BMP7 controls collecting tubule cell proliferation and apoptosis via Smad1-dependent and -independent pathways

Tino D. Piscione*, Tien Phan*, and Norman D. Rosenblum

Division of Nephrology, Program in Developmental Biology, The Hospital for Sick Children, University of Toronto, Toronto, Ontario, Canada M5G 1X8


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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(Delta 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


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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(Delta 458), shown here to inhibit signaling by endogenous wild-type Smad1 via a dominant negative mechanism. Expression of Smad1(Delta 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.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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(Delta 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(Delta 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 (Delta 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.

To identify stable cell lines expressing either Flag-wild- type (mIMCD-3WT) or Flag-mutant forms of Smad1 (mIMCD-3Delta 458, mIMCD-3G419S), cells were treated with lysis buffer containing 50 mM Tris, pH 7.4, 150 mM NaCl (TNTE), and 0.5% Triton X-100 in the presence of protease inhibitors and then centrifuged to remove debris. Lysates were immunoprecipitated with 1:3,000 anti-Smad1 antibody (generously provided by J. Wrana) for 1 h at 4°C and adsorbed onto protein A-Sepharose beads (Pharmacia). The immunoprecipitates were washed four times in ice-cold TNTE containing 0.1% Triton X-100 and separated by SDS-PAGE. Expression of Flag-Smad1 (wild-type and mutant forms) was confirmed by Western blot analysis using 1:3,000 anti-Flag M2 antibody (Sigma). Stable cell lines expressing Flag-wild-type or Flag-mutant(Delta 458 or G419S) Smad1 were subsequently maintained in DMEM/F-12 supplemented with 5% fetal calf serum and 1 mg/ml zeocin.

[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(Delta 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).


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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).


View larger version (156K):
[in this window]
[in a new window]
 
Fig. 1.   Qualitative effects of bone morphogenetic protein-2 (BMP2) and BMP7 on ureteric bud and collecting duct cell proliferation in mouse embryonic kidney explants. Shown are representative photocomposite images of embryonic day 13 (E13) embryonic kidney explant 5-µm paraffin-embedded tissue sections after treatment for 48 h with agarose beads soaked in either 10 mg/ml BSA (control; A-C), 2 µM BMP2 (D-F), or 2 µM BMP7 (G-I) and placed on the peripheral cortex. A, D, and G: brightfield images at ×100 magnification. C, F, and I: brightfield images at ×200 magnification. B, E, and H: immunofluorescence images of the corresponding sections in A, D, and G. White arrows in C, F, and I : proliferating 5-bromo-2'-deoxyuridine [(BrdU)-positive] cells. Yellow arrowheads in B, E, and H and black arrowheads in A, C, D, F, G, and I : Dolichos biflorus agglutinin (DBA)-stained ureteric bud branches or collecting ducts. Numbered semicircular hatched areas, ring zones. The radial distance between the perimeter of each zone is 75 µm. Treatment with BMP7 decreased the number of ureteric bud branches or collecting ducts and BrdU-positive cells close to the bead but increased the number of branches and BrdU-positive cells in the region distal to the bead. BMP2 decreased the number of ureteric bud or collecting duct branches close to the bead and the number of BrdU-positive cells (black arrow in I).



View larger version (142K):
[in this window]
[in a new window]
 
Fig. 2.   Qualitative effects of BMP2 and BMP7 on ureteric bud and collecting duct cell apoptosis in mouse embryonic kidney explants. Shown are representative photocomposite images of E13 embryonic kidney explant 5-µm paraffin-embedded tissue sections after treatment for 48 h with agarose beads soaked in either 10 mg/ml BSA (control; A-C), 2 µM BMP2 (D-F) or 2 µM BMP7 (G-I) and placed on the peripheral cortex. A, D, and G: brightfield images at ×100 magnification. C, F, and I: brightfield images at ×200 magnification. B, E, and H: immunofluorescence images of the corresponding sections in A, D, and G. White arrows in C, F, and I: apoptotic terminal deoxynucleotidyl transferase-mediated labeling [(TUNEL)-positive] cells. Yellow arrowheads in B, E, and H and black arrowheads in A, C, D, F, G, and I: DBA-stained ureteric bud branches or collecting ducts. Numbered semicircular hatched areas, ring zones. The radial distance between the perimeter of each zone is 75 µm. BMP7 had no apparent effect on the number of TUNEL-positive ureteric bud or collecting duct cells but decreased mesenchymal cell apoptosis. BMP2 decreased the number of ureteric bud or collecting duct branches close to the bead and increased the number of TUNEL-positive cells.



View larger version (23K):
[in this window]
[in a new window]
 
