1 Division of Gene Function in Animals, Nara Institute of Science and
Technology, 8916-5 Takayama, Ikoma, Nara 630-0101, Japan
2 Division of Metabolic Regulation of Animal Cells, Nara Institute of Science
and Technology, 8916-5 Takayama, Ikoma, Nara 630-0101, Japan
* Author for correspondence (e-mail: mkawaich{at}bs.aist-nara.ac.jp)
Accepted 21 November 2003
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SUMMARY |
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Key words: HtrA1, Tgfß, Bmp, Serine protease, Noggin, Follistatin
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Introduction |
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Human HtrA2 (omi) is the most characterized member of
mammalian HtrA genes. HtrA2 induces apoptosis in a
caspase-independent manner through its protease activity and in a
caspase-dependent manner via its ability to activate caspases
(Suzuki et al., 2001). The
crystal structure of HtrA2 has recently been elucidated
(Li et al., 2002
). In the
crystals, three monomers of HtrA2 are assembled into a pyramid-shaped
structure. The trimer structure is essential for the proteolytic activity of
HtrA2.
Human HtrA1 (L56 or RSPP11) was originally
isolated as a gene whose expression was downregulated in a human fibroblast
cell line after transformation with SV40
(Zumbrunn and Trueb, 1996).
Although HtrA1 and HtrA2 share homologous SP domains and PDZ domains in the
C-terminal regions, HtrA1 contains a signal sequence for secretion, as well as
an insulin-like growth factor binding protein (IGFBP) domain and a Kazal-type
serine protease inhibitor (KI) domain in the N-terminal region (see
Fig. 5 for HtrA1 structure).
Downregulation of human HtrA1 has been repeatedly observed in ovarian
cancers (Shridhar et al.,
2002
) and melanomas, in close correlation with the malignant
progression and metastasis of these tumors
(Baldi et al., 2002
).
Overexpression of HtrA1 in highly invasive melanomas was shown to
suppress proliferation and migration of tumor cells
(Baldi et al., 2002
).
HtrA1 is thus considered to be a tumor suppressor gene in certain
cancers.
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The Tgfß family consists of more than 30 proteins in mammals,
including Tgfßs, Bmps, growth/differentiation factors (Gdfs) and activin
(for a review, see Massagué and
Chen, 2000). These proteins are expressed in distinct and complex
spatio-temporal patterns during embryogenesis and in adult tissues, playing
pivotal roles in the development and homeostasis of these tissues. Although
Tgfß signaling can be regulated at various steps, it is largely regulated
extracellularly through receptor activation. A group of antagonists have been
identified that bind to Bmp subfamily proteins in the extracellular space,
thereby interfering the receptor activation. Noggin and chordin are
antagonists of several Bmps, whereas follistain is an antagonist of activin.
New members, like Dan family proteins, are continuously being added to the
list of Bmp antagonists (Balemans and Van
Hul, 2002
). However, there are only a few proteins known to be
antagonists of the Tgfß subfamily.
We report that the expression pattern of mouse HtrA1 shows a striking correlation with the sites where signaling of the Tgfß family regulates development. We also present evidence that HtrA1 binds to a broad range of Tgfß proteins and inhibits their signaling in vitro as well as in vivo. Surprisingly, the binding and inhibitory activities of HtrA1 on Tgfß proteins depend on the integrity of the HtrA1 serine protease domain.
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Materials and methods |
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In situ hybridization of mRNA
Whole embryos and tissues were fixed in 4% paraformaldehyde in phosphate
buffered saline (PBS) overnight at 4°C. Whole-mount and section in situ
hybridization were performed as previously described (Sasakli and Hogan, 1994;
Morimoto et al., 1996).
Non-overlapping fragments of HtrA1 cDNA, a
BalI-BamHI fragment (nucleotide 644-1191) and a
BamHI-HincII fragment (nucleotide 1191-1750), cloned in
BluescriptKSII(+) (Stratagene) were used as templates to synthesize
digoxigenin-labeled RNA probes using a DIG-RNA labeling kit (Boehringer).
