1 Department of Surgery, University of Connecticut, Farmington, Connecticut 06032; 2 Laboratory of Developmental Biology and Repair, Department of Surgery, New York University School of Medicine, New York, New York 10016; and 3 Department of Cardiology, Kyushu University School of Medicine, Fukuoka 812, Japan
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ABSTRACT |
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Angiogenesis is essential to both normal and pathological bone
physiology. Vascular endothelial growth factor (VEGF) has been implicated in angiogenesis, whereas transforming growth factor-1 (TGF-
1) modulates bone differentiation, matrix
formation, and cytokine expression. The purpose of this study was to
investigate the relationship between TGF-
1 and VEGF expression in
osteoblasts and osteoblast-like cells. Northern blot analysis revealed
an early peak of VEGF mRNA (6-fold at 3 h) in fetal rat calvarial cells
and MC3T3-E1 osteoblast-like cells after stimulation with TGF-
1 (2.5 ng/ml). The stability of VEGF mRNA in MC3T3-E1 cells was not increased
after TGF-
1 treatment. Actinomycin D inhibited the TGF-
1-induced
peak in VEGF mRNA, whereas cycloheximide did not. Blockade of TGF-
1
signal transduction via a dominant-negative receptor II adenovirus
significantly decreased TGF-
1 induction of VEGF mRNA. Additionally,
TGF-
1 induced a dose-dependent increase in VEGF protein expression
by MC3T3-E1 cells (P < 0.01).
Dexamethasone similarly inhibited VEGF protein expression. Both
TGF-
1 mRNA and VEGF mRNA were concurrently present in rat membranous
bone, and both followed similar patterns of expression during rat
mandibular fracture healing (mRNA and protein). In summary,
TGF-
1-induced VEGF expression by osteoblasts and osteoblast-like
cells is a dose-dependent event that may be intimately related to bone
development and fracture healing.
angiogenesis; bone; fracture
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INTRODUCTION |
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normal and pathological bone physiology is inexorably tied to angiogenesis. The process of bone development and repair depends on the adequate formation of new capillaries from existing blood vessels (37). The osteon forms around a haversian canal containing blood vessels that supply osteoblasts with needed oxygen and nutrients. During both intramembranous and endochondral ossification, bone spicules and osteogenic buds, respectively, delineate and surround capillaries. Additionally, osteocyte survival requires a <0.1-mm proximity to nutrient vessels (17), and interruption of the blood supply to bone results in avascular necrosis (9). Furthermore, vascularized bone grafts maintain more osseous mass than nonvascularized bone grafts (10).
As molecular mechanisms of angiogenesis become defined, cytokines and their interrelationships appear to play a crucial role in the formation, growth, and regression of blood vessels. Vascular endothelial growth factor (VEGF), a dimeric heparin-binding glycoprotein, is assuming an increasingly central role in the understanding of the development and modulation of angiogenesis. VEGF is expressed in highly vascular tissues and is an endothelial cell-specific mitogen (23). VEGF receptor knockout mice lack adequate blood vessel formation (34), whereas loss of a single VEGF allele is lethal in the mouse embryo (12). Devascularized rat islets of Langerhans cells and hypoxic human vascular smooth muscle cells demonstrate increased VEGF expression (5).
Fracture healing requires adequate angiogenesis, and it is within this
context that VEGF may also play an important role. VEGF is expressed in
the normal rat tibia (19), whereas both intramembranous and
endochondral ossification is associated with capillary development (8).
Additionally, VEGF expression in osteoblasts and osteoblast-like cells
is increased by several cytokines and growth factors, including
prostaglandin E1
(PGE1) and
PGE2, insulin-like
growth factor (IGF), platelet-derived growth factor, and
1,25-dihydroxyvitamin D3 (14,
19, 38).
Transforming growth factor-1 (TGF-
1), a ubiquitous cytokine with
wide-ranging effects, has been implicated in osteoblast proliferation
and differentiation. Moreover, TGF-
1, the largest source of which is
bone (3), is expressed at high levels during bone growth and
development (25, 33), processes that depend on an adequate blood
supply. The observation that TGF-
1 is both a potent inducer and a
potent inhibitor of angiogenesis has given rise to the concept that, to
promote angiogenesis, TGF-
1 requires an inflammatory environment
(28). It is within this inflammatory "context" that TGF-
1 is
then able to effect an angiogenic cascade. Support for this concept of
"indirect" angiogenesis is provided by the observation that
TGF-
1 increases VEGF production in human smooth muscle cells, mouse
fibroblasts, human lung adenocarcinoma cells, and human histiocytic
lymphoma cells (28).
