BMP-2 and OP-1 exert direct and opposite effects on renal
branching morphogenesis
Tino D.
Piscione1,
Thomas D.
Yager1,2,
Indra R.
Gupta1,
Branko
Grinfeld1,
York
Pei3,
Liliana
Attisano4,
Jeffrey L.
Wrana2,5, and
Norman D.
Rosenblum1,2
5 Division of Gastroenterology
and Nutrition and 2 Program in
Developmental Biology,
1 Division of Nephrology, Hospital
for Sick Children, 3 Division of
Nephrology, Toronto Hospital, and
4 Department of Anatomy and Cell
Biology, University of Toronto, Toronto, Ontario, Canada M5G
1X8
 |
ABSTRACT |
The bone
morphogenetic proteins, BMP-2 and OP-1, are candidates for growth
factors that control renal branching morphogenesis. We examined their
effects in embryonic kidney explants and in the mIMCD-3 cell model of
collecting duct morphogenesis (mIMCD-3 cells are derived from the
terminal inner medullary collecting duct of the SV40 mouse). Osteogenic
protein-1 (OP-1), at a dose of 0.25 nM, increased explant growth by
30% (P = 0.001). In contrast, 100-fold greater concentrations of OP-1 (28 nM) decreased explant growth by 10% (P < 0.001). BMP-2
was entirely inhibitory (maximum inhibition of 7% at 5 nM,
P < 0.0004). In an in vitro model
for branching morphogenesis utilizing the kidney epithelial cell line, mIMCD-3, low doses of OP-1 (<0.5 nM) increased the number of tubular structures formed by 28 ± 5% (P = 0.01), whereas concentrations >0.5 nM decreased that number by 22 ± 8% (P = 0.02). All
concentrations of BMP-2 (0.05-10 nM) were inhibitory (maximum
inhibition at 10 nM of 88 ± 3%, P < 0.0001). Stimulatory doses of OP-1 increased tubular length
(P = 0.003) and the number of branch
points/structure (3.2-fold increase, P = 0.0005) compared with BMP-2. To determine the molecular basis for
these effects, we demonstrated that BMP-2 is bound to mIMCD-3 cells by
the type I serine/threonine kinase receptor, ALK-3, and that OP-1 bound
to an ~80-kDa protein using ligand-receptor affinity assays. To
demonstrate that OP-1 can exert both stimulatory and inhibitory effects
within a developing kidney, embryonic explants were treated with
agarose beads saturated with 2 µM OP-1. OP-1 decreased
the number of ureteric bud/collecting duct branches adjacent to
the beads by 58 ± 1% (P < 0.0001). In contrast, the number of branches in tissue distal to the
OP-1 beads was enhanced, suggesting a stimulatory effect at lower doses of OP-1. We conclude that OP-1 and BMP-2 directly control branching morphogenesis and that the effects of OP-1 are dependent on its local
concentration within developing kidney tissue.
bone morphogenetic protein-2; osteogenic protein-1; bone
morphogenetic protein-7; inner medullary collecting duct; tubulogenesis
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INTRODUCTION |
RENAL BRANCHING morphogenesis is defined as the growth
and branching of epithelial tubules during embryonic development (30). In the kidney, branching morphogenesis results from reciprocal mesenchymal-epithelial tissue interactions between the mesenchymal metanephric blastema and the epithelial ureteric bud (and its derivative collecting ducts). These interactions are mediated, in part,
by peptide growth factors. Recent evidence suggests that bone
morphogenetic proteins (BMPs), a family of growth factors within the
transforming growth factor-
(TGF-
) superfamily, play a role in
controlling these tissue interactions (7, 8, 19, 34).
BMPs constitute the largest subgroup within the TGF-
superfamily.
They are related to other TGF-
- related families including the
activins and TGF-
s by the homology among their carboxy-terminal domains (reviewed in Ref. 14). The BMPs are subdivided into three
subgroups: 1) BMP-2/4,
2) BMP-5, BMP-6, and BMP-7 (also named osteogenic protein-1, OP-1), and
3) BMP-3 and BMP-8 (16). BMPs induce
cellular responses by forming heteromeric complexes with type I and
type II cell surface transmembrane serine/threonine kinase receptors
(28). The cellular response to a BMP is defined by the particular
member of the type I receptor family to which it binds, since the type
I receptor transduces a signal to downstream intracellular target
molecules (37). Binding assays in model cell systems suggest that ALK-3
and ALK-6 are candidate type I receptors for BMP-2 (18) and that ALK-2
is a candidate receptor for OP-1 (33).
A large body of evidence in multiple species indicates that BMPs are
involved in controlling morphogenetic steps at multiple stages of
development (14). Several types of evidence suggest a role for BMPs
during mammalian kidney development. BMP-2/4 and OP-1 and ALK-3 and
ALK-6 are expressed in a temporal and spatial pattern that is
consistent with a role in inductive mesenchymal-epithelial tissue
interactions (2, 6, 7, 24, 35). OP-1 induces the differentiation of
epithelial elements when added to explanted uninduced metanephric
blastema (34). Mutational inactivation of the murine Op-1
gene results in underdevelopment and disorganization of
both the mesenchymal- and epithelial-derived elements in the embryonic
kidney. This phenotype suggests that OP-1 may function to control
aspects of renal development other than inductive tissue interactions
(7, 19). However, the cellular complexity of the developing kidney and
the ongoing nature of reciprocal mesenchymal-epithelial interactions
severely limit the ability to distinguish the effects of OP-1 at the
primary versus secondary level. Therefore, the Op-1
/
renal phenotype does not provide direct evidence regarding the role of OP-1 in branching morphogenesis. The function of BMP-2 and
its candidate ALK receptors is also undefined. Mutational inactivation
of BMP-2 and its type I serine/threonine kinase receptor, ALK-3,
results in embryonic lethality before the onset of organogenesis (14,
21).
In this report, we determined the direct effects of BMP-2 and OP-1 on
renal branching morphogenesis. Our initial experiments in embryonic
kidney explants yielded the surprising result that low doses of OP-1
(0.25 nM) stimulated explant growth, specifically the ureteric
bud/collecting ducts, whereas higher doses (28 nM) were inhibitory.
BMP-2 only inhibited explant growth. We characterized the direct
effects of these BMPs on developing collecting ducts in the mIMCD-3
model (4). Consistent with our results in explants, OP-1 (<0.25 nM)
stimulated the number of tubules formed, the number of branch
points/tubule and tubular length. In contrast, concentrations of OP-1
>0.5 nM and 0.05-10 nM BMP-2 were inhibitory. To determine the
molecular basis for these opposite effects, we identified candidate
cell surface receptors for BMP-2 and OP-1. Our results demonstrate that
both BMP-2 and OP-1 are bound to mIMCD-3 cells by the type II
serine/threonine kinase receptor, ActRII/IIB. BMP-2 is also bound by
the type I serine/threonine kinase receptor, ALK-3, and OP-1 is bound
by an ~80-kDa protein. Since peptide growth factors exist in a
concentration gradient in some developing tissues (11), we tested the
possibility that OP-1 exerts a stimulatory or inhibitory effect on
branching morphogenesis in embryonic kidney tissue depending on its
local concentration. Using agarose beads saturated with micromolar
amounts of OP-1, we demonstrated an inhibition of ureteric
bud/collecting duct branching adjacent to the beads but stimulation in
tissue distal to the beads. Taken together, our results suggest that
OP-1 and BMP-2 directly control renal branching morphogenesis and that
the effects of OP-1 are dependent on its local concentrations within
developing kidney tissue.