Antibodies, immunostaining and western blot analysis
N- and C-terminal HtrA1 fragments (aa 28 to 143 and aa 385 to 480) produced
in E. coli were injected into rabbits to raise antibodies, and
antiserum was affinity purified. Purified polyclonal antibodies against the
N-terminal fragment were used for immunohistochemistry. This antibody did not
cross react with mouse HtrA3. Immunohistochemical staining was carried out
using the tyramide signal amplification-avidin-bioton-complex method as
previously described (Toda et al.,
1999). Adult bones were decalcified in 10% EDTA (pH 7.4), and
embedded in paraffin wax. For western blot analysis, protein samples were
separated by SDS-polyacrylamide gel electrophoresis (PAGE) and transferred to
nitrocellulose membranes. Membranes were probed with the appropriate primary
antibodies and then with alkaline phosphatase-labeled secondary antibodies,
followed by the BCIP/NBT color reaction.
GST pull-down assay
The cDNA fragments encoding the mature peptides of mouse Tgfß1,
Tgfß2, Bmp4, Gdf5 and activin were cloned into the pGEX-4T-1 vector
(Stratagene) in-frame using BamHI and EcoRI sites. All GST
fusion proteins were expressed in insoluble fractions. They were solubilized
in 8 M urea containing buffer and then renatured, as previously described
(Groppe et al., 1998). The
renatured proteins were dialyzed against the GST pull-down buffer [20 mM
Tris-HCl (pH 8.0), 500 mM NaCl, 0.05% NP-40, 10% glycerol]. HtrA1, its mutant
forms and myc-tagged follistatin were produced in sf9 cells using a
baculovirus system. Each sf9 culture supernatant was mixed with one of the GST
fusion proteins and incubated overnight at 4°C in 400 µl of 20 mM
Tris-HCl (pH 8.0), 1.5 mM CaCl2, 3 mM MgCl2, 5%
glycerol, 0.1% NP-40, 0.3-1.0 M NaCl, 5% fetal bovine serum (FBS) to minimize
non-specific binding. Glutathione-Sephrose 4B beads (Amersham Pharmacia
Biotech) (20 µl) were added to the mixture and incubated for 1 hour at
4°C. Beads were recovered by centrifugation and washed five times in 500
µl GST pull-down buffer. Proteins bound to the beads were separated by
SDS-PAGE and analyzed by western blotting, as described above. Pull-down
assays in the presence of 1.0 M NaCl gave the most specific and reproducible
results.
Solid phase binding assay
ELISA plate (Nunc) wells were coated with 50 µl of either 10 µg/ml of
GST or GST-Bmp4 in 20 mM Tris-HCl (pH 7.5) and 500 mM NaCl at 4°C
overnight. The wells were washed twice with a wash buffer [0.05% Tween 20 in
TBS (20 mM Tris-buffered saline, pH 7.5)] and non-specific binding sites were
blocked with 0.1% skimmed milk in wash buffer. Various amounts of myc-tagged
HtrA1 protein in Dulbecco's Modified Essential Medium (DMEM) containing 10%
FBS were added to the wells and incubated at 4°C overnight. The wells were
then washed twice with wash buffer, incubated with anti-myc antibody, washed
five times with wash buffer, and then incubated with alkaline
phosphatase-conjugated anti-mouse IgG antibody. The bound phosphatase was
assayed using p-nitrophenyl phosphate. The reaction was stopped with
100 mM EDTA, and absorbance at 405 nm was measured. The difference between the
absorbance value for GST-Bmp4 and that for GST alone was considered to
represent specific binding and was used to determine the Kd value. The
concentration of HtrA1 in the medium was quantitated by western blotting with
anti-HtrA1 antibody using bacterially produced HtrA1 as the standard.