An increasing body of evidence implicates TGF-1 in fracture healing.
TGF-
1 stimulates osteoblast migration, modulates osteoblast proliferation, and is localized to cells within the developing skeleton
(11, 29). Additionally, TGF-
1 expression is increased in fracture
healing (21), and exogenous application of TGF-
1 accelerates both
endochondral bone fracture healing and the closure of membranous bone
critical size defects (2, 24). Despite the above findings, the
relationship between TGF-
1 and VEGF in bone growth and healing
remains undefined.
Given the importance of TGF-1 and VEGF in the related processes of
angiogenesis and fracture healing, we proposed that TGF-
1 may
regulate VEGF expression in osteoblasts. We demonstrated that TGF-
1
increased VEGF mRNA in both primary and clonal osteoblasts. Significant
control of this mechanism occurred at the transcriptional level in
clonal osteoblasts. Overexpression of a dominant-negative receptor II
by adenovirus-mediated gene transfer disrupted TGF-
1 signal
transduction and significantly decreased stimulation of VEGF mRNA by
exogenous TGF-
1. Additionally, TGF-
1 increased VEGF protein
production by osteoblastic cells, and this increase was inhibited by
dexamethasone. Finally, synchronously modulating levels of both
TGF-
1 and VEGF mRNA and protein expression were found during
membranous bone fracture healing.
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MATERIALS AND METHODS |
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Materials.
Tissue culture plates and flasks were purchased from Fisher Scientific
(Pittsburgh, PA). DMEM, -modified Eagle's medium, 0.05%
trypsin-EDTA, PBS, fetal bovine serum (FBS), and cell culture reagents
were purchased from Life Technologies (Gaithersburg, MD). Recombinant
human TGF-
1 (Life Technologies) was prepared in 100 ng/5 ml PBS
aliquots and was stored at
20°C. Actinomycin D,
cycloheximide, and dexamethasone were from Sigma (St. Louis, MO).
Cell culture. Fetal rat calvarial (FRC) cells were cultured from FRC explants by a modification of the procedure described by Freshney (13). Briefly, frontal and parietal bones from gestational 21-day Sprague-Dawley fetal rats were sterilely stripped of their periosteum and minced into 1-mm3 fragments. The explants were washed with sterile PBS containing an antibiotic and antimycotic and then placed at the bottom of an upright 25-cm2 flask containing preincubated media (DMEM supplemented with 10% FBS, 100 µg/ml penicillin G, 50 µg/ml streptomycin, and 0.25 µg/ml amphotericin B). After 15 min at 37°C, the flask was slowly returned to a horizontal position, and the culture was maintained in a humidified atmosphere consisting of 95% air-5% CO2 at 37°C. Media were changed, and when the cells were confluent, the explant pieces were removed and the cells were trypsinized and transferred to 75-cm2 flasks. Passage 2 cells were used for all experiments. Verification of osteoblastic lineage was performed by mineralized bone nodule formation assay and Northern analysis for osteocalcin (data not shown).
MC3T3-E1 cells, a mouse clonal osteoblastic cell line, were grown in DMEM supplemented with 10% FBS, 100 µg/ml penicillin G, 50 µg/ml streptomycin, and 0.25 µg/ml amphotericin B. Media were changed every 2-3 days. Confluent MC3T3-E1 cell cultures 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.Animals. Adult male Sprague-Dawley rats (250-350 g) were purchased from Taconic Laboratories (Germantown, NY) and housed in separate cages. Animals were kept under a constant 12-h light-dark schedule and fed Purina rodent chow ad libitum. Surgical procedures were approved by the Institutional Care and Use Committee at New York University Medical Center. Anesthesia for all operative procedures was achieved with a mixture of Ketaset (7.5 mg/kg body wt; Fort Dodge Animal Health, Fort Dodge, IA), xylazine (1.5 mg/kg; Bayer Animal Health, Shawnee Mission, KS), and acepromazine maleate (0.25 mg/kg; Fermenta Animal Health; Kansas City, MO).
Animal surgery.