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METHODS |
Embryonic kidney organ culture and treatment of organs
with recombinant proteins. Mouse embryos were
surgically resected from embryonic day 12 (E12)
pregnant CD1 mice. Embryonic kidneys were isolated by
microdissection and were cultured on 0.45 µm polyethylene terephthalate membranes (Falcon) (40) in 12-well multiwell plates in
the presence of Richter's modification of Dulbecco's modified Eagle's medium-Ham's F-12 nutrient mixture (DMEM-F12, BRL Life Technologies) containing 50 µg/ml transferrin (Sigma) (25). Culture
medium was changed every 48 h. BMP-2 (provided by Genetech) and OP-1
(provided by Creative Biomolecules) were either added directly to the
culture medium or were absorbed at 37°C for 30 min onto Affi-Gel
blue agarose beads (100-200 mesh, 75-150 µm diameter;
Bio-Rad), previously washed once with phosphate-buffered saline, pH 7.4 (PBS). Treated beads were washed once in Richter's modified DMEM-F12
medium and manually placed on organ cultures (31). The effect on kidney
growth was defined by measuring the surface area of the explants using
image analysis software (NIH Image) and by imaging 5-µm
paraffin-embedded tissue sections stained with hematoxylin and eosin.
Whole mount immunostaining of embryonic
kidneys. Embryonic kidneys were fixed in 4%
formaldehyde in PBS for 10 min, washed with PBS four times for 10 min
each wash, and then stored in blocking buffer consisting of 1% goat
serum and 0.0001% Tween-20 in PBS. Kidneys were then incubated for 2 h
with fluorescein-conjugated Dolichos
biflorus agglutinin (20 µg/ml) (Vector Labs) in
blocking buffer and then washed with PBS four times for 10 min each
wash. The effect of ligand on the collecting system was defined by
counting the number of ureteric bud/collecting duct branches formed on either side of the ureteric bud.
mIMCD cell culture. The mIMCD-3 cell line is derived from
the terminal inner renal medullary collecting duct of the SV40
transgenic mouse. mIMCD-3 cells retain several differentiated
characteristics of this nephron segment, as previously described (26).
Monolayer cultures of mIMCD-3 cells (obtained from American Tissue
Culture Collection) were maintained in DMEM-F12 supplemented with 5%
fetal bovine serum (Hyclone), penicillin (100 U/ml), and streptomycin (100 U/ml) in 5% CO2 at 37°C.
For assays of tubulogenesis, collagen gels were prepared on ice by
mixing 4 µl of 1 M
N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (Sigma), 8 µl of 1 M
NaHCO3, 40 µl DMEM-F12, 200 µl
rat type I collagen (Collaborative Biomedical Products), and
25,000-250,000 mIMCD-3 cells. Aliquots (50 µl/well) were seeded
in 96-well culture plates. After the gels solidified at 37°C, 100 µl of DMEM-F12 containing 5% fetal bovine serum were added to each
well. Cultures were then maintained at 37°C in 5%
CO2 and the medium was changed every 48 h.
Effect of growth factors on mIMCD-3
tubulogenesis. Serial dilutions of activin A and
TGF-
1 (kindly provided by Genetech), OP-1, and BMP-2 were prepared
in DMEM-F12 and added to newly established cultures of mIMCD-3 cells
embedded in collagen gels. After 48 h in culture, gels were fixed in
4% formaldehyde in PBS for 10 min at room temperature. Fixed gels were
then washed four times in PBS and stored in blocking buffer at 4°C.
After fixation, gels were 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 (Kodak T-Max 400 or Ilford
Delta 400) at ×100 magnification. The effect of each ligand on
mIMCD-3 tubulogenesis was determined by counting the number of
continuous, elongated, linear structures per quadrant of photograph.
Morphometric analysis of mIMCD-3
structures. Collagen gels were prepared containing
5,000 cells/gel. Cell culture medium containing ligand was immediately
added to wells containing solidified gels to achieve the following
concentrations: OP-1, 0.25 nM; and BMP-2, 5 nM. Two or seven days
later, gels were fixed and stained with bis-benzamide (Hoechst No. 33258, Sigma) for 30 min at 4°C in the dark and then
destained with PBS for 15 min at room temperature in the dark. In some
experiments, mIMCD-3 nuclei were detected with a monoclonal anti-human
histone-1 antibody (Chemicon) and a rhodamine-labeled mouse anti-human
secondary antibody (BRL-Life Technologies).
Bis-benzamide or antibody-treated gels
were directly imaged by DIC and fluorescence microscopy. Fluorescence
microscopy was performed with a HBO 50-W mercury vapor short-arc lamp
using a Shott-38 band-pass filter and a 3-FL fluorescence reflector. Representative structures imaged by both DIC and fluorescence microscopy were photographed. Morphometric analysis was conducted by
making scaled measurements of the photographed structures for the
length of long axis, number of branch points per structure, length of
branches, and number of Hoechst-stained nuclei per structure and per
branch. A branch was defined by the presence of one or more nuclei
beyond the branch point, whereas a cell process was defined by the
absence of any nuclei in a structure extending beyond a branch point.
Reverse transcription-polymerase chain reaction
cloning of BMPs and receptor serine/threonine kinases.
Poly(A)+ mRNA was isolated from
E13 mouse metanephroi by the oligo(dT) method using a commercial kit (Fast Track, Invitrogen).
Oligo(dT)-primed first-strand cDNA was synthesized using 200 ng
poly(A)+ mRNA as substrate and
reverse transcriptase (Superscript II, GIBCO-BRL).
For reverse transcription-polymerase chain reaction (RT-PCR) cloning of
cDNAs encoding BMPs, E13 metanephric
cDNA was used as a substrate for PCR using degenerate oligonucleotides
directed against conserved sequences present in the subfamily of
TGF-
members that includes the BMPs, Vg-1 and decapentaplegic
(1). A 130-bp DNA band was generated and cloned into
pBluescript. Recombinant plasmids were purified and sequenced by the
dideoxy chain-termination method using a commercial kit (Pharmacia).
For RT-PCR cloning of cDNAs encoding receptor serine/threonine kinases,
first-strand E13 metanephric cDNA was
used as substrate for PCR using degenerate oligonucleotides directed
against conserved sequences present in the family of receptor
serine/threonine kinases (9). A 450-bp PCR product was generated and
cloned into pBluescript. Recombinant plasmids were isolated and
sequenced.
