Division of Nephrology, Program in Developmental Biology, The Hospital for Sick Children, University of Toronto, Toronto, Ontario, Canada M5G 1X8
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ABSTRACT |
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Bone morphogenetic
protein-7 (BMP7) controls ureteric bud and collecting duct
morphogenesis in a dose-dependent manner (Piscione TD, Yager TD, Gupta
IR, Grinfeld B, Pei Y, Attisono L, Wrana JL, and Rosenblum ND.
Am J Physiol Renal Physiol 273: F961-F975, 1997). We defined cellular and molecular mechanisms underlying these effects
in embryonic kidney explants and in the mIMCD-3 cell model of
collecting tubule morphogenesis. Low-dose (0.25 nM) BMP7
significantly increased tubule number and cell proliferation.
Similar to BMP2, high-dose (10 nM) BMP7 inhibited cell
proliferation and stimulated apoptosis. To define molecular mechanisms,
we identified signaling events downstream of BMP7. High-dose BMP7, but
not low-dose BMP7, activated Smad1 in mIMCD-3 cells. Moreover,
the inhibitory effects of high-dose BMP7 and BMP2, but not the
stimulatory effects of low-dose BMP7, on tubulogenesis and cell
proliferation were significantly reduced in mIMCD-3 cells stably
expressing Smad1(458), a dominant negative mutant form of
Smad1, but not in cells stably expressing wild-type Smad1. We conclude
that BMP7 exerts dose-dependent effects on ureteric bud or collecting
duct cell proliferation and apoptosis by signaling via Smad1-dependent
and Smad1-independent pathways.
bone morphogenetic protein-7; renal collecting system; bone morphogenetic protein-2; branching morphogenesis
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INTRODUCTION |
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CONTROL OF CELL PROLIFERATION and apoptosis is fundamental to the formation of complex tissues during embryogenesis. In the mammalian kidney, the collecting system arises via growth and branching of the ureteric bud and its daughter collecting ducts, a process termed branching morphogenesis (30). Cell proliferation and apoptosis are both spatially and temporally regulated during renal branching morphogenesis. Although cell proliferation occurs in all regions, it is highest at the tips of the branching ureteric bud and collecting ducts. In contrast, apoptosis is comparatively infrequent in the trunk and tips of the ureteric bud and collecting ducts but is prominent in the innermost regions that give rise to the calyces and pelvis (7, 29).
Secreted growth factors including bone morphogenetic proteins (BMP) regulate cell proliferation and apoptosis during morphogenesis of nonrenal tissues. The cellular response to BMP family members is dependent on the differentiated state of the cell and the particular BMP that is bound to the cell (5, 24). For example, BMP2 inhibits cell proliferation in astrocytes but promotes proliferation in mouse embryo cells cultured in the absence of serum (8). BMP7 stimulates cell proliferation in embryonic neurectoderm (3) and osteoblasts (6) but inhibits proliferation of embryonal carcinoma cells (2). BMP2 induces apoptosis in sympathoadrenal cells, but BMP7 has no effect (32). The response to BMP family members is also dependent on the stage of embryogenesis at which they are active (11). For example, at embryonic day 13 (E13), BMP2 increases apoptosis and inhibits proliferation of mouse cortical ventricular zone cells. In contrast, at E16, concentrations of 0.3-3 nM promote neural differentiation, whereas 10-fold higher concentrations promote cell death (21). Taken together, these observations suggest that the cellular response to BMPs in the kidney may be dependent on factors including the phenotype of the target cell and the morphogenetic stage.
During renal organogenesis, BMP7 and BMP2 are expressed in a spatial and temporal pattern consistent with a role in the control of branching morphogenesis (10). BMP7 is highly expressed in the ureteric bud and its branches and also in mesenchymal cells induced by the advancing tips of the ureteric bud and collecting ducts. BMP2 is expressed in an overlapping but distinct spatial domain in metanephric mesenchymal cells adjacent to ureteric bud branches and collecting ducts. We have demonstrated that BMP7 exerts dose-dependent and opposite effects on ureteric bud and collecting duct morphogenesis in embryonic kidney explants treated with BMP7-agarose beads (27). In an in vitro culture model of collecting duct morphogenesis, low doses (<0.5 nM) stimulate tubule number, length, and branching, whereas higher doses generate shorter, unbranched tubules. BMP2 acts in a monophasic dose-dependent manner via Smad1 to generate tubules that are morphologically similar to those generated by high doses of BMP7 (12, 13, 27). Our finding that BMPs exert stimulatory or inhibitory effects during kidney morphogenesis suggests the presence of distinct underlying cellular events and molecular signaling pathways.