Twenty-one adult male rats were used in this study. Three rats
underwent a sham operation with a 1-cm skin incision made along the
inferior border of the right mandible, separation of the rat masseter
muscle, and exposure of the mandible without performing an osteotomy.
In 18 experimental animals, the body of the right hemimandible was
similarly exposed; however an osteotomy was performed in a copiously
irrigated field between the second and third molars (Fig.
1) with an 8-mm, double-sided
diamond disc (Brassler, Savannah, GA). A pair of 1-mm bicortical holes
were drilled 4 mm anterior and posterior to the osteotomy and two 1.5 × 20-mm Flexi-Post pins (Essential Dental Systems, South
Hackensack, NJ) were screwed into the holes. A prefabricated external
fixator was attached to the pins (Fig. 1). The field was irrigated
copiously with sterile saline, and the skin and soft tissues were
reapproximated with resorbable sutures. Buprenorphine (0.1 mg/kg; Fort
Dodge Animal Health) was administered for 1 day postoperatively for
pain management. Twelve animals were killed on postoperative
days 3,
9, and
23; the right hemimandible was
dissected free of skin and soft tissues; and the section of the
mandible immediately bounded by the pins was resected, snap frozen in
liquid nitrogen, and homogenized with a Polytron tissue homogenizer
(Kinematica). Total cellular RNA was extracted with TRIzol solution as
outlined below. In the remaining six animals, mandibles harvested on
postoperative days 9 and
23 were fixed in 4% paraformaldehyde
and decalcified in Immunocal (Decal Chemical, Congers, NY) for 6 days.
Tissues were then placed in 30% sucrose solution for 2 days and
embedded in tissue-freezing medium (Triangle Biomedical Sciences,
Durham, NC), and 10-mm sections were prepared for immunohistochemistry as outlined below.
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Probe preparation.
Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was a 1-kb probe from
Clontech (Palo Alto, CA). A 535-bp probe against rat TGF-1 and a
411-bp probe against mouse VEGF were generated by PCR from whole rat
and mouse embryo cDNA, respectively (16). PCR primers for TGF-
1 and
VEGF have been previously described (4, 15). PCR bands were gel
purified, cloned into PCR.1 plasmids (Invitrogen, Carlsbad, CA), and
sequenced to confirm sequence identity. The probe was generated after
EcoR I digestion and gel purification.
One hundred nanograms of each probe were labeled with
32P-labeled deoxycytidine
triphosphate
([
-32P]dCTP) by
using random oligonucleotide primers and Klenow fragment (Ready To Go
labeling beads; Pharmacia Biotech, Cambridge, England). Unincorporated
nucleotides were removed with Sephadex G-50 DNA-grade nick columns
(Pharmacia Biotech). All probes had specific activities >105 cpm/ml of hybridization solution.
RNA extraction and Northern analysis.
Subconfluent FRC (passage 2) and
MC3T3-E1 cells in 100-mm2 plates
were stimulated with 2.5 ng/ml TGF-1 in antibiotic-containing, serum-free media for 0, 3, 6, or 24 h. In experiments designed to
investigate the effect of TGF-
1 on VEGF mRNA stability,
transcription was interrupted with actinomycin D (5 µg/ml) after 2 h
of stimulation with TGF-
1 (2.5 ng/ml) in serum-free media. To
investigate protein synthesis or gene transcription, subconfluent
MC3T3-E1 cells in 100-mm2 plates
underwent 3-h exposures to cycloheximide (10 µg/ml) or actinomycin D
(5 µg/ml) with or without TGF-
1 (2.5 ng/ml;
±TGF-
1). Cells in the actinomycin D group (but not
the cycloheximide group) underwent 1 h of pretreatment with actinomycin
D before TGF-
1 stimulation (14).
Northern blot analysis.
Total cellular RNA was extracted with TRIzol solution (Life
Technologies) according to the manufacturer's specifications, and
quantified with 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 by
electrophoresis. RNA was transferred to positively charged 0.45-µm
nylon membranes (Schleicher & Schuell, 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); this was followed by hybridization
with
[-32P]dCTP-labeled
cDNA probes against VEGF, TGF-
1, or GAPDH in fresh Rapid
Hybridization solution (Clontech) for 2 h at 68°C. Stringency washes were performed twice at room temperature with 2× SSC
(1× SSC = 0.15 M NaCl-15 mM sodium citrate)-0.1% SDS for 10 min
each and were followed by two washes in 0.1× SSC-0.1% SDS at
50°C for 15 min each. Membrane signal intensity was quantified with
a PhosphorImager (Molecular Dynamics, Sunnyvale, CA), and the resulting
images were analyzed with ImageQuant (Molecular Dynamics) image
analysis software. All experiments were repeated in triplicate.
