogenesis in both the rabbit corneal micropocket (58) and the chick chorioallantoic membrane (59) models. Furthermore, TGFß1 increases VEGF mRNA and protein in osteoblastic cells, and both growth factors have been colocalized in a healing fracture (26). In contrast, genetically induced TGFß1 overexpression in arteries, liver, epidermis, and respiratory epithelial cells does not result in angiogenesis (60). To account for these diverse effects, a theory has arisen that the microenvironment within which TGFß1 is expressed determines the ultimate biological actions of TGFß1. Specifically, in an inflammatory microenvironment, TGFß1 levels correlate directly with increased angiogenesis (60). This is consistent with the concept that the indirect modulation of angiogenesis by TGFß1 may occur through inflammatory cells temporarily recruited to a site of injury. Thus, given the proper environment and/or effector cells, TGFß1 can be involved in the production of direct angiogenic factors such as VEGF.
To further investigate TGFß, we have demonstrated that its other mammalian isoforms (TGFß2 and TGFß3) also increase VEGF mRNA and protein, again with physiologically relevant doses. Unlike TGFß1, which maximally increases VEGF mRNA at 3 h (26, 28), both TGFß2 and TGFß3 maximally increased VEGF mRNA at 6 h, albeit with lower maximal potency (2- to 3-fold for TGFß3 and TGFß2, respectively, vs. 6-fold for TGFß1). Similar to TGFß1, both TGFß2 and TGFß3 cause dose-dependent increases in VEGF protein production.
As all TGFß isoforms increased VEGF, we investigated the possibility that FGF-2 mediated its induction of VEGF via activation or induction of TGFß. We demonstrated that costimulation with both TGFß1 and FGF-2 yielded additive, and not synergistic, increases in VEGF protein production. Additionally, after blocking TGFß signal transduction with a dominant negative truncated TGFß1 receptor II virus, FGF-2-induced VEGF mRNA was maintained, demonstrating TGFß-independent modulation of VEGF mRNA by FGF-2. Despite these findings, it is not unlikely that FGF-2 induction of VEGF through TGFß occurs physiologically and that this may represent a redundant mechanism of VEGF up-regulation.
The mitogenic and remodeling effects of VEGF on capillary endothelial cells and the ability of FGF-2 to increase VEGF expression by the endothelium suggests that FGF-2 may play an important role in the angiogenic response evidenced by healing bone. Additionally, the increase in VEGF during bone healing (26) may be at least partially an indirect effect of FGF-2, because, as indicated above, FGF-2 increases the levels of several cytokines that have been implicated in VEGF expression. Finally, the effect of FGF-2 on VEGF may by synergistically enhanced by other cytokines and conditions (strain, hypoxia) present in the fracture milieu. A fracture creates the necessary environment for FGF-2 elaboration by osteoblasts, and it is likely that this inflammatory microenvironment sets the stage for the production of VEGF and other direct angiogenic cytokines, without which fracture vascularization, and hence healing, cannot occur. Similarly, significant evidence separately implicates FGF-2 and the TGFß isoforms in skeletal development and VEGF in embryogenesis. These in vivo circumstantial relationships have been explored by investigating possible mechanisms of control of VEGF expression in osteoblastic cells. A better understanding of these mechanisms may lay the groundwork for future manipulations of the developing bone microenvironment and ultimately improve the treatment of bone pathology.
Received May 25, 1999.
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