1 Diabetes Center, Department of Medicine, University of California, San
Francisco, CA 94143, USA
2 Department of Molecular and Cellular Biology, Harvard University, Cambridge,
MA 02138, USA
3 Cardiovascular Research Institute, University of California, San Francisco, CA
94143, USA
* Author for correspondence (e-mail: mhebrok{at}diabetes.ucsf.edu)
Accepted 6 June 2003
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SUMMARY |
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Key words: Pancreas, Islets, Hedgehog signaling, Hhip, Patched
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Introduction |
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Regulation of Hh signaling activity is essential for embryogenesis as
different levels of hedgehog signaling have been shown to specify different
cell types. In the neural tube, specification of ventral cell types is
regulated via graded levels of Hh signaling
(Chiang et al., 1996;
Ericson et al., 1997
;
Marti et al., 1995
;
Roelink et al., 1995
).
Extensive feedback mechanisms have evolved to ensure that hedgehog gradients
are established within responding tissues, in part through the expression of
cell surface proteins that bind and thus limit the diffusion of Hh ligands.
All three Hh proteins bind to Ptch (Marigo
et al., 1996
; Stone et al.,
1996
), a transmembrane receptor expressed in hedgehog target cells
(Goodrich et al., 1996
).
Ptch is a transcriptional target of hedgehog signaling
(Ingham and McMahon, 2001
) and
by increasing the level of Ptch protein in responding cells, Hh signaling
attenuates its own activity in a negative feedback loop. Similarly, another
hedgehog binding protein, Hhip (previously known as Hip1), has been identified
(Chuang and McMahon, 1999
).
Biochemical studies have shown that Hhip binds to all three Hh
ligands with affinities similar to Ptch
(Chuang and McMahon, 1999
).
Ectopic expression of Hhip in transgenic animals inhibits hedgehog
function (Chuang and McMahon,
1999
; Treier et al.,
2001
), while loss of Hhip function results in increased Hh
signaling (Chuang et al.,
2003
). Combined activities of Hhip and Ptch have been reported
during lung development (Chuang et al.,
2003
), demonstrating that the function of these regulatory
proteins during normal embryogenesis is to restrict overt Hh signaling by
sequestering of Hh ligands.
We have tested the requirement of Hhip and Ptch during pancreas formation. We show that Hhip is co-expressed with Ptch at high levels in stomach and duodenum, while only low levels of Hhip expression are detectable in pancreatic tissue. Targeted deletion of Hhip results in increased hedgehog signaling activity in pancreatic tissue, reduction of pancreas mass, endocrine cell numbers and smaller islets of Langerhans. The additional loss of one Ptch allele in Hhip/ embryos further compromises pancreas morphogenesis and endocrine cell differentiation, demonstrating that Hhip and Ptch function jointly to restrict hedgehog signaling activity during pancreas organogenesis. Thus, as it has been shown for other tissues, pancreatic tissue responds to hedgehog signaling in a dose-dependent manner.
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Materials and methods |
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Tissue preparation, immunohistochemistry and microscopy
The lower trunk of E12.5 (embryonic day 12.5) embryos or isolated pancreas
at E18.5 were fixed in 4% (w/v) paraformaldehyde (PFA) in phosphate-buffered
saline (PBS) for 1 or 4 hours at 4°C, respectively. Histological analysis,
quantification of the tissue area and counting of cells were performed as
previously described (Hebrok et al.,
2000). Hematoxylin/Eosin staining, immunohistochemical and
immunofluorescence analyses were performed on paraffin wax-embedded sections
as described previously (Kim et al.,
1997
). The following primary antibodies were used: guinea pig
anti-insulin diluted 1:500 (Linco); rabbit anti-glucagon diluted 1:500
(Linco); rabbit Pdx1 diluted 1:3000 (gift from Dr Michael German); rabbit
anti-Glut2 diluted 1:1000 (Chemicon); rabbit anti-amylase diluted 1:750
(Sigma); mouse Cy3-conjugated anti-
-smooth muscle actin diluted 1:200
(Sigma); rabbit anti-Ki-67 diluted 1:200 (Novocastra Laboratories) and mouse
anti-Isl-1 diluted 1:100 (Developmental Studies Hybridoma Bank).