Fig. 3.   Quantitative effects of BMP2 and BMP7 on ureteric bud or collecting duct cell proliferation and apoptosis in embryonic kidney explants. The effect of BSA-agarose (control), BMP7-agarose, or BMP2-agarose on ureteric bud or collecting duct proliferation (BrdU incorporation) or apoptosis (TUNEL staining) was quantitated as a function of the distance of the target cells from the bead. The mean number of ureteric bud or collecting duct cells analyzed in each grid zone is shown above the graphs. A: effect of BMP7 and BMP2 on ureteric bud or collecting duct cell proliferation. BMP7 exerted a biphasic effect on ureteric bud or collecting duct cell proliferation. In zone 1, BMP7 decreased cell proliferation 4-fold compared with control (%BrdU incorporation, BMP7 vs. control: 6.1 ± 1.6 vs. 26.7 ± 3.4, P = 0.004). In contrast, in zone 3, BMP7 increased cell proliferation 2-fold (zone 3, %BrdU incorporation, BMP7 vs. control: 9.6 ± 1.4 vs. 4.3 ± 1.0, P = 0.02); n = 3 experiments. BMP2 decreased cell proliferation in ureteric bud branches or collecting ducts, exerting a 26-fold inhibitory effect in zone 1 (%BrdU incorporation, BMP2 vs. control: 1.0 ± 0.7 vs. 26.7 ± 3.4, P = 0.0003). No significant effect was observed in more distant regions (n = 4 experiments) B: effect of BMP7 and BMP2 on ureteric bud or collecting duct cell apoptosis. BMP7 caused no significant change in ureteric bud or collecting duct cell apoptosis. In contrast, BMP2 stimulated apoptosis 2.3-fold in zone 1 (%TUNEL-positive cells, BMP2 vs. control: 11.7 ± 1.6 vs. 5.0 ± 1.4, P = 0.03) and had no significant effect at greater distances from the bead (n = 4 experiments). *P < 0.05 compared with control.

In parallel experiments, we analyzed the effects of BMP7-agarose on branch formation in DBA-stained whole-mount kidneys. The results were identical to our previously published findings (27) and demonstrated that BMP7-agarose inhibits the number of ureteric bud branches or collecting ducts present within 75 µm of the bead and stimulates branch formation in regions 150-225 µm from the bead (data not shown). This morphogenetic pattern was not observed in control (BSA-agarose) treated kidneys, indicating that these effects are specific to BMP7. BMP7 exerted a biphasic effect on ureteric bud or collecting duct cell proliferation (Fig. 3A). In the region closest to the bead (Fig. 1F), BMP7 decreased cell proliferation fourfold compared with control (%BrdU incorporation, BMP7 vs. control: 6.1 ± 1.6 vs. 26.7 ± 3.4%, P = 0.004). The high rate of cell proliferation observed in controls is consistent with the known high rate of cell division in the branching tips of the ureteric bud and collecting ducts in the peripheral kidney cortex (30) and with our analysis of E13 explants not treated with beads but mapped with zones in a manner to identical to bead-treated kidneys (%ureteric bud or collecting duct cell proliferation, zone 1: 26 ± 8; zone 2: 10 ± 3; zone 3: 9 ± 4; zone 4: 19 ± 4%). In contrast, BMP7 increased cell proliferation twofold in regions 150-225 µm from the bead (zone 3, %BrdU incorporation, BMP7 vs. control: 9.6 ± 1.4 vs. 4.3 ± 1.0%, P = 0.02). No effect of BMP7 was observed in tissue farthest from the bead. This tissue compartment included the main stem ureteric bud, characterized by a moderately high rate of cell proliferation in controls (zone 4, %BrdU incorporation: 12.4 ± 3.7%). BMP7 caused no significant change in ureteric bud or collecting duct cell apoptosis (Figs. 2, D and E, and 3B), which occurred at a low rate in control kidneys (zones 1-3, 5-6%; Figs. 2, A-C, and 3B) consistent with previous observations (18). However, BMP7 did appear to exert a protective effect on mesenchymal cell apoptosis (Fig. 2D), consistent with the massive mesenchymal cell apoptosis present in the BMP7-deficient mouse (9, 20). Taken together, these results suggest that BMP7 exerts dose-dependent control over cell proliferation in the developing collecting system in a manner commensurate with its effects on tubular growth and branching.

Similar to high-dose BMP7, BMP2 decreased cell proliferation in the embryonic renal collecting system, exerting a 26-fold inhibitory effect in the region closest to the bead (zone 1, %BrdU incorporation, BMP2 vs. control: 1.0 ± 0.7 vs. 26.7 ± 3.4%, P = 0.0003). No significant effect was observed in more distant regions (Figs. 1, G-I, and 3A). In addition, BMP2 stimulated apoptosis 2.3-fold in the region limited to within 75 µm of the bead (zone 1, %TUNEL-positive cells, BMP2 vs. control: 11.7 ± 1.6 vs. 5.0 ± 1.4%, P = 0.03) (Figs. 2F and 3B). Thus BMP2 inhibits cell number in the developing collecting system via an inhibitory effect on cell proliferation and a stimulatory effect on apoptosis.

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).


View larger version (131K):
[in this window]
[in a new window]
 
Fig. 4.   Qualitative effects of BMP2 and BMP7 on mIMCD-3 cell proliferation during collecting duct morphogenesis in vitro. mIMCD-3 cells were induced to form tubule progenitors in collagen gels for 48 h in the presence of 5 nM BMP2, 0.25 nM BMP7, or 10 nM BMP7. BrdU was added for the last 4 h of culture. Tubule progenitors were imaged by differential interference contrast (DIC) microscopy (A, D, G, and J). Nuclei were identified by ethidium homodimer-1 and fluorescence microscopy (B, E, H, and K). Proliferating cells (BrdU-positive) were identified with an anti-BrdU antibody and fluorescence microscopy (C, F, I, and L). Black arrowheads in A, D, G, and J: the position of tubule progenitors. White arrowheads in B, E, H, and K: the position of nuclei. White arrows in C, F, I, and L: the position of BrdU-positive cells. Low-dose (0.25 nM) BMP7 increased the number of tubule progenitors formed and BrdU-positive cells. In contrast, high-dose (10 nM) BMP7 and BMP2 inhibited these processes. (Magnification = ×100.)