Detection of mIMCD-3 serine/threonine kinase receptor
RNAs. Ribonuclease (RNase) protection assays were
performed using specific antisense riboprobes for T
RI (ALK-5),
T
RII, ActRIB (ALK-4), ActRII, and c-met. Riboprobes were prepared
from linearized plasmid templates treated with proteinase K (50 µg/ml
at 37°C for 30 min). Riboprobes were then synthesized by performing
in vitro transcription with T3 polymerase in the presence of
[32P]UTP
(Amersham). Following digestion of template DNA with RNase-free deoxyribonuclease I, full-length radiolabeled transcripts were isolated
by gel purification. RNase protection assays were performed using a
commercial kit (Ambion). A quantity of 10 µg of mIMCD-3 total RNA was
coprecipitated with 5 × 105
cpm of riboprobe, resuspended in 20 µl of hybridization buffer (80%
formamide, 100 mM sodium citrate, 300 mM sodium acetate, and 1 mM EDTA)
and hybridized overnight at 50°C. For a positive control, 5 µg of
mouse liver total RNA was hybridized with 5 × 104 cpm of an antisense mouse
-actin riboprobe. For a negative control, 10 µg of yeast RNA was
hybridized with each riboprobe. Following hybridization, RNase
digestion was carried out at 37°C for 30 min. The precipitated RNA
was dissolved in 8 µl of 80% loading buffer, heat-denatured at
90°C for 3 min, and electrophoresed in a 8 M urea/6% acrylamide
gel.
RT-PCR was used to detect mIMCD-3 mRNAs encoding the ALK-2, ALK-3, and
ALK-6 receptors. First-strand cDNA was synthesized as described above.
PCR reactions were then performed using the following primers, all of
which encode sequences in the extracellular domains of specific ALK
receptors: ALK-2 sense 5' GATGAGAAGCCCAAGGTCAACC 3', ALK-2
antisense 5' ATGTTCCTGTTACACCAGTCCC 3' (GenBank/EMBL accession no. L15436); ALK-3 sense 5' GTGCTATTGCTCAGGACACTGC 3', ALK-3 antisense 5' AATGAGCACAACCAGCCATCGG 3'
(GenBank/EMBL accession no. Z23154); ALK-6 sense 5'
CACCAAGAAGGAGGATGG 3', ALK-6 antisense 5'
ACAGACAGTCACAGAGATAAGC 3' (GenBank/EMBL accession no. Z32143).
Thirty-five cycles of PCR amplification were performed in a
programmable Dri-Block (Perkin-Elmer) using the following thermal-cycling protocol: 94°C for 1 min, 58°C for 2 min, and 72°C for 3 min, for 35 cycles.
Ligand-receptor affinity binding
studies. Human recombinant BMP-2, OP-1, and TGF-
1 (R
& D Systems) were iodinated as described by Frolik et al. (10). For
receptor binding studies, 0.1 nM 125I-labeled TGF-
, 10 nM
125I-OP-1, or 10 nM
125I-BMP-2 were incubated with
mIMCD-3 cell monolayers and were affinity cross-linked using
disuccinimidyl suberate as previously described (20). For
immunoprecipitations, cells were lysed in lysis buffer [20 mM
tris(hydroxymethyl)aminomethane chloride, pH 7.4, 150 mM NaCl, and
0.5% Triton X-100] in the presence of protease inhibitors and
centrifuged to remove debris.
125I-BMP-2- and
125I-OP-1-labeled cell extracts
were incubated with polyclonal antibodies to ALK-1, ALK-2, ALK-3, and
ALK-6 [generously provided by P. ten Dijke and K. Miyazono (32,
33)], and TGF-
-labeled cell extracts were incubated with
polyclonal T
R-I or T
R-II antibodies (generously provided by J. Massagué). Lysates were incubated with antibodies for 1 to 2 h at
4°C and collected on protein A-Sepharose beads (Pharmacia). The
immunoprecipitates were washed five times in cold lysis buffer and then
resuspended in sample buffer for analysis by sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and autoradiography.
Statistical analysis. Data were
analyzed using the Statview statistical analysis program (version 4.01;
Abacus Concepts, Berkeley, CA). For the dose-response analyses and
explant treatment experiments, mean differences between the effect of
the various ligands were examined by Student's two-tailed
t-test. Differences between
measurements obtained in the morphometric analyses of mIMCD branching
structures were examined by analysis of variance. Significance was
taken at a value of P < 0.05 (two-tailed).
 |
RESULTS |
BMP-2 and OP-1 control kidney
morphogenesis. We identified BMPs expressed in the
developing kidney by RT-PCR cloning using degenerate PCR primers
encoding sequences contained within the BMPs, Vg-1 and decapentaplegic
(1). Comparison of DNA sequences obtained from 28 recombinant plasmids to the nonredundant GenBank/EMBL database using
the BLAST algorithm revealed that 23/28 encode OP-1, whereas 5/28
encode BMP-2.
We first determined the overall effects of BMP-2 and OP-1 on kidney
development by measuring the growth of cultured embryonic murine kidney
explants treated with each BMP. The surface area of each explant was
computed from images obtained at 24 and 48 h after initiation of
culture (Fig. 1), and the percent change in
surface area over 24 h was determined (Fig.
2). Both OP-1 and BMP-2 exerted a direct
and rapid effect on kidney growth. Surprisingly, OP-1 exerted opposite
effects depending on dose. Whereas control kidneys increased in surface
area by ~3%, those treated with 0.25 nM OP-1 increased by 30%
(P = 0.001). In contrast, nanomolar
concentrations of OP-1 exerted a negative effect on explant growth, as
indicated by a 7% decrease in surface area with 28 nM OP-1
(P < 0.001). BMP-2 was entirely
inhibitory (5% inhibition at 5 nM, P < 0.001) and did not stimulate growth at picomolar doses.
Histological analysis indicated that 0.25 nM OP-1 stimulated the
development of both epithelial- and mesenchymal-derived elements (Fig.
3). The ureteric bud/collecting ducts were
notably longer and more highly branched compared with control. In
contrast, 10 nM OP-1 attenuated the development of the ureteric bud and
appeared to reduce the mass of mesenchymal cells. Despite a reduction
in kidney surface area, the architecture of BMP-2-treated explants was
preserved. However, the tissue appeared to be more compact than
control, and the ureteric bud was somewhat attenuated compared with
that observed in 0.25 nM OP-1-treated explants. Together these data indicate a direct role for OP-1 and BMP-2 in regulating embryonic kidney development. However, the cellular complexity of the organ culture explant limits the ability to determine the direct effects of
BMP-2 and OP-1 on cell types within the kidney and specifically on
tubulogenesis. Therefore, we developed an in vitro model using mIMCD-3
cells to investigate the direct effects of BMPs on branching morphogenesis.

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Fig. 1.
Osteogenic protein-1 (OP-1) and bone morphogenetic protein-2 (BMP-2)
control the growth of embryonic kidney explants.