For immunohistochemistry, the biotinylated anti-mouse IgG (Vector) was used as secondary antibodies at a 1:200 dilution. Staining for diaminobenzidine (DAB) was performed with the ABC Elite immunoperoxidase system (Vector). The following secondary antibodies were used for immunofluorescence: FITC-conjugated anti-guinea pig 1:750 (Molecular Probes), Cy3-conjugated anti-rabbit diluted 1:750 (Molecular Probes). Fluorescence was visualized and photographed with a Zeiss Axiphoto2 plus microscope.
Staining for ß-galactosidase activity and whole mount in situ
hybridization
Upper abdominal organs isolated from heterozygous Hhip mutant
mice, in which the lacZ gene was inserted into the Hhip
locus, were fixed for 15 minutes at 4°C in 4% PFA and then incubated
overnight in phosphate-buffered saline (PBS) supplemented with
5-bromo-4-chloro-3-indolyl-D-galactopyranoside (X-gal; 400 µg/ml) at
4°C. For whole-mount in situ hybridyzation, gastrointestinal tract
including lung, stomach, pancreas and duodenum were dissected and fixed in 4%
PFA overnight at 4°C. Whole-mount in situ hybridization with
digoxigenin-labeled Fgf10 riboprobe was performed as previously
described (Chuang et al.,
2003). Stained tissues were photographed on a Leica MZ FL3
equipped with a Leica IM500 system.
RNA preparation and RT-PCR analysis
Dissected embryonic pancreas rudiments were dissolved in Trizol (Gibco-BRL)
and total RNA was prepared according to the manufacturer's methods. RT-PCR was
performed as described elsewhere (Wilson
and Melton, 1994). Hhip PCR was performed under the
following conditions: 1 cycle of 94°C for 2 minutes; 60°C for 1
minute; 72°C for 1.5 minutes followed by 35 cycles of 94°C for 1
minute; 60°C for 1 minute; 72°C for 1.5 minutes. Mouse ribosomal
protein L19 was used as the internal control. Forward and reverse primer
sequences used are listed 5' to 3'.
Hhip: AATTGCCAAGTGTGAGCCAG and TGCCCACTGGAAAGATAGAC
L19: CTGAAGGTCAAAGGGAATGTG and GGACAGAGTCTTGATGATCTC
SYBR Green real-time quantitative PCR
PCR amplifications were performed using an ABI Prism 7700 sequence
detection system (Applied Biosystems, Foster City, CA). Reactions were
performed in a reaction mixture consisting of a 50 µl volume solution
containing 1xSYBR Green PCR master mix (Applied Biosystems) and 300 nM
of each primer. Amplification was performed by initial polymerase activation
for 10 minutes at 95°C, and 40 cycles of denaturation at 94°C for 30
seconds, annealing at 58°C for 40 seconds and elongation for 1 minute at
72°C. To exclude contamination with nonspecific PCR products such as
primer dimers, melting curve analysis was applied to all final PCR products
after the cycling protocol. Forward and reverse primer sequences used are
listed 5' to 3'.
Actin: ATGACGATATCGCTGCGCTGGT and ATAGGAGTCCTTCTGACCCATTCC
Gli: TTGTCCAGCTTGGATGAAGG and CCCAGACGGCGAGACAC
Morphometric quantification of pancreatic tissues and cell
counting
To obtain representative results, the whole pancreas was used for
quantification. The first three or five consecutive sections of E12.5 or E18.5
pancreatic tissue, respectively, were mounted on the first of a series of
three or five microscope slides, followed by the next three or five sections
placed on the second slide. A total of three individual slides (1a-3a) were
filled with consecutive sections for E12.5 embryos, and a total of five
individual slides (1a-5a) for E18.5 specimen. When necessary, additional
series of three (1b-3b, etc) or five (1b-5b, etc) slides were prepared until
all pancreatic sections were mounted.
After Hematoxylin/Eosin staining or immunohistochemistry, pancreatic epithelial areas were outlined and measured with the OpenLab software. Insulin- and glucagons-positive areas of the E18.5 pancreas were measured on every 25th section (every 150 µm) from one set of slides (1a-1e). Ki-67-, Isl1-, insulin- and glucagons-positive cells were counted using one series of the slides at E12.5 or E18.5 (e.g. 1a-c). The widest region of the posterior stomach at E12.5 was used to measure thickness of epithelium or mesenchyme. Data analysis was performed with Minitab (version 13, State College, PA). Statistical significance was assessed by employing Mann-Whitney test, except the result of Isl1-expressing cell numbers in Fig. 6L, which was analyzed using Student's t-test because of similar standard deviations.