View larger version (155K):
[in this window]
[in a new window]
 
Fig. 5.   Qualitative effects of BMP2 and BMP7 on mIMCD-3 cell apoptosis during collecting duct morphogenesis in vitro. mIMCD-3 cells were induced to form tubule progenitors in collagen gels for 48 h in the presence of 5 nM BMP2, 0.25 nM BMP7, or 10 nM BMP7. Tubule progenitors were imaged by DIC microscopy (A, D, G, and J). Nuclei were identified by Hoe-33258 and fluorescence microscopy (B, E, H, and K). Apoptotic cells (TUNEL-positive) were identified with an anti-digoxigenin antibody and fluorescence microscopy (C, F, I, and L). Black arrowheads in A, D, G, and J: the position of tubule progenitors. White arrowheads in B, E, H, and K: the position of nuclei. White arrows in C, F, I, and L: the position of TUNEL-positive cells. Low-dose (0.25 nM) BMP7 increased the number of tubule progenitors formed and had little effect on apoptosis. High-dose (10 nM) BMP7 and BMP2 inhibited formation of tubule progenitors and increased apoptosis. (Magnification = ×100.)



View larger version (23K):
[in this window]
[in a new window]
 
Fig. 6.   Quantitative effects of BMP2 and BMP7 on mIMCD-3 cell number, proliferation and apoptosis during collecting duct cell morphogenesis. A: effect of BMP2 and BMP7 on formation of tubule progenitors and mIMCD-3 cell number. Low-dose (0.25 nM) BMP7 increased both tubule number (P = 0.001) and cell number (P = 0.02), with the increase in cell number accounting for the increase in tubules. Both high-dose (10 nM) BMP7 and BMP2 decreased tubule number (P < 0.0001) and cell number (P < 0.01). The decrease in tubule number persisted after controlling for cell number (P < 0.01). B and C: effect of BMP2 and BMP7 on mIMCD-3 cell proliferation and apoptosis, respectively. The effect on cell proliferation or apoptosis was calculated by counting the no. of BrdU-labeled or TUNEL-positive mIMCD-3 cells and dividing by the total no. of cells (no. of ethidium homodimer-1 or Hoe-33258-stained cells) in an imaged field. Data are expressed as %BrdU-positive or TUNEL-positive mIMCD-3 cells/imaged field and are shown as degree of difference (fold difference on the axis) from control. BMP7 (0.25 nM) stimulated mIMCD-3 cell proliferation (P = 0.02) while 10 nM BMP7 and 5 nM BMP2 inhibited (P = 0.02 and P = 0.002, respectively). High-dose (10 nM) BMP7 and BMP2 significantly increased mIMCD-3 cell apoptosis (P = 0.02 and P = 0.01, respectively). In contrast, low-dose BMP7 exerted no significant effect (n = 3 independent experiments). Number of microscopic fields examined: 14 (0.25 nM BMP7), 20 (10 nM BMP7), 30 (5 nM BMP2), 36 (control). dagger , Dagger , *: P < 0.05 compared with controls.

BMP7 exerted dose-dependent stimulatory and inhibitory effects on the formation of tubule progenitors in vitro (Figs. 4, D-I, 5, D-I, and 6A). At a low dose (0.25 nM), BMP7 increased tubule number (no. of tubules/imaged field, low-dose BMP7 vs. control: 48 ± 2 vs. 37 ± 2, P = 0.001), whereas at a high dose (10 nM), BMP7 was inhibitory (no. of tubules/imaged field, high-dose BMP7 vs. control: 21 ± 2 vs. 37 ± 2, P < 0.0001). Consistent with these effects, low-dose BMP7 significantly increased total mIMCD-3 cell number (low-dose BMP7 vs. control: 600 ± 54 vs. 475 ± 24, P = 0.02; Fig. 6A). Analysis of the tubule and nontubule cell populations separately demonstrated a selective stimulatory effect of low-dose BMP7 on tubule cells (Table 1). In contrast, high-dose BMP7 decreased cell number (high-dose BMP7 vs. control: 379 ± 18 vs. 475 ± 24, P = 0.008; Fig. 6A), with a similar effect on tubule and nontubule cells (Table 1). Controlling the effect on tubule number for cell number demonstrated that the stimulatory effect of low-dose BMP7 on tubule number could be totally explained by its effects on cell number. In contrast, inhibition of tubule number by high-dose BMP7 was greater than its effect on cell number, suggesting a second mechanism of action in addition to regulation of cell proliferation (Fig. 6A).

                              
View this table:
[in this window]
[in a new window]
 
Table 1.   Cell proliferation and apoptosis in mIMCD-3 tubules and mIMCD-3 cells not incorporated into tubules

BMP7 controlled mIMCD-3 cell proliferation in a biphasic concentration-dependent manner similar to that observed in tissue explants (Figs. 4, D-F, and 6B). At 0.25 nM, BMP7 stimulated mIMCD-3 cell proliferation (%BrdU incorporation, 0.25 nM BMP7 vs. control: 26 ± 3 vs. 18 ± 2%, P = 0.02). In contrast, 10 nM BMP-7 inhibited mIMCD-3 cell proliferation (10 nM BMP7 vs. control: 14 ± 2 vs. 18 ± 2%, P = 0.05). Furthermore, high-dose (10 nM) BMP-7 significantly increased mIMCD-3 cell apoptosis (Figs. 5, D-F, and 6C, 10 nM BMP7 vs. control: 18.1 ± 1 vs. 12 ± 1, P = 0.02). Yet, low-dose BMP7 exerted no such effect on apoptosis. Taken together, these results indicate that BMP7 exerts a biphasic effect on collecting duct cell number in vitro. At a low dose, BMP7 stimulates via an effect on cell proliferation. At a high dose, BMP7 inhibits via an inhibitory effect on cell proliferation and a stimulatory effect on apoptosis.