Embryonic day 12 (E12) mouse kidneys were cultured in
serum-free medium containing OP-1 or BMP-2 and photographed as whole
mounts 24 and 48 h after establishing cultures. Representative images
are shown. Explants treated with 0.25 nM OP-1 grew substantially more
than control (no ligand). In contrast, 10 nM OP-1 decreased explant
size. Explants treated with 5 nM BMP-2 also decreased in size.
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Fig. 2.
BMPs control the growth of metanephric explants.
E12 embryonic mouse kidneys were
cultured in serum-free medium containing OP-1 or BMP-2. Explant surface
area was computed from images obtained 24 and 48 h after establishing
cultures (Fig. 1). Data represent the percent change in kidney surface
area between 24 and 48 h in each treatment group; 0.25 nM OP-1
increased surface area by ~30%, whereas explants cultured without
BMP (control) increased by ~3%. In contrast, treatment with 2.8 nM
and 28 nM OP-1 caused a decrease in surface area in a dose-dependent
manner. Treatment with 5 nM BMP-2 also caused surface area to decrease
at a similar quantitative level; n = 3 separate experiments.
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Fig. 3.
BMP-2 and OP-1 control development of epithelial-derived kidney
elements. E12 embryonic mouse kidneys
were cultured in serum-free medium containing OP-1 or BMP-2. After 48 h, paraffin-embedded 5-µm tissue sections were generated from fixed
explant tissue. Images are of hematoxylin-eosin-stained tissue
sections. Left: sections imaged at
×75 magnification. Right:
identical sections imaged at ×150 magnification. Arrows, ureteric
bud/collecting duct structures. Ureteric bud/collecting ducts in 0.25 nM OP-1-treated explants are longer and more highly branched than in
other treatment groups. Treatment with 10 nM OP-1 attenuates this
development. BMP-2 generates short ureteric bud/collecting ducts with
heavily stained resident epithelial cells.
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BMP-2 and OP-1 exert opposite effects on tubulogenesis
in vitro. mIMCD-3 cells are derived from the terminal
inner medullary collecting duct of the SV40 mouse (26) and form
branching structures in three-dimensional matrices (4). The mIMCD-3
cell model provides an opportunity to test the activity of controlled
quantities of ligands on collecting duct morphogenesis without
interaction with other cell types. mIMCD-3 cells cultured in type I
collagen form branching tubular structures when induced with 5% fetal
bovine serum (Fig. 4). We defined
intermediate structures formed after induction of mIMCD-3 cells in
collagen gels by directly imaging structures at different times after
cultures were initiated. Multicellular spheroid structures were
observed within 6 h of induction (Fig. 4,
A and
D), elongated structures by 18 h
(Fig. 4, B and
E), and elongated branching forms by
72 h (Fig. 4, C and
F). Subcellular structures,
specifically large cytoplasmic processes, were observed at the ends of
structures before and during branch formation (Fig. 4G). Cross sections of elongated
branched mIMCD-3 structures revealed that they consist of
interconnected cells organized around a central lumen (Fig.
4H); therefore, they can be defined
as tubules.

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Fig. 4.
mIMCD-3 cells form tubular structures in vitro. mIMCD-3 cells (50,000 cells/gel) were cultured in type I collagen in presence of serum.
A and
D: cellular aggregate after 6 h in
culture imaged by differential interference contrast microscopy (DIC)
(magnification = ×400) (A) and
immunofluorescence microscopy (IF) to identify cell nuclei
(D).
B and
E: elongated structure imaged by DIC
(B) and IF
(E) by 18 h in culture
(magnification = ×400). C and
F: elongated structure with a branch
imaged by DIC (C) and IF
(F) by 72 h in culture
(magnification = ×200). G:
high-power view (magnification = ×400) of a branch point to
identify long, thin cellular processes (arrowhead) arising from cells
at the branch point (left is DIC;
right is IF).
H: cross section of mIMCD-3 structure
stained with hematoxylin and eosin 72 h after initiation of culture.
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We determined the effect of BMP-2 and OP-1 on mIMCD-3 tubulogenesis by
adding these ligands to the culture medium which bathed the collagen
gels. The apoptotic response of mIMCD-3 cells cultured in collagen gels
in the absence of serum precluded analysis of the effect of these
ligands in the absence of serum. Therefore, in these experiments the
effects of BMP-2 and OP-1 were compared in the presence of 5% serum.
To determine whether mIMCD-3 tubulogenesis was regulated in vitro in a
manner consistent with effects observed in whole organ explants
cultured in vitro, we first tested the effects of known positive and
negative regulators of tubulogenesis. TGF-
and activin A are known
inhibitors of tubulogenesis in embryonic kidney explants (4, 27), and
hepatocyte growth factor (HGF) is stimulatory (29). Consistent with
these effects on intact organs, serum alone (Fig.
5A) and
20 ng/ml HGF (Fig. 5B) stimulated mIMCD-3 tubulogenesis. In contrast, 0.5 nM TGF-
and 1 nM activin A
inhibited mIMCD-3 tubulogenesis (Fig. 5,
D and
E). BMP-2 and OP-1 altered both the
number and phenotype of the structures that were formed within 48 h.
This is precisely quantitated below; here, we focus on the broad
phenotype. Treatment with 0.25 nM OP-1 appeared to increase the number
of structures formed compared with serum alone (Fig.
5C). In contrast, 2.5 nM BMP-2
produced a marked reduction in the number of mIMCD-3 structures formed (Fig. 5F). These results suggested
that BMP-2 and OP-1 have opposite effects on the morphogenesis of
mIMCD-3 tubular structures in collagen gels.

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Fig. 5.
Members of the transforming growth factor- (TGF- ) family modulate
the phenotype of mIMCD-3 structures formed in vitro. mIMCD-3 cells
(50,000 cells/gel) were cultured in type I collagen in presence of
serum alone (A) or with hepatocyte growth factor (HGF,
B), TGF- 1
(D), activin A
(E), BMP-2
(F), or OP-1
(C) and imaged by DIC 48 h after
initiation of cultures. Bar in A = 200 µm.
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To precisely define the effects of BMP-2 and OP-1 on mIMCD-3 tubule
formation, we performed dose response analyses of mIMCD-3 tubule
formation in the presence of varying doses of these ligands. We
assessed the number of mIMCD-3 tubular structures formed 48 h
after induction in collagen gels in the presence of serum supplemented medium containing OP-1, BMP-2, TGF-
, or activin A (Fig.
6). For each ligand, four experiments were
performed, and we counted up to 144 mIMCD-3 linear structures in 4 different scaled photographic fields. OP-1 affected mIMCD-3 tubule
formation in a biphasic concentration-dependent manner (Fig.
6A). At low concentrations, OP-1
increased the number of mIMCD-3 tubular structures formed above control
values with a maximum effect observed at 0.25 nM (28 ± 5%
increase, P = 0.01). However, at
concentrations greater than 0.5 nM, OP-1 inhibited the number of
mIMCD-3 tubular structures formed with maximal inhibition observed at
10 nM (81 ± 8% inhibition, P = 0.02). In contrast, the effect of BMP-2 was entirely inhibitory with
maximal inhibition at a dose of 10 nM (88 ± 3% inhibition,
P < 0.0001) (Fig.