VEGF concentration in conditioned media.
A mouse VEGF quantitative sandwich enzyme immunoassay was purchased
from R&D Systems (Minneapolis, MN). Assay and controls were performed
in accordance with the manufacturer's recommendations. Briefly, 2 × 104 MC3T3-E1 cells were
plated in each well of a 24-well plate and allowed to reach confluence
over a period of 2-3 days in DMEM supplemented with 10% FBS as
described above. Once at confluence, media were removed and cells were
washed with PBS. Serum free media (400 µl) containing an antibiotic
and antimycotic and recombinant human TGF-1 in concentrations of 0, 0.62, 1.25, 2.5, 5, 12.5, and 25 ng/ml were then added to the cultures.
Additionally, as a separate experiment, dexamethasone (1 × 10
7, 2 × 10
7, and 4 × 10
7 M) was coadded to
separate wells stimulated with 5 ng/ml TGF-
1. Each cytokine dose was
repeated four times per experiment. After 24 h, the media were removed
and centrifuged to remove particulate matter. Equal cell numbers
between wells were further verified by a crystal violet colorimetric
assay (see below). All experiments were repeated in triplicate.
Crystal violet staining. To minimize the effect of alterations in cellular proliferation or equal plating, the number of plated cells was estimated by crystal violet staining as described by Kueng et al. (22). Briefly, cells were washed in PBS and fixed in ice-cold 3.7% paraformaldehyde (Sigma) for 20 min. Cells were washed with PBS, permeablized 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 optical density of the dye was measured at 650 nm with a SPECTRAmax 250 spectrophotometer (Molecular Devices, Sunnyvale, CA).
Immunohistochemistry.
Affinity-purified rabbit polyclonal anti-TGF-1 and anti-VEGF
antibodies with no cross-reactivity were used in all experiments (R&D
Systems). After fixing fracture tissues from days
9 and 23, immunohistochemistry was performed as previously described (32). Briefly, 10-µm tissue sections were placed on SuperFrost Plus slides
(Fisher Scientific), endogenous peroxidase activity was quenched with
0.6% hydrogen peroxide in methanol, and antigen unmasking was
performed with 10 mM sodium citrate. Nonspecific staining was blocked
by incubating sections with normal goat serum (1.5%; Vector
Laboratories, Burlingame, CA) followed by antisera against TGF-
1 or
VEGF overnight at 4°C. Biotinylated goat anti-rabbit secondary antibodies (Vector Laboratories) and avidin-biotin peroxidase complex were successively applied, and positive staining was visualized with 3,3'diaminobenzidine (Sigma) as the substrate to cause brown staining of positively stained tissues. Sections were counterstained with Harris hematoxylin. Control slides were incubated in nonimmune rabbit serum or no primary antibody and processed identically to
experimental sections. All experiments were performed in triplicate.
Statistical analysis.
All data from the quantitative VEGF sandwich enzyme immunoassay are
expressed as means ± SD. Additionally, the quantitative sandwich enzyme immunoassay and the crystal violet assay underwent statistical significance testing with one-way ANOVA to compare levels
of VEGF protein production by the different doses of TGF-1. Post hoc
tests consisted of the Tukey-Kramer multiple comparison test, with
P < 0.05 considered significant.
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RESULTS |
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TGF-1 increased VEGF mRNA levels in MC3T3-E1
osteoblast-like cells and primary osteoblasts.
MC3T3-E1 mouse clonal osteoblasts express osteoblastic features such as
collagen type I and alkaline phosphatase and they behave similarly to
primary osteoblasts in response to TGF-
1 (3). When osteoblastic
cells were stimulated with 2.5 ng/ml TGF-
1, VEGF mRNA was increased
at all time points compared with levels in unstimulated cells (Fig.
2). Maximal VEGF mRNA occurred early, with
a 6-fold increase in VEGF mRNA at 3 h followed by decreases to 2.5- and
1.5-fold inductions of VEGF mRNA expression at 6 and 24 h respectively.