The present model of BMP signaling suggests that BMPs modulate gene expression by activating a signaling pathway involving receptor-dependent Smad proteins. Binding of BMPs by type I and type II serine/threonine kinase receptors induces phosphorylation of receptor-activated Smad proteins bound to the cytoplasmic domain of the type I receptor. Phosphorylated Smad dissociates from the receptor and forms a complex with a common Smad binding partner, Smad4. The heteromeric Smad complex then translocates to the nucleus and modifies gene expression [reviewed in 16]. Although this model accounts for the functions of the signaling intermediates described to date, it does not explain the dose-dependent effects of BMPs observed in the kidney.
In this paper, we identify distinct cellular and molecular mechanisms
that underlie the effects of stimulatory and inhibitory BMPs in the
embryonic kidney. We demonstrate that BMP7 controls ureteric bud or
collecting duct cell proliferation in a dose-dependent manner in
embryonic kidney explants and in an vitro culture system for collecting
duct morphogenesis (12, 27). High-dose BMP7 is inhibitory,
whereas low-dose BMP7 is stimulatory. To determine the mechanisms
underlying these effects, we defined signaling events downstream of
BMP7. Our results demonstrate that high-dose BMP7, but not low-dose
BMP7, activates Smad1 in collecting duct cells. To determine the
functional consequences of Smad1 activation, we generated mIMCD-3 cell
lines that stably express a mutant form of Smad1, Smad1(458), shown
here to inhibit signaling by endogenous wild-type Smad1 via a dominant
negative mechanism. Expression of Smad1(
458) partially rescued
inhibition of tubule formation and cell proliferation and stimulation
of apoptosis by high-dose BMP7 but had no significant effect on
stimulation of tubule formation and cell proliferation by low-dose
BMP7. Taken together, our results suggest that BMP7 exerts
dose-dependent effects on ureteric bud or collecting duct cell
proliferation by signaling via Smad1-dependent and -independent pathways.
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MATERIALS AND METHODS |
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Treatment of cultured embryonic kidneys with recombinant BMPs. Mouse embryonic kidneys were surgically resected from E13 pregnant CD1 mice (Charles River), transferred onto 0.45-µm polyethylene terephthalate membranes (Falcon), and cultured in improved modified Eagle medium, Zn2+ option (Richter's modification, BRL Life Technologies) supplemented with 50 µg/ml transferrin (Sigma) (37). AffiGel Blue agarose beads (100-200 mesh, 75-150 µm diameter; Bio-Rad) were incubated in 2 µM BMP2 (provided by Genetics Institute), 2 µM BMP7 (provided by Creative Biomolecules), or 10 mg/ml BSA (BRL Life Technologies) at 37°C for 30 min. Next, beads were washed with PBS, pH 7.4, and then manually placed on the peripheral kidney cortex. Explants were cultured for 48 h in 5% CO2 at 37°C.