6B). BMP-2 was also inhibitory at
concentrations as low as 0.05 nM and was never observed to stimulate
the formation of mIMCD-3 tubules. Both TGF-
and activin A inhibited
the formation of serum-induced mIMCD-3 tubular structures in a
concentration-dependent manner consistent with previous studies (4, 27)
(Fig. 6, C and
D). These results demonstrate that
OP-1 exerts direct but opposite effects on mIMCD-3 tubulogenesis in a
dose-dependent manner. In contrast, BMP-2 is inhibitory and more potent
than high-dose OP-1 and other members of the TGF-
superfamily.

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Fig. 6.
BMP-2 and OP-1 modulate mIMCD-3 tubulogenesis in a dose-dependent
manner. mIMCD-3 cells (50,000 cells/gel) were cultured in type I
collagen in presence of serum with TGF- 1
(C), activin A
(D), BMP-2
(B), or OP-1
(A) and imaged by DIC 48 h after
initiation of cultures. Numbers of tubules (linear structures) formed
are expressed as percentage of control (5% fetal bovine serum alone)
in 4 independent experiments. A: OP-1
is stimulatory at concentrations 0.5 nM [28 ± 5% increase
(max), P = 0.01] and inhibitory at concentrations
>0.5 nM (1 nM, 13 ± 4% inhibition,
P = 0.05; 2.5 nM, 19 ± 11%
inhibition, P = 0.16; 10 nM, 22 ± 8% inhibition, P = 0.02).
B: BMP-2 is inhibitory at all
concentrations tested (1 nM, 45 ± 13% inhibition,
P = 0.04; 2.5 nM, 62 ± 8%
inhibition, P = 0.005; 5 nM, 80 ± 3% inhibition, P = 0.0001; 10 nM, 88 ± 3% inhibition, P < 0.0001).
C: TGF- 1 is inhibitory at all
concentrations tested (0.5 nM, 45 ± 12% inhibition,
P = 0.03; 1 nM, 67 ± 12%
inhibition, P = 0.01).
D: activin A is inhibitory at all
concentrations tested (2.5 nM, 45 ± 14% inhibition,
P = 0.05; 5.0 nM, 39 ± 7% inhibition, P = 0.009; 10 nM, 55 ± 13% inhibition, P = 0.02).
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BMP-2 and OP-1 modulate the phenotype of mIMCD-3
tubular structures. Our initial results suggested that
BMP-2- and OP-1-treated structures differ in their phenotype.
Therefore, we analyzed their structural characteristics. Representative
structures imaged simultaneously by DIC and immunofluorescence
microscopy are shown in Fig. 7. Morphometric measurements were made using photographed images of 155 structures generated by 2 days and 7 days after induction in collagen
gels consisting of a low density of cells (5,000 cells/gel). At this
low concentration of cells, mIMCD-3 structures can be imaged at high
resolution without optical interference by other adjacent structures.
Compared with the structures imaged in higher cell-density gels (Fig.
5), structures formed in low-density gels contain fewer cells and
develop more slowly as a function of time.

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Fig. 7.
Morphometric analysis of BMP-2- and OP-1-treated mIMCD-3 structures.
mIMCD-3 cells (5,000 cells/gel) were cultured in type I collagen in
presence of serum and 5 nM BMP-2 or 0.25 nM OP-1. Two days or seven
days after initiation of culture, fixed structures were stained with
bis-benzamide and imaged by DIC
(A,
C, E,
and G) and IF
(B,
D, F,
and H) microscopy. As exemplified in
E, the long axis (solid bar) and the
number of branch points (arrowheads) were determined for each imaged
structure. Arrow in C marks the
position of a cellular process extending from a branch point.
A and
B: BMP-2-treated structure after 2 days in culture (length = 58 µm). Structure is short and unbranched.
C and
D: OP-1-treated structure after 2 days
in culture (length = 96 µm). Structure is long relative to BMP-2
structure and consists of a branch point.
E and
F: BMP-2-treated structure after 7 days in culture (length = 148 µm). Structure is short and
consists of short, stubby branches. G
and H: long, multibranched
OP-1-treated structure after 7 days in culture (length = 364 µm).
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We analyzed the morphometric characteristics of mIMCD-3 tubules during
the early phase of tubulogenesis (2 days) and when mature tubules with
lumens were formed (7 days). OP-1-treated structures differed from
BMP-2-treated structures in the number of branch points per structure
and in length (Table 1). Two days after
induction of tubulogenesis, OP-1-treated structures were more highly
branched than BMP-2-treated structures [OP-1
(n = 48) vs. BMP-2
(n = 44): 0.6 ± 0.1 vs. 0.07 ± 0.05 branch points/structure, respectively; P < 0.0001] (Fig. 7,
A-D).
Only 3/44 BMP-2-treated structures contained branch points or were
branched. Further analysis of structures arising from branch points
revealed that only three of all the branches generated in OP-1- and
BMP-2-treated structures contained nuclei (Fig. 7, C and
D). This suggests that an early step in the
formation of a branch is the formation of a cell process and that BMP-2
inhibits the formation of these cell processes. In addition,
OP-1-treated structures were longer than BMP-2-treated structures
[OP-1 (n = 48) vs. BMP-2
(n = 44): length, 40.6 ± 4.9 vs.
29.5 ± 3.1 µm; P = 0.06].
Despite this difference in length, the number of nuclei/tube did not
differ between OP-1- and BMP-2-treated structures [OP-1
(n = 48) vs. BMP-2
(n = 44), 1.8 ± 0.3 vs.
1.8 ± 0.4 nuclei/tube]. This suggests that OP-1 increases
cell size.
In contrast to 2-day structures, 7-day structures contained a much
larger number of cells [OP-1 (n = 43) and BMP-2 (n = 20), 76 ± 12 and 38 ± 14 nuclei/structure, respectively]. Consistent with
our analysis of 2-day structures, OP-1-treated structures were more
highly branched than BMP-2-treated structures [OP-1 vs. BMP-2, 13 ± 2 vs. 4 ± 1 branch points/structure;
P < 0.0001] (Fig. 7,
E-H).
The branches arising from these branch points consisted of true
branches (contain nuclei) and pseudo branches (consist of cell
processes only). Both true and pseudo branches were more numerous in OP-1-treated structures compared with
BMP-2-treated branches [OP-1 vs. BMP-2 true branches, 7 ± 1 vs. 2 ± 1 true branches/structure (P = 0.0003); pseudo branches, 5 ± 1 vs. 1 ± 1 pseudo branches/structure (P < 0.0001)]. Also
consistent with our analysis of 2-day structures was our finding that
7-day OP-1-treated structures were longer than BMP-2-treated
structures, probably secondary to the larger number of cells in
OP-1-treated tubules (OP-1 vs. BMP-2: length, 191 ± 14 vs. 118 ± 16 µm; P = 0.003). However,
although BMP-2 inhibited tubular growth and branching, the smooth
borders and width of BMP-2-treated tubules were more similar to native
tubules than those treated with OP-1 (Fig. 7,
E and G) (30). This suggests that BMP-2 plays a
role in modulating the final shape of developing branched tubules.