Similarly, when the FRC cells were stimulated with 2.5 ng/ml TGF-
1,
VEGF mRNA was increased at all time points compared with levels in
unstimulated cells and the peak increase occurred at 3 h
(Fig. 3). Importantly, the concentration of
TGF-
1 added to the cell cultures falls within the range of previously reported physiologically relevant levels (~1 ng/ml).
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Effect of mRNA and protein synthesis inhibitors on
TGF-1 stimulation of VEGF mRNA.
The short, sharp rise of VEGF mRNA followed by its rapid decline in
TGF-
1-stimulated primary osteoblasts and osteoblastic cells was
consistent with the pattern shown by other osteogenic cytokines (19)
and suggested high turnover and low stability of VEGF mRNA. To further
define the mechanisms of action of TGF-
1 stimulation of VEGF
expression in osteoblastic cells, we employed inhibitors of RNA
polymerase and protein synthesis to block transcription and
translation, respectively (Fig. 4).
Blockade of transcription with actinomycin D (5 µg/ml) decreased the
baseline VEGF mRNA. When protein production was blocked with
cycloheximide (10 µg/ml), VEGF mRNA was still produced. The
TGF-
1-induced increase in VEGF expression was blocked by actinomycin
D. In contrast, disruption of translation with cycloheximide did not
substantially reduce VEGF mRNA expression in response to TGF-
1
stimulation, suggesting that TGF-
1 exerted its effects at the
transcriptional level.
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TGF- signal blockade reduced TGF-
1
stimulation of VEGF mRNA.
TGF-
has three defined isoforms in mammals, TGF-
1, TGF-
2, and
TGF-
3. In addition to effecting differential modulation of gene
expression, these isoforms may act through autocrine mechanisms to
regulate both their own expression and the expression of their common
receptor, transforming growth factor receptor II.
Although neutralizing antibody studies are an effective way to isolate cytokine effects, the multiple isoforms of TGF-
, and the biphasic effects of TGF-
1 in particular, make TGF receptor II an elegant target for manipulation. Overexpression of a dominant-negative TGF
receptor II via an adenovirus vector allowed for disruption of TGF-
signal transduction because this construct binds all TGF-
isoforms
but does not allow phosphorylation of TGF-
receptor I. When
exogenous TGF-
1 was added to control (uninfected or
-galactosidase adenovirus-infected) cells, the previously described
increase in VEGF expression (peak at 3 h) was identified (Fig.
6). However, when cells were transfected
with truncated dominant-negative TGF receptor II, the expression of
VEGF was significantly curtailed at all time points. These data are
further evidence that TGF-
1 acts through its secondary receptors to
directly effect the upregulation of VEGF mRNA.
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Effect of TGF-1 on VEGF concentration in conditioned
medium.
We then proceeded to examine the production of VEGF protein as a result
of TGF-
1 stimulation. Additionally, we examined the dose-response relationship between TGF-
1 and VEGF. The basal level
of VEGF production by MC3T3-E1 cells was 300 pg/ml at 24 h (Fig.
7). To control for the effect that TGF-
1
may have had on cell proliferation, only identically seeded, confluent
wells were stimulated with TGF-
1. Additionally, a crystal violet
assay, performed at the time of medium collection, did not demonstrate statistically significant well-to-well variation in cell number (data
not shown). TGF-
1 produced a dose-dependent increase in VEGF
production with a maximal increase to 1,200 pg/ml at 25 ng/ml TGF-
1.
At 2.5 ng/ml TGF-
1, VEGF production was 1,050 pg/ml. Thereafter, the
slope of VEGF production decreased to a plateau, suggesting receptor
saturation. TGF-
1 had a significant effect on VEGF protein
production at the physiologically relevant doses of 0.63 and 1.25 ng/ml, with increases in VEGF production to 600 and 900 pg/ml,
respectively.
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Effect of dexamethasone on TGF-1-stimulated VEGF
production in conditioned medium.
To consider the effects of glucocorticoids on TGF-
1-induced VEGF
expression in bone, dexamethasone was added to the culture system.
Dexamethasone significantly inhibited TGF-
1 stimulation of VEGF
expression in a dose-dependent fashion (Fig.
8). This decrease in a potential angiogenic
response may provide an insight into the molecular mechanisms of
glucocorticoid impairment of bone healing.