In situ assay of cell proliferation in organ explants. 5-Bromo- 2'-deoxyuridine (BrdU; final concentration 10 µM, Roche Molecular Biochemicals) was added to the culture medium of kidney explants 4 h before fixation in 4% formaldehyde/PBS. Detection of BrdU was performed in 5-µm paraffin-embedded sections according to the manufacturer's instructions with some modifications. DNA denaturation was performed by microwave heating tissue sections in 0.01 M citrate buffer, pH 6.0, for 25 min in a microwave pressure cooker (Nordic Ware) at the "HI" setting using a 900-W Dual III Microwave Oven (General Electric). This was followed by two 5-min buffer exchanges at room temperature. Specimens were then preincubated in 1 mM Tris · HCl, pH 7.4, at 37°C for 15 min and treated with 2 µg/ml proteinase K in Tris · HCl for 5 min at 37°C. Endogenous peroxidase activity was quenched in 3% hydrogen peroxide for 5 min. BrdU detection was preceded by a 1-h incubation in 5% goat serum, 0.1% Tween 20 in PBS, pH 7.4 (blocking buffer), at room temperature. Next, sections were incubated with 100 µl of 1.5 U/ml mouse monoclonal anti-BrdU-peroxidase F(ab) fragments (Roche Molecular Biochemicals) for 1 h in a humidified chamber at 37°C. Sections were then washed three times in PBS for 2 min and incubated with 100 µl of either diaminobenzidine (DAB) peroxidase substrate (Pierce) diluted 1:10 in 1× DAB buffer (Pierce) or undiluted aminoethylcarbazole (AEC Histostain-Plus; Zymed) for 5-7 min at room temperature to produce the color reaction. Collecting duct cells were identified by incubating sections with Dolichos biflorus agglutinin (DBA; Vector Laboratories; 1:100) in blocking buffer for 1 h at 37°C. Slides were counterstained with hematoxylin and mounted in either DPX Mountant (DAB-stained, VWR Scientific) or GVA aqueous mounting medium (AEC-stained, Zymed).
In situ assay of apoptosis in organ explants. Apoptotic cells were identified by the terminal deoxynucleotidyl transferase-mediated labeling (TUNEL) assay using the ApopTag kit (Oncor) with some modifications. Sections were pretreated by microwave heating in 0.01 M citrate buffer, pH 6.0, in a microwave pressure cooker at the HI setting for 12 min (just until boiling). This was followed by incubation with 2 µg/ml proteinase K for 5 min at 37°C. After preincubation with 100 µl terminal deoxynucleotidyl transferase (TdT) equilibration buffer (Oncor) for 15 min at 37°C, sections were treated with 100 µl of TdT mixed in reaction buffer (Oncor) according to the manufacturer's instructions for 60 min at 37°C. The reaction was stopped by immersing slides in 300 mM NaCl, 30 mM sodium citrate buffer, pH 8.0 (Stop buffer), for 30 min at 37°C. Endogenous peroxidase activity was quenched in 3% hydrogen peroxide for 5 min, followed by incubation in blocking buffer for 1 h. After excess blocking buffer was rinsed off, sections were treated with 100 µl sheep monoclonal anti-digoxigenin F(ab) fragments antibody (Roche Molecular Biochemicals) diluted 1:20 in blocking buffer for 1 h at 37°C. After sections were washed in PBS, the peroxidase substrate reaction was performed as described for the BrdU incorporation assay. Sections were then counterstained with DBA and hematoxylin as described above and mounted.
Quantitation of collecting duct cell proliferation and apoptosis in organ explants. Tissue sections of E13 organ explants were prepared to include cortical and medullary elements of the ureteric bud and its derivative collecting duct branches. Sections were imaged at ×400 magnification and photographed in their entirety using fine-grain color film (Kodak Royal Gold 400). DBA staining was imaged by fluorescent microscopy with a HBO 50-W mercury-vapor, short-arc lamp using a Shott 38 band-pass filter and a 3-FL fluorescence reflector. A composite of the brightfield and fluorescent images of the entire kidney section was then constructed. We quantified the effects on ureteric bud or collecting duct cell proliferation and apoptosis relative to the proximity to the ligand-coated bead by overlaying a series of concentric rings of increasing diameter (scale 75 µm) over the composite, with the innermost ring positioned over the bead. The number of BrdU- or TUNEL-labeled cells and the number of hematoxylin-stained cells within ureteric bud branches or collecting ducts, identified by DBA, located within each ring zone were counted. The effect of each ligand on collecting duct cell proliferation or apoptosis was calculated from the mean ratio of BrdU- or TUNEL-labeled cells, respectively, to total cells (BrdU- or TUNEL-labeled plus hematoxylin-stained cells) within a ring zone.
Mouse inner medullary collecting duct cell culture. Mouse inner medullary collecting duct (mIMCD-3) cells (American Tissue Culture Collection) grown in monolayers were maintained in DMEM/F-12 Nutrient Mixture (BRL Life Technologies) supplemented with 5% fetal bovine serum (Hyclone), penicillin (100 U/ml), and streptomycin (100 U/ml) in 5% CO2 at 37°C.