Taken together, our analysis of early and mature tubular structures
suggests that 1) BMP-2 and OP-1
exert different activities during the formation of branched tubules,
2) the formation of a true branch is
preceded by the elaboration of a cytoplasmic process by the cell, and
3) BMP-2 inhibits the formation of
these branches possibly by interfering with cell process formation.
mIMCD cells express mRNAs encoding BMP
receptors. Having demonstrated that BMP-2 and OP-1
control the number and type of mIMCD-3 structures formed in collagen
gels, we sought to define the molecular basis for these effects. Since
the specificity of the effects of these ligands resides, in part, with
the cell surface receptors which bind them, we first identified
receptors that are expressed in the
E13 metanephros, a stage at which
collecting duct morphogenesis is occurring. Since members of the
TGF-
superfamily are known to signal via a highly conserved family
of cell surface transmembrane serine/threonine kinases, we used
a RT-PCR cloning strategy to identify receptor mRNAs (9).
By this method we identified cDNAs encoding T
RI (ALK-5), T
RII,
ActR1B (ALK-2), ActRIIB, ALK-3, and ALK-6 (Table
2). Next, we determined by RNase protection and RT-PCR assays whether these mRNAs are also expressed by mIMCD-3 cells (Fig. 8; Table 2). RNase protection
assays using exact match cRNA probes demonstrated protection of RNA
fragments of 200, 380, 390, and 400 bp in size predicted for T
RI
(ALK-5), T
RII, ActR1B (ALK-2), and ActRIIB, respectively. These are
the known receptors for TGF-
and activin A, respectively. Positive controls in this assay included
-actin and c-met, the receptor for
HGF (3). RT-PCR and Southern analysis using exact match primers and
cDNA probes demonstrated that mIMCD-3 cells express ALK-3 and ALK-6,
type I receptors known to bind BMP-2 (Fig. 8, B and
C). Interestingly, we did not detect
expression of ALK-2, a candidate OP-1 receptor. Together, these results
demonstrate that the E13 metanephros
and mIMCD-3 cells express mRNAs encoding type I and type II receptors,
which bind TGF-
superfamily members including BMP-2.

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Fig. 8.
mIMCD-3 cells express a broad repertoire of type I and type II
serine/threonine kinase receptor mRNAs.
A: RNase protection assay
demonstrating protection of mIMCD-3 RNAs using
32P-labeled antisense RNA probes
for c-met, T R1, T RII, ActR1B, ActRIIB, and actin.
B: agarose gel of DNA bands generated
by reverse transcription-polymerase chain reaction (RT-PCR) of mIMCD-3
poly(A)+ mRNA using exact-match
oligonucleotide primers for ALK-3 and ALK-6.
C: Southern analysis of agarose gel,
shown in B, demonstrating
hybridization of 32P-labeled ALK-3
and ALK-6 cDNA probes to candidate ALK-3 and ALK-6 DNA bands generated
by RT-PCR of mIMCD-3 RNA. MW, molecular weight markers.
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BMP-2 binds to mIMCD-3 cells via
ALK-3. To further define, at the protein level, the
receptors that bind BMP-2 and OP-1, we performed ligand-receptor
cross-linking with immunoprecipitation using antisera directed against
type I and type II receptors (Fig. 9).
Isolated receptors were resolved by SDS-PAGE. In these experiments, mIMCD-3 cell receptor affinity labeled with
125I-TGF-
served as a positive
control. As expected, we observed three labeled products corresponding
to betaglycan, type II and type I receptors, typically
observed in a wide variety of cell lines responsive to TGF-
(38).
The identities of T
RII and T
RI were confirmed by
immunoprecipitation of total cell lysates with specific receptor
antibodies (Fig. 9, middle). Since
TGF-
receptors typically form stable heteromeric
complexes, immunoprecipitates using anti-T
RII antibodies contained
affinity-labeled species corresponding to T
RII and additional
species corresponding to betaglycan and T
RI, as
observed previously (36). Similarly, immunoprecipitates of anti-T
RI
antibodies contain labeled products corresponding to both T
RI and
T
RII.

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Fig. 9.
BMP-2 and OP-1 bind to mIMCD-3 cell surface proteins.
125I-labeled ligand was
cross-linked to mIMCD-3 cells in monolayer. Labeled ligand-receptor
complexes were immunoprecipitated using antisera directed against type
I and type II receptors and analyzed by SDS-PAGE.
Middle: total cell lysate after
labeling of cells with
125I-TGF- 1 contains 3 labeled
products, >180, 80-85, and 69 kDa in size. Immunoprecipitation
of cell lysate with anti-T RII yielded two bands, the size of T RI
and T RII. Immunoprecipitation of cell lysate with anti-T RI
yielded a major band the size of T RI and a minor band the size of
T RII. Left: total cell lysate after
labeling of cells with 125I-BMP-2
contains 3 labeled products, 180, 80-85, and 75 kDa in size.
Immunoprecipitation of the cell lysate with specific type I receptor
antibodies demonstrating that only anti-ALK-3 immunoprecipitates
contain affinity-labeled complexes.
Right: total cell lysate after
labeling of cells with 125I-OP-1
contains five labeled products, 200, 180, 150, 85, and 80 kDa in size.
Immunoprecipitation of the cell lysate with ActRII/IIB antibodies
demonstrates that OP-1 binds this type II receptor as in the cell line
C5.18. Immunoprecipitation of cell lysate with specific type I receptor
antibodies demonstrates that the 80-kDa protein (p80) is not ALK-1,
ALK-2, ALK-3, or ALK-6.
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mIMCD-3 cells labeled with
125I-BMP-2 yielded three labeled
products, a high-molecular-weight band at ~180 kDa, a diffuse 80- to
85-kDa band, and a faster migrating 75-kDa band (Fig. 9,
left). The two larger proteins
likely represent the two alternatively spliced forms of BMPR-II (18).
Alternatively, the 80- to 85-kDa form may also represent ActRII and
ActRIIB, which under the appropriate conditions also bind BMP-2 (L. Attisano and J. L. Wrana, unpublished observations). To identify the
specific BMP-2 binding type I receptors, lysates from affinity-labeled
cells were subjected to immunoprecipitation with specific type I
receptor antibodies. Only anti-ALK-3 immunoprecipitates contained
affinity-labeled products. Furthermore, as expected from the known
ability of BMP receptors to form heteromeric complexes (17), additional
affinity-labeled products of 80-85 kDa and 180 kDa were also
observed. These data indicate that the candidate cell surface receptor
in the BMP-2 signaling pathway is ALK-3.
Affinity labeling of mIMCD-3 cell surface proteins with
125I-OP-1 yielded five labeled
protein bands, 200, 180, 150, 85, and 80 kDa in size (Fig. 9,
right). The 180- and 150-kDa bands
may represent different forms of BMPR-II (18). Immunoprecipitation of
mIMCD-3 proteins in the cell lysate with antibodies specific to
ActRII/IIB yielded the 85-kDa band, as in the control cell line C5.18.