BMP2 inhibited formation of tubule progenitors in vitro (Figs. 4, J-L, 5, J-L, and 6A) (no. of tubules/imaged field, BMP2 vs. control: 10 ± 1 vs. 37 ± 2, P < 0.0001). Similar to high-dose BMP7, BMP2 decreased cell number (BMP2 vs. control: 350 ± 16 vs. 475 ± 24, P < 0.0001, Table 1). Control of tubule number for cell number indicated that BMP2-mediated inhibition was only partially explained by a decrease in cell number (Fig. 6A). BMP2 decreased cell proliferation (%BrdU incorporation, BMP2 vs. control: 12.5 ± 1.1 vs. 18.6 ± 1.5%, P = 0.002; Fig. 6B). However, BMP2 exerted its inhibitory effects predominately in the non-tubule cell population (Table 1), consistent with the near-total inhibition of tubule formation in the presence of BMP2. Finally, BMP2 increased apoptosis (%TUNEL-positive cells, BMP2 vs. control: 20 ± 2 vs. 12 ± 1%, P = 0.01; Fig. 6C) with a selective effect on tubule cells (Table 1).

Taken together, these data demonstrate that BMP2 acts in an inhibitory manner to control collecting duct cell number, via a combined effect on cell proliferation and apoptosis.

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.


View larger version (9K):
[in this window]
[in a new window]
 
Fig. 7.   Inhibitory doses of BMP7 activate Smad1 in collecting duct cells. mIMCD-3 cells were incubated in 5% FBS+DMEM/F-12 and then treated for 1 h with either 5 nM BMP2, 0.25 nM BMP7, or 10 nM BMP7. Protein lysates were immunoprecipitated with an antibody to Smad4 (far-right lane) or an antibody to Smad1 and then analyzed by immunoblotting using an antibody to Smad4 (arrow). Smad1-Smad4 complexes were induced by treatment with 5 nM BMP2 and 20 nM BMP7 but not with 0.25 nM BMP7.

A mutant Smad1, Smad1 (Delta 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(Delta 458), lacks the COOH-terminal serines that are phosphorylated by activated type I BMP receptors. Smad1(Delta 458) binds BMP type I receptor but is not phosphorylated by the type I receptor. Consequently, Smad1(Delta 458) does not dissociate from the receptor in the manner observed with wild-type Smad1 (23). We predicted that Flag-Smad1(Delta 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.

To test this prediction, we generated cell lines stably expressing Flag-Smad1(Delta 458). Because genetic overexpression of Smad proteins can activate Smad-dependent signaling in a ligand-independent manner, we generated control cell lines including mIMCD-3 cells stably expressing Flag-tagged wild-type Smad1 or the mutant Smad1, Smad1(G419S). The G419S Smad1 mutation abolishes the ability of Smad1 to bind to its type I receptor. Thus in contrast to Smad1(Delta 458), Smad1(G419S) is predicted not to interfere with signaling via endogenous wild- type Smad1. We determined the effect of Flag-Smad1(Delta 458) on Smad1-dependent signaling by examining discrete molecular events downstream of BMP2/BMP-receptor activated Smad1 signaling, namely, Smad1 phosphorylation and Smad1-Smad4 complex formation (12, 13). First, we determined the effect of Flag-Smad1(Delta 458) expression on Smad1 phosphorylation. mIMCD-3 cell monolayer cultures were labeled with [32P]phosphate and incubated in the absence or presence of 5 nM BMP2. Cell lysates were immunoprecipitated with anti-FLAG antibody and subjected to SDS-PAGE and autoradiography. As shown in Fig. 8A (top), Flag-Smad1 (wild type) was phosphorylated on induction with BMP2. However, neither Flag-Smad1(Delta 458) nor Flag-Smad1(G419S) was phosphorylated above basal levels in BMP2-stimulated cultures despite the presence of equivalent amounts of Flag-Smad1 in each cell line. Next, we determined whether expression of Smad1(Delta 458) prevented BMP2-induced activation of endogenous Smad1 by measuring phosphorylation of both endogenous and Flag-Smad (total Smad1). In these experiments, cell lysates were immunoprecipitated with anti-Smad1 antibody and analyzed by SDS-PAGE and autoradiography (Fig. 8A, bottom). After induction with 5 nM BMP2, mIMCD-3WT and mIMCD-3G419S cells demonstrated a marked increase in total Smad1 phosphorylation. In contrast, mIMCD-3Delta 458 cultures demonstrated only a weak induction of total Smad1 phosphorylation after stimulation with 5 nM BMP2. Differences in the amount of phosphorylated Smad1 in each cell line were not accounted for by variable levels of expression of total Smad1 protein, as demonstrated by immunoblot analysis of the lysates (Fig. 8B, bottom). These data suggest that Smad1(Delta 458) significantly and partially blocks phosphorylation of endogenous Smad1 in mIMCD-3 cells.