Tubulogenesis assay. mIMCD-3 cells were induced to form tubule-like structures in three-dimensional type I collagen gels, as previously described (27). Briefly, mIMCD-3 cells (10,000 cells/gel) were suspended in collagen gels. After 48 h in culture, the gels were fixed in 4% formaldehyde in PBS for 10 min at room temperature, washed four times with PBS, and directly imaged by differential interference contrast (DIC) microscopy using an Axioskop microscope and plan-neofluar objectives (Carl Zeiss). Representative microscopic fields were photographed with a MC80 magnetic shutter camera (Carl Zeiss) using fine-grain black and white film (Ilford XP-2 400) at ×100 and ×400 magnification. The effect of 0.25 nM BMP7, 10-20 nM BMP7, or 5 nM BMP2 was determined by counting the number of continuous, elongated linear structures in four randomly selected photographic fields.
Cell proliferation in mIMCD-3 collagen gel cultures. Cells were labeled with 10 µM BrdU for 4 h at 37°C before fixing. Next, gels were treated in sequence with 100 µl 0.1% trypsin in PBS for 2 h at 37°C, 4N HCl for 2 h to denature DNA, and eight 10-min washes in blocking buffer. Cells were then incubated overnight at 4°C in 100 µl of 1.5 U/ml mouse monoclonal anti-BrdU-peroxidase antibody F(ab) fragments in blocking buffer. Cells were subsequently washed three times in blocking buffer and then incubated with either 0.1 mg/ml goat anti-mouse IgG-AMCA-S secondary antibody (Molecular Probes) for 4 h or AEC peroxidase substrate for 5-10 min. After four 10-min washes in PBS, cells were counterstained with 4 µM ethidium homodimer-1 (Molecular Probes) for 1 h at room temperature.
Apoptosis in mIMCD-3 collagen gel cultures. mIMCD-3 cell apoptosis was determined using the TUNEL assay (ApopTag Kit, Oncor) with some modifications. Cells were incubated in blocking buffer for 30 min, preincubated in equilibration buffer for 1 h, and treated with TdT in reaction buffer for 5 h at 37°C. Stop buffer was then added for 30 min at 37°C. Cells were left overnight in blocking buffer at 4°C and then treated for 4 h with either 100 µl of Texas red-conjugated IgG fraction mouse monoclonal anti-digoxigenin antibody (Jackson ImmunoResearch Laboratories) diluted 1:80 in blocking buffer at room temperature or AEC for 5-10 min. After four 10-min washes in PBS, cells were counterstained with 1 µg/ml bisbenzamide (Hoechst no. 33258; Sigma) for 30 min at 4°C and then washed twice in PBS.
Construction of Smad1 expression vectors.
Plasmids encoding pCMV5b/Flag-Smad1, pCMV5b/Flag-Smad1(458), and
pCMV5b/Flag-Smad1(G419S) were constructed, as described previously
(22, 23) (generously provided by Dr. Jeffrey Wrana). A
Mlu I-BamH I Flag-Smad1 fragment from each vector
was subcloned into pSE280 (Invitrogen). Apa I-Spe
I Flag-Smad1 fragments were then cloned into pTRACER-SV40
(Invitrogen), which contains a zeocin selection cassette as well as a
green fluorescent protein expression cassette.
Stable transfection of mIMCD-3 cell lines
with Flag-Smad1,
Flag-Smad1(458), or
Flag-Smad1(G419S).
mIMCD-3 cells were transfected with 20 µg DNA of either
pTRACER-SV40, pTRACER-SV40/Flag-Smad1,
pTRACER-SV40/Flag-Smad1 (
458), or pTRACER-SV40/ Flag-Smad1(G419S)
using the calcium phosphate-DNA precipitation method. Transfected cells
were selected and maintained in DMEM/F-12 with 5% fetal calf serum
containing 1 mg/ml zeocin. During the selection process, stable
transfectants were identified by noninvasive monitoring of green
fluorescent protein expression under direct fluorescence microscopy.