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Results |
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To understand if increased hedgehog signaling influences ß-cell differentiation, we examined the expression of mature ß-cell markers, including Pdx1 and glucose transporter 2 (Glut2; Slc2a2 Mouse Genome Informatics), a low-affinity transporter present in the plasma membrane of pancreatic ß-cells, in Hhip mutant embryos (Fig. 4F-I). Expression of these markers is unchanged in Hhip mutant ß-cells, indicating that elevated hedgehog signaling impairs ß-cell proliferation but not differentiation.
Fgf10 expressions in pancreatic mesenchyme are affected in
Hhip mutants
The relative loss of pancreatic mass and endocrine cell numbers is
reminiscent of defects reported in mice lacking Fgf10
(Bhushan et al., 2001).
Transient expression of Fgf10 in pancreatic mesenchyme between E9.5
to E11.5 is essential for proliferation of Pdx1-positive epithelial progenitor
cells. We performed in situ hybridization to test if pancreatic Fgf10
expression is affected by increased hedgehog signaling in Hhip
homozygous mutants. Although Fgf10 expression is rapidly detectable
in control pancreatic and lung buds at E10.5, Fgf10 transcripts are
significantly reduced in Hhip/ tissue
(Fig. 5A,B). However,
Fgf10 expression is not completely abolished, as similar levels of
expression are detected in control and Hhip mutant pancreas at E11.5
(Fig. 5C,D). The partial
reduction of Fgf10 expression is likely to account for some of the
pancreatic defects observed in Hhip mutant mice.
|
Defects in organ formation at the fore-midgut area in
Hhip/Ptch+/
mutants
In addition to defects in pancreas development,
Hhip/ mutants also showed structural
abnormalities in stomach and spleen (Figs
2,
7)
(Apelqvist et al., 1997). In
control embryos, wild-type and Hhip+/, the proximal
third of the stomach (fore-stomach) is lined by a squamous epithelium, while
the distal two-thirds are covered by a glandular epithelium. In Hhip
mutants, there was a tendency that the thickness of the posterior epithelium
was reduced. Further reduction was observed in
Hhip/Ptch+/
embryos, indicating that combined activities of Hhip and
Ptch are required for proper stomach development. In contrast to the
epithelial reduction, a stepwise increase in posterior stomach mesenchyme was
noted with decreasing levels of Hhip and Ptch
(Fig. 7). These morphological
changes are accompanied by molecular changes in marker gene expression, as
demonstrated by changes in Isl1 expression in stomach mesenchyme.
Isl1 is normally found in pancreatic and posterior stomach mesenchyme
but is excluded from anterior stomach mesenchyme
(Kim et al., 2000
). In
Hhip/ embryos, and even more pronounced in
Hhip/Ptch+/
embryos, the expression in posterior stomach is reduced
(Fig. 7A-C), suggesting that
stepwise increase of Hh signaling leads to a gradual change in cell
differentiation.
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Discussion |
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Embryos carrying homozygous deletions of the Hhip gene develop
without any obvious changes in gross morphology but newborn mice die shortly
after birth because of deficiencies in lung development that result in
respiratory failure (Chuang et al.,
2003). Closer inspection of the developing lungs revealed elevated
Hh levels as demonstrated by increased expression of the Hh target genes,
Ptch and Hhip (elevated levels of Hhip were
measured by comparing the intensities of ß-galactosidase staining in
Hhip+/ and Hhip/
mutants). Similarly, Hhip also regulates hedgehog signaling in the
pancreas anlage as the expression levels of the lacZ gene knocked
into the Hhip locus was substantially higher in stomach and duodenum
of E12.5 Hhip/ embryos compared with that in
Hhip+/ mutant embryos. The increase in
ß-galactosidase activity was significant, even when the differences in
gene dosage of the lacZ gene in heterozygous and homozygous mutants
was taken into account (Fig.
2A,B). Quantitative measurement of the hedgehog signaling activity
with `Real time' PCR detection of Gli transcripts revealed an
approximately eightfold increase in homozygous Hhip mutants
(Fig. 2C). The approximately
threefold increase observed in heterozygous mutants indicates that loss of one
Hhip allele is sufficient to substantially increase Hh signaling
activity. Thus, these data demonstrate that Hhip functions as an
inhibitor of Hh signaling within the developing organs of the fore-midgut
region, including the pancreas proper.