View larger version (31K):
[in this window]
[in a new window]
 
Fig. 8.   Smad1(Delta 458) inhibits Smad1-dependent signaling in mIMCD-3 cells. mIMCD-3WT, mIMCD-3G419S , or mIMCD-3Delta 458 cells were incubated in 5% FBS+ DMEM/F-12 and then treated for 1 h with 5 nM BMP-2. A: effect of Smad1(Delta 458) on BMP-2-dependent Smad1 phosphorylation. mIMCD-3 cells were labeled with [32P]phosphate during treatment with BMP-2. Top: after immunoprecipitation (IP) with an antibody to FLAG, proteins were analyzed by SDS-PAGE and autoradiography. BMP-2 induced phosphorylation of FLAG-Smad1 [wild-type (WT)] but not FLAG-Smad1 (Delta 458) or FLAG-Smad1 (G419S) (arrow), despite equivalent expression of these Smad1 proteins. Bottom: after immunoprecipitation with an antibody to Smad1, proteins were analyzed by SDS-PAGE and autoradiography. Smad1 phosphorylation was significantly decreased in mIMCD-3Delta 458 cells but not in mIMCD-3WT or mIMCD-3G419S cells, despite equivalent expression of Smad1 (alpha -Smad1 blot). B: BMP-2-dependent Smad1-Smad4 complex formation by FLAG-tagged Smad1 isoforms. Protein lysates were immunoprecipitated with an antibody to FLAG and immunoblotted with an antibody to Smad4. Smad4 formed a complex with FLAG-Smad1(WT) but not with FLAG-Smad1(Delta 458) or FLAG-Smad1- (G419S) despite the presence of equivalent amounts of FLAG-Smad1 species and Smad4, where WT is wild type. C and D: effect of Smad1(Delta 458) expression on Smad1-Smad4 complex formation. Protein lysates were immunoprecipitated with an antibody to Smad1 and immunoblotted with an antibody to Smad4. The amount of Smad1-Smad4 complex detected was quantified by measuring the intensity of the bands by counting pixels (D). C, i, and D, i: expression of FLAG-Smad1(Delta 458) decreased total Smad1-Smad4 complex formation 2-fold (P < 0.05) compared with expression of FLAG-Smad1(WT) and FLAG-Smad1(G419S), despite equivalent expression of Smad1 (C, ii, and D, ii) and Smad4 (C, iii, and D, iii). *P < 0.05, Smad1(Delta 458) vs. Smad1(WT).

Next, we determined whether partial inhibition of endogenous Smad1 phosphorylation was sufficient to inhibit receptor-regulated Smad1-Smad4 complex formation. As expected, Smad1(Delta 458) and Smad1(G419S) were not able to participate in Smad1-Smad4 complex formation after induction with BMP2 (Fig. 8B), consistent with the prior demonstration that only phosphorylated Smad1 interacts with Smad4 (19). To determine the effect of Smad1(Delta 458) on formation of complexes between total Smad1 (endogenous or Flag-tagged) and Smad4, cell lysates were purified by immunoprecipitation with anti-Smad1 antibody, separated by SDS-PAGE, and immunoblotted with anti-Smad4. The amount of Smad1-Smad4 complex detected was quantified by measuring the intensity of the bands. Expression of Flag-Smad1(Delta 458) decreased total Smad1-Smad4 complex formation twofold (P < 0.05) compared with expression of Flag-Smad1(wild type) and Flag-Smad1(G419S) (Fig. 8C, i). This finding was not accounted for by differences in the total amount of Smad1 or Smad4, as demonstrated by immunoprecipitation and immunoblot analysis for total Smad1 and Smad4 protein, respectively (Fig. 8C, ii and iii). Taken together, these data provided additional evidence that Flag-Smad1(Delta 458) exerted partial dominant negative effects on receptor-regulated endogenous Smad1 signaling in mIMCD-3 cells.

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(Delta 458). Under control conditions (no ligand added), tubule formation by mIMCD-3Delta 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-3Delta 458 cell tubule formation by only 68% (%inhibition, mIMCD-3Delta 458 vs. mIMCD-3WT: -68 ± 9 vs. -97 ± 3%, P = 0.04; Figs. 9 and 10A), suggesting that Smad1 (Delta 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-3Delta 458 cultures (%inhibition, mIMCD-3Delta 458 vs. mIMCD-3WT: -50 ± 6 vs. -77 ± 5%, P = 0.03; Fig. 10B). In contrast to the effects of Smad1(Delta 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.


View larger version (113K):
[in this window]
[in a new window]
 
Fig. 9.   Effect of Smad1(Delta 458) expression on mIMCD-3 morphogenesis. mIMCD-3WT, mIMCD-3Delta 458, and mIMCD-3G419S cells were cultured in type I collagen in culture medium supplemented with 5 nM BMP-2, 20 nM BMP7, or 0.25 nM BMP7 for 48 h and then imaged by DIC microscopy (×100 magnification). Under control conditions (no ligand added), all 3 cell lines were capable of generating tubule-like structures. Treatment with BMP2 inhibited tubule formation by mIMCD-3WT and mIMCD-3G419S cells but only partially by mIMCD-3Delta 458 cells. Similarly, expression of Smad1(Delta 458) partially rescued mIMCD-3 cells (mIMCD-3Delta 458 cell line) from the inhibitory effect of high-dose (20 nM) BMP7. In contrast, low-dose (0.25 nM) BMP7 stimulated tubule morphogenesis in mIMCD-3Delta 458 cells as well as in mIMCD-3WT and mIMCD-3G419S cells. (Magnification = ×100.)