Cell culture, transfection and luciferase assay
Mouse C2C12 myoblasts were cultured in DMEM supplemented with 15% FBS in
24-well plates. Four hours before transfection, growth medium was replaced
with fresh medium containing 2% FBS. Transfection of cells was carried out
using the calcium phosphate method with 125 ng of a ß-galctosidase
expression vector as an internal control and other plasmid DNAs, as described
in the figure legends. Cell extracts were prepared 24 hours after transfection
and assayed for luciferase activity using the Picagene kit (Toyo Ink Co. Ltd,
Tokyo). Luciferase activity was corrected for transfection efficiency using
ß-galactosidase activity. Transfection of C2C12 cells with the
Bmp4 or caBMPR-IB expression vector stimulated the
pGL3-Id985WT reporter several fold, as described previously
(Pearce et al., 1999;
Katagiri et al., 2002
).
Smad1 and/or Smad4 expression further stimulated the
pGL3-Id985WT or pGL3ti-(SBE)4 reporters in the presence of Bmp4 or
Tgfß1, respectively, although Smad expression alone did not
activate those reporters.
Misexpression of HtrA1 in chick embryo
HtrA1 cDNA was inserted into the pCAGGS vector in two orientations
to express sense and antisense strands. Cultured 293T cells were
co-transfected with an EGFP expression vector and a pCAGGS vector, containing
sense or antisense cDNA, using Lipofectamine 2000 (Invitrogen). The
pEF-Bos-noggin vector was used to express noggin. Cells were harvested 48
hours after transfection and suspended in a small amount of culture medium.
Cell suspension (less than 0.1 µl) was injected into the extraocular
mesenchyme.
The pCAGGS-HtrA1 vector and the EGFP vector were electroporated into the
optic vesicles, as described previously
(Koshiba-Takeuchi et al.,
2000). After incubation for 24-27 hours, embryos were subjected to
whole-mount in situ hybridization using a 1.4-kb cVax probe.
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Results |
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In order to precisely locate HtrA1 expression, we carried out in
situ hybridization and immunostaining of embryos and adult tissues. Two
non-overlapping probes were used for in situ hybridization and both probes
gave identical results. For immunostaining, we used polyclonal antibodies
raised against the N-terminal region of HtrA1 (aa 28-143). The results of
immunostaining were consistent with those of in situ hybridization in various
tissues (see Fig. 2D,E),
ensuring the specificity of this antibody. In the 12.5 dpc embryo,
HtrA1 mRNA was expressed in a pattern closely associated with
skeletal elements of the vertebral column and limbs
(Fig. 1A). In the sagittal
section of the 12.5 dpc embryo, expression of HtrA1 was also found in
the brain, the heart, the lungs, the gonads and the skin of the
thoraco-abdominal region (Fig.
1B). Expression of HtrA1 became enhanced at 14.5 dpc
(Fig. 1C). In the skeletal
tissues, HtrA1 was expressed in rudiments of tendons and ligaments
along the vertebrae, as well as in mesenchymal cells surrounding precartilage
condensations (Fig. 1B,C,D). In
the brain, HtrA1 was expressed in specific regions of the
neuroepithelium in the forebrain and hindbrain
(Fig. 1B), in close association
with development of the choroid plexus. This was clearly seen in the hindbrain
of the 14.5 dpc embryo (Fig.
1E), where expression of HtrA1 was detected in the
neuroepithelium adjacent to the forming choroid plexus. The mature choroid
plexus, however, did not express HtrA1
(Fig. 1E). It has been reported
that several members of the Bmp subfamily are expressed in the developing
brain with patterns characteristic to each member. Interestingly, choroid
plexus development sites are those regions where all Bmps are co-expressed
(Furuta et al., 1997).
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A striking correlation between the expression patterns of HtrA1 and the functioning sites of various Tgfß proteins, together with the presence of a domain similar to follistatin in HtrA1 protein, suggests that HtrA1 plays a role in Tgfß-mediated signaling.
HtrA1 expression in developing skeletal elements
Because skeletal tissues are the most intensively studied organs in
relation to Tgfß signaling during morphogenesis, and because expression
of HtrA1 was reported to increase in articular cartilage cells of
human arthritic patients (Hu et al.,
1998), we conducted a detailed examination of HtrA1
expression during development of bones and related tissues. As described above
for 12.5 and 14.5 dpc embryos, HtrA1 was expressed in
undifferentiated mesenchymal cells surrounding precartilage condensations but
was not detected in the condensation cores
(Fig. 1B,C,D). However, within
the precartilage condensations, joint interzones, which later form joint
cavities, expressed HtrA1 (Fig.