[32P]phosphate labeling and immunoprecipitation of
wild-type and mutant forms of Smad1.
mIMCD-3 cell lines stably expressing Flag-Smad1 (wild type),
Flag-Smad1(458), and Flag-Smad1(G419S) were labeled with
[32P]PO4 as previously described
(36). Briefly, cells were grown to confluency in six-well
culture plates, washed twice with phosphate-free medium containing
0.2% dialyzed fetal calf serum, and incubated in the same medium for
15 min in 5% CO2 at 37°C. Cells were washed twice with
phosphate-free medium containing 0.2% dialyzed fetal calf serum and
then incubated with media containing 1 mCi/ml
[32P]PO4 for 1 h at 37°C. BMP2 (5 nM)
was then added to the wells, and cells were returned to the incubator
for another hour. Afterward, the
[32P]PO4-containing medium was removed, and
the cells were washed twice with calcium-free, magnesium-free PBS.
Cells were then lysed, and lysates were subjected to
immunoprecipitation with either 1:3,000 anti-Flag M2 antibody followed
by adsorption onto 10 mg/ml protein G-Sepharose (Pharmacia) or 1:3,000
anti-Smad1 antibody followed by adsorption onto 10 mg/ml protein
A-Sepharose (Pharmacia) for 2 h at 4°C. The immunoprecipitates
were washed four times with TNTE containing 0.1% Triton X-100 and two
times with SDS-RIPA (TNTE: 0.5% Triton X-100, 1% deoxycholic acid,
0.1% SDS), separated by SDS-PAGE, and visualized by autoradiography.
In some experiments, analysis of immunoprecipitated proteins was
performed by immunoblotting using a rabbit anti-Smad4 antibody [kindly
provided by Dr. Jeffrey Wrana (23); 1:1,000 dilution]
followed by anti-rabbit HRP (1:10,000) and chemiluminescence.
Statistical analysis. Data were analyzed by using StatView 4.01 (Abacus). Between-ligand comparisons for cell proliferation and apoptosis were determined by ANOVA (P < 0.05). Post hoc comparisons to determine individual differences between ligands were made by Bonferroni-Dunn (P < 0.05).
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RESULTS |
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BMP7 exerts dose-dependent effects on cell
proliferation in ureteric bud derivatives in embryonic kidney explants.
We hypothesized that BMP7 controls cell proliferation and apoptosis
during collecting duct morphogenesis. Because analysis of cellular
events at the level of single cells is largely precluded in whole-mount
explant preparations, we measured cell proliferation by BrdU
incorporation and apoptosis by TUNEL assay in histological sections of
embryonic kidney explants treated with BMP-agarose beads. (Figs.
1 and 2).
The effects of ligand at specific positions relative to the bead were
determined by constructing composite images of explant tissue sections
(Figs. 1 and 2). These composite images were subdivided into concentric
zones relative to the position of the bead. The radial distance between
the perimeter of each zone was 75 µm. The relative concentration of
ligand in each zone could be predicted on the basis of known kinetics
of binding and release of proteins to agarose (31) and the
effect on ureteric bud or collecting duct morphogenesis
(27). Thus zone 1, the area closest to the bead, was
predicted to contain the highest concentration of ligand, whereas zones
incrementally farther from the bead were predicted to contain
incrementally lower concentrations. The effects on cell proliferation
and apoptosis were expressed as the fraction of BrdU- or TUNEL-positive
ureteric bud or collecting duct cells, identified by DBA (Fig.
3).
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BMP7 exerts dose-dependent effects on cell
proliferation and apoptosis during collecting duct morphogenesis in
vitro.
The cellular complexity of the embryonic kidney and the ongoing nature
of mesenchymal-epithelial tissue interactions limit the ability to
discriminate between direct and indirect effects of ligands.
Furthermore, although the concentration-dependent effects of ligands
can be assessed on a relative basis in explant tissue, ligand
concentration cannot be directly measured. In contrast, dose-specific
effects can be measured in the mIMCD-3 model of collecting duct
morphogenesis. mIMCD-3 cells form branched tubules (tubule progenitors
by 48 h) when suspended in extracellular matrix and respond to
growth factors in an identical manner to that observed in embryonic
kidney explants and a ureteric bud cell line (12, 27, 28).
mIMCD-3 cells were induced to form tubular progenitors in the presence
of stimulatory (0.25 nM) or inhibitory (10 nM) doses of BMP7. The
effect of each ligand was determined by observing the number of
tubule-like structures formed and by quantitating the number of
BrdU-labeled or TUNEL-labeled cells compared with the number of
nonlabeled cells identified by bisbenzamide or ethidium homodimer-1
(Figs. 4 and
5). The effect of ligand on cell
proliferation or apoptosis was determined by calculating the mean
fraction of BrdU-labeled or TUNEL-labeled cells within the photographed
fields and expressing this value relative to control (Fig.