View larger version (24K):
[in this window]
[in a new window]
 
Fig. 10.   Quantitative analysis of effect of Smad1(Delta 458) expression on mIMCD-3 morphogenesis. mIMCD-3WT, mIMCD-3Delta 458, and mIMCD-3G419S cells were cultured in type I collagen in culture medium supplemented with 5 nM BMP-2, 20 nM BMP7, or 0.25 nM BMP7 for 48 h. The number of tubule-like structures was counted in four randomly selected microscopic fields, imaged by DIC microscopy (×100 magnification). The response to ligand is expressed as the percentage of the number of tubules formed divided by the number formed in the absence of ligand. A: effect of Smad1(Delta 458) expression on BMP2-mediated inhibition. BMP2 (5 nM) significantly inhibited mIMCD-3 tubulogenesis in all cell lines (%inhibition, mIMCD-3WT: 97 ± 3%, P = 0.001; mIMCD-3G419S: 96 ± 4%, P = 0.002; mIMCD-3Delta 458: 68 ± 9%, P = 0.02). However, the percent inhibition of tubule formation in mIMCD-3Delta 458 cultures was significantly lower compared with the inhibition of tubulogenesis in other cell lines (mIMCD-3Delta 458 vs. mIMCD-3WT, P = 0.04; vs. mIMCD-3G419S, P = 0.05). B: effect of Smad1(Delta 458) expression on BMP7-mediated inhibition. BMP7 (20 nM) significantly inhibited mIMCD-3 tubule formation in all cell lines (%inhibition, mIMCD-3WT: 77 ± 5%, P = 0.003; mIMCD-3Delta 458: 50 ± 6%, P = 0.02; and mIMCD-3G419S: 67 ± 13%, P = 0.04). However, the amount of inhibition in the mIMCD-3Delta 458 cultures was significantly less than that observed in mIMCD-3WT cultures (P = 0.03). C: effect of Smad1(Delta 458) expression on BMP7-mediated stimulation. Expression of Smad1(Delta 458) did not alter the stimulatory effect of low-dose (0.25 nM) BMP7 on tubule formation (%stimulation, mIMCD-3Delta 458: 44 ± 7%, vs. mIMCD-3WT: 38 ± 11, P = 0.64; vs. mIMCD-3G419S: 38 ± 6, P = 0.52). dagger , *: P < 0.05 compared with control.

Having demonstrated a function for Smad1 downstream of BMP2 and high-dose BMP7 at the morphogenetic level, we determined the role of Smad1 in BMP-dependent collecting duct cell proliferation and apoptosis. mIMCD-3Delta 458 or mIMCD-3WT cells were cultured in collagen gels in the presence of BMP2 or BMP7. Cell proliferation and apoptosis were measured by BrdU incorporation and TUNEL assays, respectively. For each cell line, the effect of each ligand relative to untreated cells (controls) was calculated as the %BrdU-positive or %TUNEL-positive cells relative to its control and expressed as the degree of difference from control (Fig. 11). BMP2 significantly inhibited cell proliferation in mIMCD-3WT cells (degree of difference from control: -0.67 ± 0.01, P = 0.006) (Fig. 11A). The degree of inhibition was greatly reduced in mIMCD-3Delta 458 cells (degree of difference from control: -0.07 ± 0.15) and was significantly different from that observed in mIMCD-3WT cells (P = 0.05). Similarly, high-dose BMP7 significantly inhibited cell proliferation in mIMCD-3WT cells (degree of difference from control: -0.53 ± 0.1, P = 0.0007) but exerted no significant effect in mIMCD-3Delta 458 cells. In contrast, low-dose BMP7 stimulated BrdU incorporation to a similar degree (~20%) in mIMCD-3WT and mIMCD-3Delta 458 cells. Stable expression of Smad1(Delta 458) abrogated the stimulatory effect of BMP2 on apoptosis (degree of difference in %TUNEL-positive cells, mIMCD-3WT vs. mIMCD-3Delta 458 cells: 1.47 ± 0.3 vs. 0.5 ± 0.2, P = 0.001). A similar effect was observed for high-dose BMP7 (degree of difference in %TUNEL-positive cells, mIMCD-3WT vs. mIMCD-3Delta 458 cells: 1.31 ± 0.3 vs. -0.2 ± 0.1, P = 0.004). In contrast, low-dose BMP7 exerted no significant effect on apoptosis in either cell line. Taken together, these results strongly suggest that Smad1 functions downstream of inhibitory BMPs, that is, high-dose BMP7 and BMP2, but is not required for the actions of the stimulatory BMP, low-dose BMP7.