2A,B). Rudiments of tendons and ligaments that develop from
mesenchyme in the vicinity of the precartilage condensations expressed
HtrA1 (Fig. 1D; Fig. 2B,C). In the process of
endochondral ossification, matured chondrocytes become hypertrophic,
degenerate and are then replaced by calcified bones. Although precartilage
condensations and chondrocytes did not express HtrA1, ossification
centers were found to strongly express HtrA1. The primary
ossification centers expressing HtrA1 mRNA as well as the protein in
the ribs of 14.5 and 15.5 dpc embryos are shown in
Fig. 2, parts D and E,
respectively. This figure also shows the rib perichondorium and periosteum
expressing HtrA1 (Fig.
2D,E). In the hind limb of a 17.5 dpc embryo, significant
HtrA1 expression persisted in tendons and ligaments, but expression
in the forming joints was reduced and weak signals were detected only in the
thin layer of articular surfaces (Fig.
2F). In adult bones, HtrA1 protein seemed to be deposited in the
bone matrix (Fig. 2G,H,I),
where Tgfß1 is known to accumulate at high concentrations
(Kresse and Schönherr,
2001
). Cells in the tendons continued producing HtrA1 protein
(Fig. 2H). No significant HtrA1
protein was detectable in the articular cartilage
(Fig. 2G,I) or in the
chondrocytes of the growth plate (Fig.
2G,H). However, when mice were challenged with an anti-collagen
antibody cocktail in order to induce experimental arthritis, articular
chondrocytes, especially those in the deep layer, started producing high
levels of HtrA1 (A.T., unpublished).
The observed spatio-temporal expression patterns of HtrA1 strongly suggest that HtrA1 regulates signaling of Tgfß proteins.
Binding of HtrA1 to Tgfß family proteins
In order to confirm the involvement of HtrA1 in Tgfß signaling, we
examined the binding activity of HtrA1 to Tgfß family proteins. The
mature forms of mouse Tgfß1, Tgfß2, Bmp4, Gdf5 or activin were
expressed as GST-fusion proteins in E. coli. HtrA1 protein was
expressed in a baculovirus expression system. First, we tested the binding of
follistatin to these Tgfß proteins. Follistatin bound most strongly to
activin, weakly to Bmp4, but not to any other proteins
(Fig. 3A,a). This binding
specificity of follistatin was consistent with previous reports
(Iemura et al., 1998), thus
confirming the validity of our binding assay. The GST pull-down assay revealed
that HtrA1 protein bound to all Tgfß family proteins tested
(Fig. 3A,b).
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To measure the binding affinity quantitatively, we developed a solid phase
binding assay for HtrA1 (Fig.
3C). Kd of HtrA1 for Bmp4 was estimated to be
1.3 nM. This affinity was comparable with the reported affinities of
follistatin (FS-315) for activin (Kd=430 pM)
(Hashimoto et al., 2000), or
of follistatin (FS-288) for Bmp4 (Kd=23 nM)
(Iemura et al., 1998
).
Inhibition of Bmp4 and Tgfß1 signaling
We next examined whether HtrA1 was able to inhibit signaling mediated by
Tgfß proteins. We assayed the effects of HtrA1 on Bmp2, Bmp4 and
Tgfß1 signaling by transcriptional assay in mouse C2C12 myoblast cells
(Katagiri et al., 1994). For
Bmp signaling we used a reporter plasmid, pGL3-Id985WT, in which a luciferase
gene was driven by a Bmp-responsive promoter element derived from murine
Id gene (Katagiri et al.,
2002
). Transfection of C2C12 cells with the Bmp4
expression vector stimulated the Id promoter several fold, similar to
as described by Pearce et al. (Pearce et
al., 1999
), and cotransfection of Smad1 and
Smad4 expression vectors further stimulated the Id promoter
up to 11-fold (Fig. 4A).