6, B and
C).
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Inhibitory doses of BMP7 activate Smad1 in collecting duct cells. The present model of BMP signaling suggests that BMPs modulate gene expression by activating receptor-dependent Smad proteins (16). Recently, we reported that BMP2 inhibits collecting tubule morphogenesis by activating the receptor-dependent Smad, Smad1 (13). Although evidence from cell culture models suggests that both BMP2 and BMP7 signal via Smad1 (23), the doses of BMP7 used in these experiments are those that exert an inhibitory response during renal branching morphogenesis. Because low-dose BMP7 generates a stimulatory response, we hypothesized that inhibitory BMPs (BMP2 and high-dose BMP7) signal via Smad1, whereas low-dose BMP7 signals via a pathway independent of Smad1.
To test our hypothesis, we first determined the effect of low-dose and high-dose BMP7 on the formation of Smad1-Smad4 molecular complexes (Fig. 7). Formation of these complexes is dependent on phosphorylation of Smad1 by activin-like-kinase (ALK) receptors. Monolayer cultures of mIMCD-3 cells were incubated with either low (0.25 nM) or high (10 nM) concentrations of BMP7 for 1 h. Cell lysates were immunoprecipitated with anti-Smad1 antibody and analyzed for Smad1-Smad4 complex formation by immunoblotting with anti-Smad4 antibody. Smad1-Smad4 complexes were detected after treatment with BMP2, as previously reported (13). Similarly, high-dose BMP7 induced formation of Smad1-Smad4 complexes. In contrast, Smad1-Smad4 complexes could not be detected after treatment with low-dose BMP7. These results supported a differential role for Smad1 in signaling via low-dose and high-dose BMP7.
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A mutant Smad1, Smad1 (458), exerts a partial dominant negative
effect on the activation of Smad1 in mIMCD-3 cells.
Our finding that Smad1 is differentially activated by low and high
doses of BMP7 provided a basis for determining the function of Smad1
downstream of BMP7 during collecting tubule formation. Our strategy was
to interrupt Smad1 signaling in mIMCD-3 cells and determine the effects
on BMP7-mediated collecting tubule morphogenesis. The Smad1
mutant, Smad1(
458), lacks the COOH-terminal serines that
are phosphorylated by activated type I BMP receptors. Smad1(
458) binds BMP type I receptor but is not phosphorylated by the type I
receptor. Consequently, Smad1(
458) does not dissociate from the
receptor in the manner observed with wild-type Smad1 (23). We predicted that Flag-Smad1(
458) could act in a dominant negative manner to inhibit binding and subsequent activation of endogenous wild-type Smad1 by its type I receptor in mIMCD-3 cells.
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Smad1 is required for the action of inhibitory
BMPs but not for the action of low-dose
BMP7.
We determined the function of Smad1 during collecting tubule
morphogenesis in mIMCD-3 cells coexpressing Smad1(458). Under control conditions (no ligand added), tubule formation by
mIMCD-3
458 cells was qualitatively
(Fig. 9) and quantitatively (Fig.
10) similar to
tubule formation by mIMCD-3WT and
mIMCD-3G419S cell clones. Treatment with BMP2
inhibited tubulogenesis in mIMCD-3WT and
mIMCD-3G419S cell lines by 96-97%
(P < 0.01, Figs. 9 and 10A). This was
consistent with the inhibitory effect of BMP2 in cells transfected with
empty vector (
100%) in parallel experiments (data not shown). In
contrast, BMP2 inhibited mIMCD-3
458 cell
tubule formation by only 68% (%inhibition,
mIMCD-3
458 vs.
mIMCD-3WT:
68 ± 9 vs.