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TGF-1 and VEGF mRNA are coexpressed during
membranous bone fracture healing in vivo.
Having identified the effect of TGF-
1 on VEGF
expression by osteoblasts in vitro, we investigated the presence of
TGF-
1 and VEGF in vivo in both unfractured and fractured membranous bone. An osteotomy was created behind the second molar of the rat right
hemimandible, and a mandibular fixation device was applied as shown in
Fig. 1. Total cellular RNA was isolated from fractured rat mandibles
(3, 9, and 23 days after the operation) and was analyzed by Northern
blotting with a TGF-
1 probe. TGF-
1 mRNA was identified at all
time points and increased with time after fracture (Fig.
9). The greatest TGF-
1 mRNA signal
occurred at 23 days, the middle period of fracture consolidation. We
next sought to identify the concurrent presence of VEGF mRNA.
Therefore, identical specimens of total cellular RNA from fractured rat
mandibles were analyzed by Northern blotting with a mouse VEGF probe
(Fig. 9). As found for TGF-
1, VEGF mRNA increased during fracture
healing with the strongest increase also occurring 23 days after
fracture. Interestingly, at 9 days after the operation,
the increase in VEGF mRNA appeared greater than the increase in
TGF-
1 mRNA. Finally, unfractured mandibles revealed levels of
TGF-
1 and VEGF mRNA similar to those at 23 days after the operation
(data not show).
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TGF-1 and VEGF proteins are both present in the
fracture callus during membranous fracture healing.
Immunohistochemical staining of a fracture site revealed the presence
of both TGF-
1 and VEGF (Figs. 10 and
11). As the bone healed,
TGF-
1 was strongly localized to the fracture callus and the
proliferating osteoblasts within and immediately surrounding the callus
(Fig. 10,
A-D).
Likewise, throughout fracture healing, the fracture callus demonstrated
strong VEGF staining, as did the proliferating osteoblasts within and
immediately surrounding the callus (Fig. 11,
A-F).
TGF-
1 staining was both cytoplasmic and matrix associated, whereas
VEGF staining was primarily cytoplasmic. Furthermore, although
osteoblasts stained strongly for VEGF, osteocytes did not appear to be
expressing VEGF, suggesting tightly localized control of VEGF
expression. Control groups (nonimmune rabbit serum or no primary
antibody) did not demonstrate positive staining (data not shown).
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DISCUSSION |
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The wide-ranging presence and effects of TGF-1 underscore its
fundamental role as an orchestrator of biomolecular events. Furthermore, its ubiquitous nature and sometimes discrepant effects both encourage and complicate its study. It has been observed that
TGF-
1 can stimulate or inhibit angiogenesis in vivo. For example,
TGF-
1 promotes angiogenesis in both the rabbit corneal micropocket
(30) and the chick chorioallantoic membrane (40) models. In contrast,
genetically induced TGF-
1 overexpression in arteries, liver,
epidermis, and respiratory epithelial cells does not result in
angiogenesis (28). Additionally, TGF-
1 inhibits both cultured
endothelial cell proliferation (1) and fibroblast growth factor
(FGF)-induced angiogenesis in a subcutaneous sponge system (27). These apparently discrepant effects have given rise to the
notion that the microenvironment within which TGF-
1 is expressed
determines TGF-
1's ultimate biological effects. More specifically,
it has been observed that, within the context of inflammation,
increased TGF-
1 levels correlate with increased angiogenesis (28).
This biology may be due, in part, to the presence of a specialized,
recruited subset of inflammatory cells, which are known to produce both
inflammatory and angiogenic cytokines. This, in turn, is consistent
with the concept that TGF-
1 is an indirect cytokine of angiogenesis.
Thus, given the proper environment and/or effector cells, TGF-
1 can
be involved in the production of direct angiogenic factors such as VEGF.
Because the fracture milieu contains many of the conditions and factors
that have, in other tissues, been found to promote VEGF expression
[hypoxia (20, 35), elevated TGF-1 (5), FGF-2 (26), and
interleukin-1 (20)], we sought to isolate the
effect of TGF-
1 on VEGF production by osteoblastic cells in vitro.
We found that TGF-
1 increased the expression of VEGF mRNA by primary
osteoblasts and osteoblastic cells. This effect occurred at a dose of
2.5 ng/ml TGF-
1, which lies within the mammalian physiological range
and is similar to the concentration that upregulates VEGF mRNA in
vascular smooth muscle cells (5).