To further characterize binding of OP-1 to p80, we performed
competition assays using unlabeled OP-1. As previously observed (39),
addition of unlabeled OP-1 did not displace binding of labeled OP-1 but
rather increased binding to the cell lysates (data not shown). The
basis for this phenomenon is currently unclear (39). To attempt to
determine the identity of p80, we performed immunoprecipitation with
antibodies directed against the type I receptors, ALK-1, ALK-2, ALK-3,
and ALK-6. However, none of these antibodies identified p80. These
results suggest that the OP-1 signaling pathway in mIMCD-3 cells
includes ActRII/IIB and another as yet unidentified receptor that may
correspond to p80.
OP-1 exerts both stimulatory and inhibitory effects on
branching morphogenesis in the developing kidney. On
the basis of our results in embryonic kidney explants and in the
mIMCD-3 model of collecting duct morphogenesis, we hypothesized that
OP-1 exerts opposite effects on branching morphogenesis in different
areas of developing kidney tissue depending on its local concentration. To test this directly, we delivered recombinant OP-1 and BMP-2 in a
spatially restricted manner to developing murine
E12 kidneys by saturating agarose
beads with a micromolar amount of ligand and placing ligand-saturated
beads on the kidney surface (Fig. 10)
(25). Consistent with our previous results, the growth of kidney
explants treated with high doses of OP-1 or BMP-2 was significantly less than that of kidneys treated with beads saturated with bovine serum albumin (Table 3). Since branched
tubular structures are formed symmetrically on either side of the
ureteric bud, we tested the effect of these BMPs on branching
morphogenesis by applying them to one side of the kidney. We then
compared the number of ureteric bud/collecting duct branches that
formed in the side adjacent to the beads to that on the untreated
contralateral side. On the sixth day in culture, control kidneys
contained a nearly equivalent number of D. biflorus agglutinin-stained collecting system branches
on either side of the ureteric bud (ratio of mean number on each side = 0.92 ± 0.01; Table 3). In contrast, the number of branches on the
side receiving either 2 µM BMP-2 or 2 µM OP-1 beads was 64 ± 2% and 58 ± 1% less than that on the untreated contralateral side
(P < 0.0001). Furthermore, in the tissue immediately adjacent to BMP-2 and OP-1 beads, which presumably received the highest dose of ligand, no ureteric bud/collecting duct
structures were observed. In contrast, we consistently observed that
the collecting system in the region of the kidney distal to the OP-1
beads was more highly branched than in any region of the control or
BMP-2-treated explants. In this region, the concentration of OP-1 is
predicted to be lower than in areas adjacent to the beads secondary to
diffusion of the ligand. We attempted to demonstrate directly that
agarose beads soaked in low doses of OP-1 stimulate branching
morphogenesis in explants. We did not observe any stimulation under
these conditions. This negative result is likely explained by the
relatively narrow dosage range in which OP-1 is stimulatory, the strong
affinity of OP-1 for the extracellular matrix (35), and the slow
kinetics of peptide release from agarose beads (13, 31). Polypeptides
such as epidermal growth factor and fibroblast growth factor are bound to agarose beads with an efficiency of 90-95%. Approximately 50% of bound protein is released within the first 24 h, whereas 10% is
released during every subsequent 24-h period. Thus these release kinetics combined with the narrow dose response range and sequestration of growth factor by the extracellular matrix could easily mitigate against a stimulatory effect in explant tissue. However, taken together
with our in vitro data, our results strongly support our hypothesis
that OP-1 exerts different morphogenetic effects, depending on its
local concentration in developing kidney tissue.

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Fig. 10.
OP-1 and BMP-2 modulate branching morphogenesis in embryonic kidney
explants. E12 embryonic mouse kidneys
were cultured in vitro in serum-free medium and photographed on the
second and sixth days after establishing cultures. On
day 6 of culture, explants were fixed
and stained with fluorescein-conjugated D. biflorus agglutinin (DBA) to identify ureteric
bud/collecting duct branches (C,
F, and
I). Whole mount kidney explants were
photographed on day 2 (A,
D, and
G) and day
6 (B,
E, and
H) of culture, respectively. Agarose
beads are positioned in the upper pole of the explant treated with
BMP-2 (D and
E). Positions of beads in
DBA-stained BMP-2-treated explant are marked by the two arrows
overlying tissue on right ureteric bud
(F). Branching structures are absent
in tissue adjacent to these beads. Position of agarose beads in
OP-1-treated kidneys is marked by arrowhead next to the ureteric bud
(G and
H). Position of these agarose beads
in DBA-stained OP-1-treated explant is marked by 2 arrows at
top. Branching structures are absent
in tissue adjacent to these beads. In contrast, a multibranched
structure (arrowhead) is present distal to the OP-1 beads
(I).
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 |
DISCUSSION |
Previous studies have demonstrated the spatial expression of OP-1 and
BMP-2 early in mouse kidney development and have suggested a role for
these BMPs in mesenchymal-epithelial interactions and their
morphogenetic consequences (2, 8). During early murine kidney
development, OP-1 mRNA is expressed in the ureteric bud and in the
induced metanephric mesenchyme. In contrast, BMP-2 mRNA expression is
restricted to the metanephric mesenchyme. Later, during maturation of
mesenchymal-derived elements, OP-1 is expressed in maturing glomeruli,
whereas BMP-2 expression decreases markedly. Although functional tests
have indicated a role for OP-1 in the induction of the metanephric
blastema (7, 19, 34), there is no direct evidence that either OP-1 or
BMP-2 controls branching morphogenesis.
Our experiments in embryonic kidney explants (Figs. 1-3) and in an
in vitro model of collecting duct morphogenesis (Figs. 5 and 6)
demonstrate that OP-1 exerts opposite effects on branching morphogenesis in a dose-dependent manner. Picomolar amounts of OP-1
stimulate, whereas nanomolar amounts inhibit. In vitro, low doses of
OP-1 (0.25 nM) increase mIMCD-3 tubule length and branching, and
100× higher doses attenuate these effects. The effects of 0.25 nM
OP-1 and 10 nM OP-1 on embryonic kidney explants are consistent with
the effects in vitro. In contrast, the effects of BMP-2 are monophasic
and inhibitory. BMP-2 inhibits the formation of tubules. However, its
qualitative effects are more complex. Although BMP-2 inhibits branch
formation and linear growth, it generates a more sculpted, thicker
tubular trunk in vivo and in vitro (Figs. 3 and 7). Thus OP-1 and BMP-2
may act cooperatively to regulate both the number and the structural
characteristics of developing branched tubules.