97 ± 3%,
P = 0.04; Figs. 9 and 10A), suggesting that
Smad1 (
458) exerts a dominant negative effect on signaling via
Smad1 during mIMCD-3 cell morphogenesis. High-dose BMP7 exerted a
similar profile of inhibitory action on tubule formation, decreasing tubule formation by 77 and 67% in the mIMCD-3WT
and mIMCD-3G419S cell lines, respectively, but
only by 50% in mIMCD-3
458 cultures
(%inhibition, mIMCD-3
458 vs.
mIMCD-3WT:
50 ± 6 vs.
77 ± 5%,
P = 0.03; Fig. 10B). In contrast to the effects of Smad1(
458) on signaling via high-dose BMP7, low-dose BMP7
stimulated tubule formation equally well in mIMCD-3 cells stably
expressing wild-type or mutant forms of Smad1 (Figs. 9 and
10C). Taken together, these data suggest that the inhibitory BMPs, BMP2 and high-dose BMP7, are dependent on Smad1, whereas low-dose
BMP7 acts via a Smad1-independent mechanism.
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DISCUSSION |
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The formation of complex tissues during embryogenesis is a tightly regulated process controlled by the simultaneous actions of growth factors that exert stimulatory or inhibitory effects on their cellular targets (5, 24). Development of the mammalian kidney is dependent on the formation of a branched network of collecting ducts, the number and pattern of which must be coordinated with the formation of the more proximal components of the nephron (1). Extensive evidence derived from genetic analysis of tracheal development in Drosophila suggests that growth factors, including the BMP homolog decapentaplegic, control tubular growth and the establishment of boundaries between tubular and nontubular tissues. In the murine kidney, the temporal and spatial expression patterns of members of the BMP family strongly suggest that these growth factors regulate formation of collecting ducts from their progenitor, the ureteric bud (10). Indeed, mutational inactivation of murine BMP7 results in an arrest of branching morphogenesis after initial branching of the ureteric bud (9, 20). Our analysis of the direct actions of BMP7 suggests that it directly controls renal branching morphogenesis in a dose-dependent manner. In low doses (<0.05 nM) BMP7 stimulates tubule number, length, and branching; at higher doses (>0.05 nM) it inhibits these processes (27). However, the mechanisms underlying these differential effects on branching morphogenesis have been previously undefined.
In this paper, we defined the control of renal branching morphogenesis by BMP7 at the level of cellular events and molecules that control these events. We demonstrate that BMP7 exerts differential control over ureteric bud or collecting duct cell proliferation in embryonic kidney explants and in the mIMCD-3 culture model in a manner consistent with its effects on tubular morphogenesis. Low doses of BMP7 are stimulatory, whereas high doses are inhibitory. Furthermore, we demonstrate that the inhibitory BMP, BMP2, also inhibits cell proliferation and, similar to high-dose BMP7, stimulates apoptosis. At a molecular level, our results show that Smad1 functions downstream of inhibitory BMPs (high-dose BMP7 and BMP2) but is not required for the stimulatory effects of low-dose BMP7.
The previous identification of a family of type I and type II cell surface serine/threonine kinase transmembrane receptors that bind to and are activated by BMPs (reviewed in Ref. 16) provides a basis for further defining the dose-dependent actions of BMP7. BMPs signal by activating one or more type I receptors including ALK2, ALK3, and ALK6. An activated type I receptor phosphorylates one or both of the receptor-bound Smad proteins, Smad1 and Smad5. The phosphorylated Smad is then released, binds the common Smad, Smad4, and translocates to the nucleus, where it acts to regulate gene transcription. In cell culture models, BMP7 has been shown to bind and activate the type I receptors, ALK2, ALK3, and ALK6 (35), albeit with different affinities and to induce activation of Smad1 (23) and Smad5 (34). Thus it is possible that low and high doses of BMP7 elicit different cellular responses by activating a different combination of type I receptors and downstream Smads.