The TGF-1-induced increase in VEGF mRNA appears to be primarily a
transcriptionally mediated event. As found for IGF-I,
PGE1, and
PGE2, VEGF mRNA stability was
unaffected by TGF-
1, whereas actinomycin D, but not cycloheximide,
strongly decreased TGF-
1 induction of VEGF mRNA. When TGF-
1
signal transmission was disrupted by an overexpressed dominant-negative
receptor II, the increase in VEGF mRNA was greatly attenuated.
Although the peak increase in VEGF mRNA was acute, VEGF mRNA remained
elevated throughout the stimulation period. Furthermore, TGF-1
stimulated a dose-dependent increase in VEGF protein production by
MC3T3-E1 osteoblastic cells, and this response began at physiologically relevant TGF-
1 doses. These observations were consistent with our in
vivo findings of VEGF mRNA expression during fracture healing and
localization of VEGF protein to a healing fracture callus. The
mitogenic and remodeling effect of VEGF on capillary endothelial cells
and the ability of TGF-
1 to increase VEGF expression by the
endothelium suggest that TGF-
1 plays an important role in the
angiogenic response evidenced by healing bone.
The impairment of the vascular supply to bone results in avascular
necrosis (6). Traditional theories regarding the mechanism of
glucocorticoid-induced avascular necrosis center on abnormal fat
metabolism, with resultant fat embolism causing vascular occlusion and
subsequent bone death (7). Smith (36) has hypothesized that the
pathogenesis of avascular necrosis of the femoral head is based on the
inhibition of angiogenesis. Dexamethasone has been shown to inhibit
angiogenesis in vitro (18) and to block prostaglandin stimulation of
VEGF production in osteoblastic cells (19). In vitro, dexamethasone
produced a dose-dependent inhibition of TGF-1-induced VEGF protein
production and may provide an additional molecular
explanation for the well-observed phenomenon of both impaired fracture
vascularization and healing in glucocorticoid-treated patients.
We also demonstrated the concurrent presence of TGF-1 and VEGF mRNA
in bone. Additionally, we found that, during membranous bone fracture
healing, TGF-
1 and VEGF mRNAs have similar patterns of expression,
with both being expressed during early fracture healing and both
increasing during mineralization. Although the increase in VEGF mRNA at
9 days after the operation was greater than that of TGF-
1 mRNA, this
may simply be testimony to the potency of TGF-
1 as an inducer of
VEGF production. Additionally, the increase of VEGF during this
inflammatory phase may be, at least partially, an indirect effect of
TGF-
1, because TGF-
1 increases the levels of several cytokines
(FGF-2, and TGF-
1 itself) that have been implicated in VEGF
expression. Finally, the effect of TGF-
1 on VEGF may by
synergistically enhanced by other cytokines and conditions (strain,
hypoxia) present in the fracture milieu. Immunohistochemical analysis
of the fracture site was consistent with the mRNA findings and revealed
strong staining of osteoblasts within the fracture callus for both
TGF-
1 and VEGF protein. Administration of exogenous recombinant
TGF-
1 protein has been shown to promote healing of both endochondral
fractures (24) and membranous defects (2). A fracture creates the
necessary environment for release of TGF-
1 from platelets and bone
(the largest two reservoirs of latent TGF-
1), activation of latent
TGF-
1 by acidic conditions and plasminogen (41), and upregulation of
TGF-
1 expression in an autocrine and paracrine fashion (31). 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.
These data further advance the concepts of contextual and indirect
activity of TGF-1 as a promoter of angiogenesis. Both the in vivo
and the in vitro findings supplement the current understanding of the
fracture microenvironment. It is likely that VEGF is produced and acts
in concert with a multitude of other factors involved in angiogenesis
during fracture healing. We are actively defining these factors and
their interrelationships. We remain hopeful that further understanding
of angiogenic mechanisms in bone healing will provide the foundation
for therapeutic molecular manipulations to improve bone healing.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: M. T. Longaker, Laboratory of Developmental Biology and Repair, Rm. H-169, New York Univ. Medical Center, 550 First Ave., New York, NY 10016 (E-mail: michael.longaker{at}med.nyu.edu).
Received 5 February 1999; accepted in final form 2 June 1999.
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REFERENCES |
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Baird, A.,
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