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 11.   Effect of Smad1(Delta 458) expression on BMP-dependent control of mIMCD-3 cell proliferation and apoptosis. mIMCD-3Delta 458 (stippled bars) or mIMCD-3WT (solid bars) cells were cultured in collagen gels in the presence of 5 nM BMP2 or 20 nM BMP7. Cell proliferation and apoptosis were measured by BrdU incorporation and TUNEL assays (A and B, respectively), and the number of proliferating and apoptotic cells was determined as a percentage of all cells. Data are expressed as degree of (fold) difference from control (no ligand). A: expression of Smad1(Delta 458) significantly rescued inhibition of cell proliferation by BMP2 (degree of inhibition in mIMCD-3WT vs. mIMCD-3Delta 458 cells: -0.67 ± 0.01 vs. -0.07 ± 0.15, P = 0.05) and by high-dose (20 nM) BMP7 (degree of inhibition in mIMCD-3WT vs. mIMCD-3Delta 458 cells:-0.53 ± 0.1 vs. -0.28 ± 0.09, P = 0.03). Low-dose BMP7 stimulated cell proliferation in both mIMCD-3WT vs. mIMCD-3Delta 458 cells: [degree of stimulation from control: mIMCD-3WT: 0.21 ± 0.04 (P < 0.05); mIMCD-3Delta 458: 0.19 ± 0.05, P < 0.05]. B: expression of Smad1(Delta 458) significantly reduced BMP2-stimulated apoptosis (degree of inhibition in mIMCD-3WT vs. mIMCD-3Delta 458 cells: 1.47 ± 0.3 vs. 0.5 ± 0.2, P = 0.001), and high-dose BMP7 stimulated apoptosis (degree of inhibition in mIMCD-3WT vs. mIMCD-3Delta 458 cells: 1.31 ± 0.3 vs. -0.2 ± 0.1, P = 0.004). Low-dose BMP7 exerted no significant effect in either cell line (degree of stimulation from control: mIMCD-3WT: 0.16 ± 0.04; mIMCD-3Delta 458: 0.18 ± 0.05). *P < 0.05 compared with control.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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.


    ACKNOWLEDGEMENTS

We thank Dr. Jeffrey Wrana for providing plasmids encoding Smad1(WT), Smad1(G419S), Smad1(Delta 458), Smad5, anti-Smad1, and anti-Smad4 antibodies and for helpful discussions during these studies.


    FOOTNOTES

* 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.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Al-Awqati, Q, and Goldberg MR. Architectural patterns in branching morphogenesis in the kidney. Kidney Int 54: 1832-1842, 1998[ISI][Medline].

2.   Andrews, PW, Damjonov I, Berends J, Kumpf S, Zappavigna V, Mavillo F, and Sampath K. Inhibition of proliferation and induction of differentiation of pluripotent human embryonal carcinoma cells by osteogenic protein-1 (or bone morphogenetic protein-7). Lab Invest 71: 243-251, 1994[ISI][Medline].

3.   Arkell, R, and Beddington RSP BMP-7 influences pattern and growth of the developing hindbrain of mouse embryos. Development 124: 1-12, 1997[Abstract/Free Full Text].

4.   Ashe, HL, and Levine M. Local inhibition and long-range enhancement of Dpp signal transduction by Sog. Nature 398: 427-431, 1999[ISI][Medline].

5.   Buckland, RA, Collinson JM, Graham E, Davidson DR, and Hill RE. Antagonistic effects of FGF4 on BMP induction of apoptosis and chondrogenesis in the chick limb bud. Mech Dev 71: 143-150, 1998[ISI][Medline].

6.   Chen, P, Vukicevic S, Sampath TK, and Luyten FP. Osteogenic protein-1 promotes growth and maturation of chick sternal chondrocytes in serum-free medium. J Cell Sci 108: 105-114, 1995[Abstract/Free Full Text].

7.   Coles, HSR, Burne JF, and Raff MC. Large-scale normal cell death in the developing rat kidney and its reduction by epidermal growth factor. Development 117: 777-784, 1993.

8.   D'Alessandro, J, and Wang EA. Bone morphogenetic proteins inhibit proliferation, induce reversible differentiation and prevent cell death in astrocyte lineage cells. Growth Factors 11: 45-52, 1994[ISI][Medline].

9.   Dudley, AT, Lyons KM, and Robertson EJ. A requirement for bone morphogenetic protein-7 during development of the mammalian kidney and eye. Genes Dev 9: 2795-2807, 1995[Abstract].

10.   Dudley, AT, and Robertson EJ. Overlapping expression domains of bone morphogenetic protein family members potentially account for limited tissue defects in BMP7 deficient embryos. Dev Dyn 208: 349-362, 1997[ISI][Medline].

11.   Graham, A, Francis-West F, Brickell P, and Lumsden A. The signalling molecule BMP4 mediates apoptosis in the rhombencephalic neural crest. Nature 372: 684-686, 1994[ISI][Medline].

12.   Gupta, IR, Macias-Silva M, Kim S, Zhou X, Piscione TD, Whiteside C, Wrana JL, and Rosenblum ND. HGF rescues BMP-2-mediated inhibition of renal collecting duct morphogenesis without interrupting Smad1 dependent signaling. J Cell Sci 113: 269-278, 2000[Abstract/Free Full Text].

13.   Gupta, IR, Piscione TD, Grisaru S, Phan T, Macias-Silva M, Zhou X, Whiteside C, Wrana JL, and Rosenblum ND. Protein kinase A is a negative regulator of renal branching morphogenesis and modulates inhibitory and stimulatory bone morphogenetic proteins. J Biol Chem 274: 26305-26314, 1999[Abstract/Free Full Text].

14.   Haerry, TE, Khalsa O, O'Connor MB, and Wharton KA. Synergistic signaling by two BMP ligands through the SAX and TKV receptors controls wing growth and patterning in Drosophila. Development 125: 3977-3987, 1998[Abstract/Free Full Text].

15.   Jazwinska, A, Kirov N, Wieschaus E, Roth S, and Rushlow C. The Drosophila gene brinker reveals a novel mechanism of Dpp target gene regulation. Cell 96: 563-573, 1999[ISI][Medline].