Our results provide insight into the mechanisms that control formation of the renal collecting system. During renal branching morphogenesis, the highest rate of cell proliferation is observed in the tips of the ureteric bud branches and collecting ducts, whereas a lower but measurable rate is observed in the cells of the trunks (30, data from the present study). These differential rates of proliferation are thought to determine morphogenetic events, namely, growth and branching at the tips and extension of trunk segments. These events must be tightly regulated because an uncontrolled rate of cell proliferation at ureteric bud branch or collecting duct tips results in disorganized, overgrown tubules (33). It is likely that differential rates of cell proliferation are mediated by growth factors that are expressed in the vicinity of tip and trunk cells. The stimulatory or inhibitory activities of these growth factors must be integrated by ureteric bud or collecting duct cells during formation of the collecting system. Indeed, our recent results suggest how competing signals downstream of hepatocyte growth factor and BMP2 modulate tubular growth and branching (12). Our results reported here suggest that ureteric bud or collecting duct cell proliferation and apoptosis are regulated, as well, by BMP7 in a manner dependent on the delivered dose.
The cellular response to a particular BMP is modulated by the differentiated state of the target cell (2, 3, 6). Thus it is possible that the dose-dependent effects of BMP7 we observed in embryonic kidney explants are due to responses by collecting duct cells with distinct phenotypes. Indeed, cells at the branching tips express different molecular markers of differentiation compared with cells in the trunks (17, 26). Although we have not directly compared the responses of trunk cells and tip cells to BMP7, our results suggest that collecting duct cells of a particular phenotype exhibit dose-dependent effects. In embryonic kidney explants, high-dose BMP7 inhibited cell proliferation in tip cells adjacent to BMP7-agarose beads. However, no effect on apoptosis in ureteric bud-derived cells was observed. In contrast, high-dose BMP7 stimulated apoptosis in mIMCD-3 cells, a differentiated line of medullary collecting duct cells. These observations suggest a need to integrate the dose of delivered ligand with the differentiated state of the target cell in developing a model of BMP7 activity in vivo.
Our finding that BMP7 can control ureteric bud or collecting duct cell proliferation in a dose-dependent manner suggests the presence of a BMP7 activity gradient within the embryonic kidney. Analysis of BMP7 mRNA expression suggests that this gradient may be established by expression of different amounts of BMP7 in distinct spatial domains (10). At the interface of mesenchyme and ureteric bud or collecting duct tips, BMP7 mRNA is expressed by both tissue elements, whereas in the trunk domains of ureteric bud branches and collecting ducts BMP7 is generated only by collecting duct cell types. Thus spatial differences in BMP7 mRNA expression may lead to zones of high and low BMP7 protein expression. Alternatively, as has been observed during decapentaplegic signaling in Drosophila, an activity gradient could be modulated by posttranslational modulation of BMP7 distribution or signaling capability (4, 14, 15, 25). Elucidation of the possible role of these mechanisms in the embryonic kidney will require identification of molecular signaling pathways downstream of BMP7 and interacting extracellular proteins.
Taken together, our results provide a basis for defining the dose-specific effects of BMP7 in vivo in the context of distinct populations of differentiating collecting duct cells and for determining the molecular targets that act downstream of stimulatory concentrations of BMP7. The generation of experimental models in which the expression of BMP7 is manipulated in specified spatial domains of developing ureteric bud branches and collecting ducts in vivo should provide new insights into the functions of BMP7 during branching morphogenesis.
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ACKNOWLEDGEMENTS |
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We thank Dr. Jeffrey Wrana for providing plasmids encoding
Smad1(WT), Smad1(G419S), Smad1(458), Smad5, anti-Smad1, and
anti-Smad4 antibodies and for helpful discussions during these studies.
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FOOTNOTES |
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* T. D. Piscione and T. Phan contributed equally to this work.
BMP2 was provided via a material transfer agreement with Genetics Institute. BMP-7 (osteogenic protein-1) was provided via a material transfer agreement with Creative Biomolecules. This work was supported by grants from the Kidney Foundation of Canada/Canadian Society of Nephrology Fellowship Program and The Hospital for Sick Children Research Training Centre (to T. D. Piscione) and the Medical Research Council of Canada/Canadian Institutes of Health Research (to N. D. Rosenblum).
Address for reprint requests and other correspondence: N. D. Rosenblum, Div. of Nephrology, The Hospital for Sick Children, 555 University Avenue, Toronto, Ontario, Canada M5G 1X8 (E-mail: norman.rosenblum{at}sickkids.on.ca).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 4 January 2000; accepted in final form 30 August 2000.
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