Addition of the HtrA1 expression vector inhibited Bmp4 signaling in a
dose-dependent manner (Fig.
4A). The inhibitory activity of HtrA1 on Bmp4 signaling appeared
to be weaker than that of noggin (Fig.
4A). The same promoter was similarly activated by addition of
purified human recombinant Bmp2 protein to the culture medium. HtrA1 inhibited
Bmp2 signaling to a similar degree as it did Bmp4 signaling (data not
shown).
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The constitutively active Bmp4 receptor, caBMPR-IB, stimulates the
Id promoter in the absence of ligands
(Akiyama et al., 1997;
Katagiri et al., 2002
). HtrA1
did not inhibit the signal that originated from caBMPR-IB
(Fig. 4D). This result excludes
the possibility that HtrA1 inhibits intracellular signaling pathways.
Overexpression of HtrA2, a diverse member of the mammalian HtrA
family, was shown to induce apoptosis (Li
et al., 2002). We assayed cell death by propidium iodide staining
after transfection of C2C12 with the HtrA1 expression vector. The
number of dead cells was not increased by HtrA1 overexpression (data
not shown). This excluded the possibility that HtrA1 overexpression
induces cell death in C2C12.
Domains necessary for binding and inhibition
The domains of HtrA1 protein involved in the binding to and the inhibition
of signaling by Tgfß proteins were determined by producing a series of
HtrA1 mutant proteins. A schematic presentation of the mutants is shown in
Fig. 5. Because wild-type (WT)
HtrA1 protein binds to Bmp4 and Gdf5 with the highest and the lowest
affinities, respectively, we investigated the binding of mutants to these two
ligands. Inhibition on Bmp4 signaling was examined using the
Bmp4/Smad1/Smad4 co-transfection assay. Unexpectedly, we found that
the protease domain and the linker region (aa 155-202) that precedes it, but
not the follistatin-like domain, were essential for both activities.
A mutant (FS) with a deletion in both the IGFBP and KI domains (here
we combine these two domains and refer to them as FS) bound to both Bmp4 and
Gdf5 (Fig. 3B,a) with
affinities equivalent to those of wild-type HtrA1.
FS inhibited Bmp4
signaling quite efficiently (Fig.
4C). The deletion mutant (
linker/SP/PDZ), retaining only
the FS domain, did not show binding activity
(Fig. 3B,c) or inhibitory
activity on Bmp4 signaling (Fig.
4C). The PDZ domain was not involved in the binding; a deletion
mutant in both the FS domain and the PDZ domain (
FS/PDZ) retained
binding activity to Bmp4 and Gdf5, as well as inhibitory activity on Bmp4
signaling (Fig. 3B,b;
Fig. 4C). Results with other
mutants are summarized in Fig.
5. When the N-terminal deletion was extended to the linker region
(
FS/linker), binding and inhibitory activities were both diminished.
The mutant (
SP/PDZ) lacking the SP domain was completely devoid of
binding and inhibitory activities. These results indicate that the linker and
SP domains are essential for binding to and inhibition of Tgfß
proteins.
We then examined whether serine protease activity was necessary for binding
and inhibition of Tgfß family proteins. The serine residue at position
328 in the catalytic center was substituted with alanine. This mutant, S328A,
exhibited no protease activity (data not shown) (see also
Hu et al., 1998). S328A showed
binding activity with an affinity comparable to that of wild-type HtrA1
(Fig. 3B,d). The serine
protease activity, therefore, is not essential for the binding activity. By
contrast, this mutant was completely inactive with regard to inhibition of
Bmp4 signaling (Fig. 4C).