The mechanism that underlies the stimulatory effect of OP-1 (<0.25
nM) on tubulogenesis is undefined. Further studies are needed to
determine the effect of OP-1 on cellular mechanisms which appear to be
critical during tubulogenesis. These include cell proliferation, cell
process formation, and cell-cell adhesion (22). The mechanisms that
underlie the inhibitory effect of BMP-2 and OP-1 (>0.25 nM) are also
undefined. However, the observation that activation of the ALK-3
receptor mediates apoptosis in the developing limb (41) suggests that
treatment of mIMCD-3 cells in collagen gels with BMP-2 may induce
apoptosis via activation of ALK-3, thereby decreasing the number of
cells capable of forming tubular structures. Further studies are
required to test this possibility and to determine whether the
inhibitory effect of high-dose OP-1 is mediated by a similar mechanism.
Our data provide insights into the mechanisms of branch formation. The
structures that arise from branch points in immature OP-1-treated
structures are cellular processes. More mature structures consist of
true branches (consisting of one or more cells) that arise from these
cellular processes. Immature BMP-2-treated structures rarely consist of
a branch point. Thus our data suggest that BMP-2 controls branch
formation by inhibiting the formation of cellular processes. Recent
studies have defined some of the molecular elements involved in
filopodia formation (23) and in the formation of tubulin-based
processes (22), and this information may provide a basis to test the
effects of BMP-2 and OP-1 on these pathways. Thus further studies are
required to characterize whether these cellular processes are actin or
microfilament based.
OP-1 also increases the length of mIMCD-3 structures. In our analysis
of 2-day, immature structures and 7-day, mature structures, we found no
significant difference in the number of resident nuclei within OP-1-
vs. BMP-2-generated tubules. This suggests that
properties other than cell number (e.g., cell shape, cell size)
determine the length of these structures. Further studies will be
required to test whether the effects of these BMPs on elements of the
cytoskeleton are related to this observed effect on cell shape.
The differential effects of BMP-2 and OP-1 on branching morphogenesis
are surprising, since both of these growth factors signal via a highly
conserved family of cell surface receptors. To determine the molecular
basis for these effects, we identified type I and II serine/threonine
cell surface receptors, which can serve as binding partners for BMP-2
and OP-1 (Fig. 9). These results provide a biochemical basis for the
phenotypic effects we observed. We demonstrate that the BMP-2 cell
surface receptor complex consists of ALK-3 and a type II receptor, the
size of which is consistent with either BMPRII or ActRIIB. These
results are consistent with the recent demonstration that
ligand-dependent phosphorylation of MADR1, a downstream effector of
BMP-2, requires the coexpression of type I and type II BMP-2 receptors
and can be mediated by either ActRIIB or BMPRII in
conjunction with either ALK-3 or ALK-6 (15). Our results strongly
suggest that in mIMCD-3 cells, ALK-3 is the type I receptor that
signals in the BMP-2 pathway. Our results also demonstrate that the
mIMCD-3 OP-1 receptor complex consists of the type II receptor,
ActRII/IIB. We were not able to identify type I receptors (ALK-2,
ALK-3, ALK-6) in this complex using specific antibodies. This may be
because receptors such as ALK-2 are expressed in lower than detectable
quantities. Our inability to detect receptors with these antibodies is
not explained by cross-species differences, since the ALK-6 antibody is
directed against a mouse peptide (32), the ALK-3 antibody identifies
ALK-3 in the mIMCD-3 BMP-2 receptor complex, and the ALK-2 antibody is
directed against a human peptide that is identical in mouse ALK-2. We
did identify an unknown protein, p80, in the OP-1 receptor complex.
However, the failure of unlabeled OP-1 to competitively displace
radiolabeled OP-1 as previously described (39) limited our ability to
determine whether p80 is a specific OP-1 receptor. This unusual
property of OP-1 is possibly due to its high affinity for the
extracellular matrix (35).
Our demonstration that BMP-2 and OP-1 have opposite effects on mIMCD-3
tubulogenesis is consistent with previous observations that members of
the TGF-
superfamily can induce different biological responses (16).
For example, during patterning of the
Xenopus embryo, activins can function
to induce dorsal mesoderm while BMP-4 induces ventral mesoderm (12).
The specificity of these responses appears to be mediated by the
particular type I receptor that is activated by a particular BMP-type
II receptor complex. Type I receptors with highly related kinase
domains can mediate similar biological responses, whereas more
distantly related type I receptors may mediate distinct responses (5).
Such is the case for BMP-2 signaling, which is regulated by ALK-3 and
ALK-6 but not by ALK-2 and ALK-5 (15). At the level of protein
homology, the kinase domains of ALK-3 and ALK-6 are highly related to
each other but share less identity with those of ALK-1, ALK-2, ALK-4, and ALK-5 (33). These observations support a hypothesis that predicts
that at low concentrations, OP-1 signals via a type I receptor other
than ALK-3 and ALK-6. Our studies demonstrate that the mIMCD-3 OP-1
receptor, p80, is not ALK-1, ALK-2, ALK-3, or ALK-6. In addition, our
demonstration that OP-1 stimulates mIMCD-3 tubulogenesis and increases
the number of branches and the length of structures suggests that the
OP-1 receptors signal via a stimulatory pathway distinct from the BMP-2
pathway. The elements in both stimulatory and inhibitory pathways
downstream of BMP-2 are largely unknown. Future studies
aimed at identifying these elements will provide a necessary
biochemical basis for the effects of these BMPs at the cellular level.
The growth and branching of the ureteric bud and its daughter
collecting ducts appears to be tightly regulated during
renal development. The expression of growth factors that signal via stimulatory or inhibitory pathways may serve as a mechanism to control
these processes. Our observations that BMP-2 and OP-1 differentially
regulate both the number and the phenotype of branched tubules in a
dose-dependent manner suggest that both these BMPs are important
regulatory molecules for these morphogenetic events. This is consistent
with the known spatial expression of BMP-2 and OP-1 in the developing
kidney and the knowledge that BMPs play diverse roles during the
development of nonrenal tissues. Further studies aimed at determining
the cellular targets of these growth factors in vivo during metanephric
development will further define their role in regulating collecting
duct development.
 |
ACKNOWLEDGEMENTS |
We thank Ivanka Antoniewicz for expert technical assistance in the
performance of these studies and Judith McNicoll for expert secretarial
support. We also thank Herman Yeger for helpful assistance with
metanephric organ culture and P. ten Dijke, K. Miyazano, K. Heldin, W. Vale, and J. Massagué for kindly providing antibodies used in
these studies.
 |
FOOTNOTES |
This work was supported, in part, by a Kidney Foundation of Canada
Scholarship, I'Anson Professorship, and Pediatric Consultants (Hospital for Sick Children) Grant (to N. D. Rosenblum),
Medical Research Council of Canada Scholarships and Operating Grants
(to L. Attisano and J. L. Wrana), and a National Cancer Institute of
Canada Operating Grant (to J. L. Wrana).
Address for reprint requests: N. D. Rosenblum, Division of Nephrology,
Program in Developmental Biology, The Hospital for Sick Children, Univ.
of Toronto, 555 Univ. Ave., Toronto, Ontario, Canada M5G 1X8.
Received 26 July 1997; accepted in final form 18 August 1997.
 |
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