16.   Kawabata, M, Imamura T, and Miyazono K. Signal transduction by bone morphogenetic proteins. Cytokine Growth Factor Rev 9: 49-61, 1998[ISI][Medline].

17.   Kispert, A, Vainio S, Shen L, Rowitch DH, and McMahon AP. Proteoglycans are required for maintenance of Wnt-11 expression in the ureter tips. Development 122: 3627-3637, 1996[Abstract/Free Full Text].

18.   Koseki, C, Herzlinger D, and Al-Awqati Q. Apoptosis in metanephric development. J Cell Biol 119: 1327-1333, 1992[Abstract].

19.   Lagna, G, Hata A, Hemmati-Brivanlou A, and Massague J. Partnership between DPC4 and SMAD proteins in TGF-beta signalling pathways. Nature 383: 832-836, 1996[ISI][Medline].

20.   Luo, G, Hofmann C, Bronckers AL, Sohocki M, Bradley A, and Karsenty G. BMP-7 is an inducer of nephrogenesis, and is also required for eye development and skeletal patterning. Genes Dev 9: 2808-2820, 1995[Abstract].

21.   Mabie, PC, Mehler MF, and Kessler JA. Multiple roles of bone morphogenetic protein signaling in the regulation of cortical cell number and phenotype. J Neurosci 19: 7077-7088, 1999[Abstract/Free Full Text].

22.   Macias-Silva, M, Abdollah S, Hoodless PA, Pirone R, Attisano L, and Wrana JL. MADR2 is a substrate of the TGF-beta receptor and its phoshorylation is required for nuclear accumulation and signalling. Cell 87: 1215-1224, 1996[ISI][Medline].

23.   Macias-Silva, M, Hoodless PA, Tang SJ, Buchwald M, and Wrana JL. Specific activation of Smad1 signaling pathways by the BMP7 type I receptor, ALK2. J Biol Chem 273: 25628-25636, 1998[Abstract/Free Full Text].

24.   Neubüser, A, Peters H, Balling R, and Martin GR. Antagonistic interactions between FGF and BMP signaling pathways: a mechanism for positioning the sites of tooth formation. Cell 90: 247-255, 1997[ISI][Medline].

25.   Nguyen, M, Park S, Marques G, and Arora K. Interpretation of a BMP activity gradient in Drosophila embryos depends on synergistic signaling by two type I receptors, SAX and TKV. Cell 95: 495-506, 1998[ISI][Medline].

26.   Pachnis, V, Mankoo B, and Costantini F. Expression of the c-ret proto-oncogene during mouse embryogenesis. Development 119: 1005-1017, 1993[Abstract/Free Full Text].

27.   Piscione, TD, Yager TD, Gupta IR, Grinfeld B, Pei Y, Attisano L, Wrana JL, and Rosenblum ND. BMP-2 and OP-1 exert direct and opposite effects on renal branching morphogenesis. Am J Physiol Renal Physiol 273: F961-F975, 1997[Abstract/Free Full Text].

28.   Sakurai, H, Barros EJ, Tsukamoto T, Barasch J, and Nigam SK. An in vitro tubulogenesis system using cell lines derived from the embryonic kidney shows dependence on multiple soluble growth factors. Proc Natl Acad Sci USA 94: 6279-6284, 1997[Abstract/Free Full Text].

29.   Savill, J. Apoptosis and the kidney. J Am Soc Nephrol 5: 12-21, 1994[Abstract].

30.   Saxen, L. Organogenesis of the Kidney. Cambridge, UK: Cambridge Univ. Press, 1987.

31.   Schreiber, AB, Winkler ME, and Derynk R. Transforming growth factor-alpha : a more potent angiogenic mediator than epidermal growth factor. Science 232: 1250-1253, 1986[ISI][Medline].

32.   Song, Q, Mehler MF, and Kessler JA. Bone morphogenetic proteins induce apoptosis and growth factor dependence of cultured sympathoadrenal progenitor cells. Dev Biol 196: 119-127, 1998[ISI][Medline].

33.   Srinivas, S, Wu Z, Chen CM, D'Agati V, and Costantini F. Dominant effects of RET receptor misexpression and ligand-independent RET signaling on ureteric bud development. Development 126: 1375-1386, 1999[Abstract/Free Full Text].

34.   Tamaki, K, Souchelnytskyi S, Itoh S, Nakao A, Sampath K, Heldin CH, and Dijke PT. Intracellular signaling of osteogenic protein-1 through Smad5 activation. J Cell Physiol 177: 355-363, 1998[ISI][Medline].

35.   Ten Dijke, P, Yamashita H, Sampath TK, Reddi AH, Estevez M, Riddle DL, Ichijo H, Heldin CH, and Miyazono K. Identification of type I receptors for osteogenic protein-1 and bone morphogenetic protein-4. J Biol Chem 269: 16985-16988, 1994[Abstract/Free Full Text].

36.   Wrana, JL, Attisano L, Wieser R, Ventura F, and Massague J. Mechanism of activation of the TGF-beta receptor. Nature 370: 341-347, 1994[ISI][Medline].

37.   Yeger, H, Forget D, Alami J, and Williams BR. Analysis of WT1 gene expression during mouse nephrogenesis in organ culture. In Vitro Cell Dev Biol 32: 496-504, 1996[ISI].


Am J Physiol Renal Fluid Electrolyte Physiol 280(1):F19-F33
0363-6127/01 $5.00 Copyright © 2001 the American Physiological Society