Taken together, these results indicated that the linker region and the SP
domain are necessary for both binding and inhibitory activities, and that the
serine protease activity is required for the inhibitory activity of HtrA1 on
Tgfß-mediated signaling. We examined the expression levels of all
mutants, except for FS/PDZ, which our antisera did not detect, in the
C2C12 culture media by western blotting (see insets in
Fig. 4C). Although some
variations in the production levels for each mutant were observed, they were
not significant and thus do not affect our conclusion. It was noticeable,
however, that the expression levels obseverved with the mutant
FS in
the C2C12 medium were less than one third of those observed with wild type
(data not shown). Taking into account these expression levels, the inhibitory
activity of
FS may be higher than that of wild type. We also measured
the protease activities of the mutants used in this study (M.Y., unpublished)
and found that protease activity correlates well with signal inhibition.
Inhibition of chick eye development in ovo
In an attempt to determine whether HtrA1 inhibits signaling by Tgfß
proteins in vivo, we investigated the effects of exogenous HtrA1 during in ovo
chick eye differentiation. Another purpose of this experiment was to examine
whether the secreted HtrA1 protein inhibited Tgfß proteins
extracellularly. Both Bmp4 and Bmp7 play important roles in
early stages of lens development (Dudley
et al., 1995; Wawersik et al.,
1999
; Furuta and Hogan,
1998
) (for a review, see Chow
and Lang, 2001
). Noggin misexpression at the optic vesicle was
shown to inhibit endogenous Bmp signaling, severely suppress the development
of the lens and retina, and result in micropthalmia
(Adler and Belecky-Adams,
2002
).
Cultured 293T cells were transiently tranfected with either a mouse HtrA1 expression vector or a control vector expressing the antisense strand of HtrA1 cDNA. A suspension of 293T cells was injected into the extraocular mesenchyme near the optic vesicle of chick embryos at stage (St.) 9-10. A GFP expression vector was co-transfected in order to monitor the injected 293T cells (see Fig. 6A,J). At St. 23-24, eye morphology of the embryo was examined. Twenty-three percent of chick embryos that were injected with HtrA1-expressing 293T cells had smaller lenses in the injected side than in the untreated side (n=43) (Fig. 6B,C), whereas none of the chick embryos that were injected with the control 293T cells exhibited this phenotype (n=25). As for the retina, 33% of embryos injected with HtrA1-expressing cells had smaller pigmented retinas (Fig. 6B,C). On the contrary, only 8% of embryos injected with the control 293T cells showed smaller pigmented retinas than the control side, and this may be due to physical damage caused by the injection procedure. When noggin-expressing 293T cells were injected, 80% of embryos developed smaller lenses and 87% of embryos showed smaller pigmented retinas (n=15; Fig. 6K,L). The effects of HtrA1 were thus apparently weaker than those of noggin. To estimate the potency of HtrA1, we diluted noggin-expressing cells with the control 293T cells. An approximate 3- to 10-fold dilution of noggin-expressing cells gave rise to the same degree of inhibition as HtrA1-expressing cells (data not shown), suggesting that the potency of HtrA1 is several-fold lower than that of noggin.
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Retinal development was also abnormal in HtrA1- or noggin-treated eyes; the
neural retina was not arranged in the correct layers, was detached from the
retinal pigment epithelium (Fig.
6E,I,N,R) and, in some cases, retinal cells were aggregated (data
not shown). In the ventral optic cup of the HtrA1-treated eye, a
neuroepithelium-like tissue replaced the normal retinal pigment epithelium
(Fig. 6H,I). This abnormal
transition of epithelial cell types was observed in noggin-treated eyes in our
experiment (Fig. 6Q,R) and was
also reported previously (Adler and
Belecky-Adams, 2002).
In conclusion, all abnormalities observed in the HtrA1-treated eyes were indistinguishable from those caused by noggin, but differed in severity. This result strongly suggests that secreted HtrA1 is able to extracellularly inhibit Bmp signaling in vivo.