Loss of Hhip function results in obvious changes in organ
morphology in the pancreas anlage. These defects are reminiscent of, but more
moderate than, changes found in transgenic mice ectopically expressing sonic
hedgehog under control of the Pdx1 promoter (Pdx-Shh)
(Apelqvist et al., 1997).
Although the spleen, an organ derived from posterior stomach mesenchyme, is
missing in Pdx-Shh mice, it is misshapen and reduced in size in
Hhip/ embryos. In Pdx-Shh
transgenics pancreatic mesenchyme transforms into duodenal mesoderm and
clusters of pancreatic cells are dispersed in duodenal tissue. In
Hhip mutants, ventral pancreatic tissue extends laterally towards the
duodenum and in some cases small patches of ectopic pancreas are found within
the gut (Fig. 2D-K). The border
between the dorsal and ventral pancreas is diminished, leading to a contiguous
mass of pancreatic tissue in severe cases
(Fig. 3F,G). However, smooth
muscle-like structures, indicative of transformation of pancreatic into
duodenal tissue, were not observed in the pancreatic region of
Hhip/ mutants
(Fig. 3E,F). Although the
exocrine component appears normal, islet mass and architecture is
significantly affected. Endocrine cells leave the pancreatic epithelium and
aggregate in cell clusters but the total number of endocrine cells is reduced
by 45%, a decrease predominately caused by the loss of larger islets
(Fig. 4D). Thus, loss of
Hhip function results in specific pancreatic phenotypes that are
similar to, but less severe than, those observed in Pdx-Shh mice.
Level of hedgehog signaling regulates pancreas development
A possible explanation for the milder phenotype is that loss of
Hhip might not increase hedgehog signaling to the level observed in
Pdx-Shh transgenic mice. In addition to Hhip, Ptch, another
attenuator of hedgehog signaling, is also expressed in pancreatic tissue and
inactivation of Ptch has previously been shown to impair pancreatic
marker expression (Hebrok et al.,
2000). Thus, it is likely that loss of Hhip function is
partially compensated for by the remaining function of Ptch. This
hypothesis is supported by our studies of pancreas development in
Hhip/;Ptch+/
mice. Although the analysis of these compound mutant embryos is restricted to
early developmental stages because of lethality before E13, pancreas
development is more severely compromised compared with
Hhip/ mutants. Progressive loss of
Hhip and Ptch alleles results in reduction of pancreas
epithelium and endocrine marker genes in a graded fashion
(Fig. 6). These results suggest
that Hhip and Ptch function jointly during pancreas
organogenesis. They also indicate that Hh signaling effects in the pancreas
are concentration dependent, an observation that has previously been noted
during the development of other organs, including limb bud and neural tube
formation (Ericson et al.,
1997
).
Hhip function regulates endocrine cell mass
Our data indicate that moderate elevation of hedgehog signaling in the
pancreas anlage interferes with proper organ formation
(Apelqvist et al., 1997;
Hebrok et al., 1998
). General
loss of Hhip and reduction of Ptch affects endocrine cell
development and organ morphogenesis at early stages of pancreas organogenesis.
In addition, proliferation of ß-cells in E18.5
Hhip/ embryos is significantly reduced,
suggesting that elevated Hh levels throughout development are sufficient to
impair expansion of ß-cells. By contrast, treatment of established
ß-cell lines with hedgehog agonists has been shown to activate
Pdx1 and insulin gene transcription and to stimulate insulin
secretion (Thomas et al.,
2001
; Thomas et al.,
2000
), suggesting a requirement of Hh signaling in adult pancreas.
Differences between developing and fully matured ß-cells with regard to
their response to Hh signaling could explain these discrepancies.
Unfortunately, Hhip/,
Ptch/ and
Hhip/;Ptch+/
mutants die early during embryogenesis or shortly after birth
(Chuang et al., 2003
;
Goodrich et al., 1997
),
thereby preventing the analysis of their function in the adult pancreas. To
address unequivocally the question of whether ectopic activation of this
pathway affects glucose homeostasis in vivo, spatial and temporal deregulation
of Hh signaling exclusively in the mature pancreas would be required.
Interaction between hedgehog and Fgf signaling pathways
How does elevated hedgehog signaling affect pancreas morphogenesis and
endocrine cell proliferation? Previous studies have shown that hedgehog
signaling attenuates Fgf signaling during lung development
(Bellusci et al., 1997;
Litingtung et al., 1998
;
Pepicelli et al., 1998
). Sonic
hedgehog-mediated repression of Fgf10 expression within localized
areas of the lung mesenchyme is required for proper branching morphogenesis.