Induction of Vax expression by HtrA1 overexpression
We further examined whether expression of Vax was affected by
HtrA1 misexpression in vivo. Expression of Vax, which
ventralizes the retina, is restricted to the ventral half of the retina,
whereas expression of Tbx5, which dorsalizes the retina, is
restricted to the dorsal half. Bmp4 signaling regulates the determination of
dorsoventral polarity in the neural retina by regulating the expression of
these two transcription factor genes (Schutle et al., 1999;
Koshiba-Takeuchi et al.,
2000). Overexpression of a Bmp antagonist, like noggin or
ventroptin, leads to upregulation of Vax expression and
downregulation of Tbx5 expression
(Koshiba-Takeuchi et al.,
2000
; Sakuta et al.,
2001
). We electroporated an HtrA1 expression vector into
the optic vesicle at St. 10-11 and examined Vax expression after
incubation for 24-28 hours at St. 17-18. Expression of Vax was
restricted to the ventral half of the retina in the untreated eye
(Fig. 6T). However, in the
retina electroporated with the HtrA1 expression vector, Vax
expression was upregulated in the ventral half and expanded into the dorsal
half (Fig. 6U). Electroporation
of a GFP expression vector alone had no effect on Vax
expression (data not shown). These data indicate that HtrA1
misexpression ventralizes the retina by inhibiting Bmp4 signaling. Taken
together, the results obtained from in ovo experiments showed that HtrA1 is a
novel inhibitor that attenuates Bmp signaling extracellularly.
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Discussion |
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Expression of HtrA1 during morphogenesis
In the brain, HtrA1 expression was observed in regions where the
choroid plexus arises from the neuroepithelium. The importance of the Bmp
signaling in choroid plexus development was recently demonstrated in a study
using conditional knockout mouse of the type I Bmp receptor (Bmpr1a)
gene; disruption of Bmpr1a in the telencephalon led to defects in
choroid plexus formation (Hebert et al.,
2002).
In the developing heart and gonad, expression of HtrA1 seems to be closely
associated with EMT, a process regulated by Tgfß signaling. The
relationship between HtrA1 and EMT was also proposed from several studies on
differentially expressed genes in malignant cancers. Downregulation of
HtrA1 expression has been repeatedly observed during the late stages
of progression or metastasis of melanomas
(Baldi et al., 2002) and
ovarian cancers (Shridhar et al.,
2002
). The process of cancer invasion and metastasis has been
shown to recapitulate EMT, and, in certain types of cancer, is enhanced by
Tgfß signaling (Moustakas et al.,
2002
). Therefore, it is plausible that HtrA1 suppresses EMT of
cancer cells by antagonizing Tgfß signaling.
During the development of bones and their appending tissues, HtrA1 is
likely to suppress chondrocyte differentiation and divert the fate of
mesenchyme to non-chondral types, such as periosteum, perichondrium, joints,
tendons and ligaments. This is consistent with the fact that Bmps and
Tgfßs enhance differentiation of chondrocytes from mesenchymal cells
(Capdevila and Belmonte, 2001).
Development of tendons and ligaments also depend on signaling by Bmp/Gdf
proteins. Bmp signaling seems to specify regions from which tendon progenitors
arise, whereas additional signals, for example Gdf6, Gdf7 and Tgfß2, may
be required for further maturation of tendons
(Schweitzer et al., 2001
;
Wolfman et al., 1997
;
Merino et al., 1998
). Because
HtrA1 is expressed in tendons at later stages of development, it is possible
that HtrA1 may regulate the maturation of tendons.
HtrA1 is expressed in joint interzones and may regulate joint development,
a process closely regulated by Bmp/Gdf signaling
(Brunet et al., 1998;
Francis-West et al., 1999
). It
has also been proposed that signaling mediated by the Tgfß subfamily
participates in the maintenance of adult joints. Tgfß is required to keep
articular chondrocytes in their normal, undifferentiated state. Transgenic
mice carrying a dominant-negative, kinase-defective Tgfß type II receptor
gene (Serra et al., 1997
), or
Smad3 knockout mice (Yang et al.,
2001
), showed phenotypes very similar to human osteoarthritis. In
these mice, suppression of Tgfß signaling probably promoted terminal
differentiation of articular chondrocytes, leading to osteoarthritis-like
phenotypes. Interestingly, expression of human HTRA1 increased
substantially in articular cartilage cells of patients with osteoarthritis
(Hu et al., 1998
). It is
possible that HtrA1 antagonizes Tgfß and aggravates osteoarthritis.
Functional domains of HtrA1
Unexpectedly, we found that the linker region and the SP domain, but none
of the IGFBP and KI (or follistain-like) domains, were required for HtrA1
binding to Tgfß family members.
The linker regions are highly conserved among the four HtrA members
(Li et al., 2002). The linker
region of HtrA2 is essential for trimer formation. This suggests that HtrA1
may form a homotrimer, similar to HtrA2. The binding activity of each HtrA1
mutant to Tgfß family proteins seems to closely correlate with its
ability to form a possible trimeric structure. For example, the mutant
FS/linker, which lacked the linker region, was incapable of binding to
Tgfß family proteins. By contrast, S328A, which did not possess protease
activity but was possibly able to form the trimer
(Li et al., 2002
),
demonstrated binding activity to Tgfß family proteins, though it lacked
inhibitory activity.
The PDZ domain of HtrA1 was not required for either binding or inhibitory
activity. The PDZ domain of HtrA2 negatively regulates proteolytic activity. A
hypothetical trimeric apoptosis receptor is thought to bind to the PDZ domain
and enhances the proteolytic activity of HtrA2
(Li et al., 2002). The PDZ
domain of HtrA1 may have a similar regulatory function. Indeed, the two
mutants that lacked PDZ domains,
PDZ and
FS/PDZ, seemed to have
stronger inhibitory activity on Tgfß signaling than wild-type HtrA1
(Fig. 5). Our yeast two-hybrid
screening isolated trimeric extracellular matrix (ECM) proteins most
frequently as binding factors for the PDZ domain of HtrA1 (Murwantoko,
unpublished). Tgfß signaling has a close and mutual correlation with the
ECM; production of ECM is regulated by Tgfß signaling and vice versa
(Kresse and Schönherr,
2001
). Secreted HtrA1 may bind to ECM around the cell, become
fully active and, subsequently, regulate Tgfß signaling. The inhibiting
activity of HtrA1 was 3- to 10-fold weaker than that of noggin, based on in
vitro (Fig. 4A) and in vivo
experiments (Fig. 6). One
explanation is that HtrA1 must be bound to and activated by ECM proteins, and
thus it functions fully as a semi-autocrine factor.
The function of the IGFBP and KI domains remains to be elucidated.
FS mutants lacking these domains seemed to have enhanced inhibitory
activity on Tgfß signaling (see Fig.
5 and Results). The N-terminal FS domain may have another
regulatory function on the protease activity of HtrA1.
The way in which HtrA1 protease activity inhibits signaling by Tgfß proteins has yet to be clarified. During the GST pull-down assay we did not detect any degradation of GST-Tgfß proteins by HtrA1. Furthermore, untagged human recombinant Tgfß1 and Bmp2, as well as myc-tagged Bmp4 produced by 293T cells, were all resistant to HtrA1 (data not shown). This does not exclude the possible degradation of Tgfß proteins in vivo, because degradation of Tgfß proteins may require cofactors, such as ECM proteins that bind to the PDZ domain. Alternatively, HtrA1 may degrade the receptors for Tgfß family proteins or the extracellular proteins that positively regulate Tgfß signaling. In these cases, the specific and high-affinity binding of HtrA1 to Tgfß may bring HtrA1 into close vicinity of the substrate proteins, and may thus facilitate efficient degradation.
Another possibility we must consider is that binding to Tgfß proteins is irrelevant to the apparent inhibition of their signaling. So far, we have not obtained evidence indicating that binding is essential for the inhibition of Tgfß signaling. It is possible that HtrA1 digests ECM, indirectly inhibiting Tgfß signaling. In fact, we found that major proteoglycans of ECM, decorin and biglycan, were digested by HtrA1 in vitro (M.Y., unpublished). These two proteoglycans bind to Tgfßs, and at least decorin can modulate physiological signaling of Tgfßs. In order to understand the biological functions of HtrA1, it will first be necessary to identify the physiological substrates of this interesting serine protease.
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ACKNOWLEDGMENTS |
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