Uniform elevation of sonic hedgehog activity in
Hhip/ lung epithelium results in general
inhibition of Fgf10 expression and disruption of secondary branching
(Chuang et al., 2003
). Our
results suggest a similar relationship between hedgehog and Fgf10 signaling
during pancreas development (Fig.
5). Loss of Fgf10 function reduces proliferation of
Pdx1-positive epithelial cells, thereby impairing pancreas growth and
expansion of endocrine cell types (Bhushan
et al., 2001
). Pancreatic defects in Hhip homozygous
mutant mice are less severe than those found in Fgf10 mutant embryos,
a difference that might be explained by residual Fgf10 activity that
is maintained in Hhip mutants
(Fig. 5). Nonetheless, these
data suggest that pancreatic organ size and endocrine cell proliferation could
potentially be controlled by the interaction between hedgehog and Fgf
signaling during early stages of pancreas formation. Inhibition of Fgf
signaling mediated through FGF receptor 1c has been shown to impair mature
ß-cell function (Hart et al.,
2000
). Hedgehog signaling is active in adult islets and future
studies will address if this activity controls expression of other Fgf ligands
within islets and if interactions between these pathways affect mature
endocrine functions.
Combined activities of Hhip and Ptch regulate
stomach development
Hedgehog signaling is essential for proper development of other intestinal
organs within the pancreas anlage, including stomach and spleen
(Harmon et al., 2002;
Ramalho-Santos et al., 2000
;
Sukegawa et al., 2000
). The
mammalian stomach is patterned along its AP axis and posterior stomach
contains mucin-negative vacuoles in columnar epithelium, adjacent to a thick
mesenchymal layer, whereas anterior stomach epithelium is normally squamous
and nonvacuolated (Larsson et al.,
1996
). During embryogenesis, Shh becomes restricted to
the anterior stomach epithelium, while Ihh is mainly expressed in the
posterior compartment (Aubin et al.,
2002
; Bitgood and McMahon,
1995
; Ramalho-Santos et al.,
2000
). In addition, Hhip and Ptch are found
within the surrounding stomach mesenchyme
(Fig. 1)
(Ramalho-Santos et al., 2000
),
indicating that Hh signaling is active in these tissues. Loss of Shh
leads to stomach epithelial overgrowth and increased numbers of glucagon
producing endocrine cells (Hebrok et al.,
2000
; Ramalho-Santos et al.,
2000
). By contrast, reduction of activin signaling or loss of the
transcription factor Hox5a, disturbs mesenchymal-epithelial
patterning of posterior stomach regions, most probably because of an increase
in Hh signaling (Aubin et al.,
2002
; Kim et al.,
2000
). Our results support this hypothesis as we observe more
severe defects in posterior stomach morphology and marker expression in
correlation with the progressive loss of Hhip and Ptch
alleles (Fig. 7). Posterior
epithelial thickness increases gradually while mesenchymal thickness decreases
in response to the additive loss of Hhip and Ptch alleles.
The morphological changes are reflected in the decrease of Isl1 expression in
posterior mesenchyme, indicating a possible anterior transformation of this
tissue. In addition, Hhip/ mutants display a
severe reduction in spleen size (Fig.
2), a phenotype that might be explained by alterations in
epithelial-mesenchymal interactions and marker expression in posterior stomach
mesenchyme.
In summary, this study demonstrates that Hhip and Ptch
act jointly to control Hh signaling within the embryonic fore-midgut region.
One potential implication of these findings is that hypomorphic mutations in
Hh inhibitors could affect adult pancreatic functions. Although adult
heterozygous Ptc1+/ mice display the inability to
maintain glucose homeostasis after injection of a concentrated glucose
solution (Hebrok et al.,
2000), adult Hhip+/ mice are
indistinguishable from wild-type littermates (data not shown). The different
requirement for Ptch and Hhip function during maintenance of
glucose homeostasis is in agreement with the severity of phenotypes in
homozygous mutants. While Hhip/ mice
complete embryogenesis and only die shortly after birth,
Ptch/ embryos die before E10.5
(Goodrich et al., 1997
),
demonstrating different requirements for Ptch and Hhip
function during embryonic development. Future studies might improve our
mechanistic understanding of the different requirements for Ptch and
Hhip in diverse tissues.
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ACKNOWLEDGMENTS |
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