ORIGINAL
RESEARCH
Connective Tissue Growth Factor (CTGF) Inactivation
Leads to Defects in Islet Cell Lineage Allocation and
␤-Cell Proliferation during Embryogenesis
Laura A. Crawford,* Michelle A. Guney,* Young Ah Oh, R. Andrea DeYoung,
David M. Valenzuela, Andrew J. Murphy, George D. Yancopoulos, Karen M. Lyons,
David R. Brigstock, Aris Economides, and Maureen Gannon
Departments of Medicine (Y.A.O., M.G.), Division of Diabetes, Endocrinology, and Metabolism and Molecular
Physiology and Biophysics (L.A.C., M.A.G., M.G.), Vanderbilt University Medical Center, Nashville, Tennessee 37232;
Department of Orthopaedic Surgery (R.A.D., K.M.L., A.E.), David Geffen School of Medicine at UCLA, University of
California, Los Angeles, California 90095; Center for Cell and Vascular Biology (D.R.B.), Children’s Research Institute,
The Ohio State University, Columbus, Ohio 43205; and Regeneron Pharmaceuticals, Inc. (D.M.V., A.J.M., G.D.Y., A.E.),
Tarrytown, New York 10591
The factors necessary for normal pancreatic islet morphogenesis have not been well characterized.
Here we report that connective tissue growth factor (CTGF) is involved in the establishment of
normal islet endocrine cell ratio and architecture. CTGF is a secreted protein known to modulate
several growth factor-signaling pathways including TGF-␤, BMP, and Wnt. Although its role in
pancreatic diseases such as pancreatitis and pancreatic cancer are well documented, a role for
CTGF in normal pancreas development and function has heretofore not been examined. Using a
lacZ-tagged CTGF allele, we describe for the first time the expression pattern of CTGF in the
developing pancreas and the requirement of CTGF for normal islet morphogenesis and embryonic
␤-cell proliferation. CTGF is highly expressed in pancreatic ductal epithelium and vascular endothelium, as well as at lower levels in developing insulin⫹ cells, but becomes down-regulated in
␤-cells soon after birth. Pancreata from CTGF null embryos have an increase in glucagon⫹ cells
with a concomitant decrease in insulin⫹ cells, and show defects in islet morphogenesis. Loss of
CTGF also results in a dramatic decrease in ␤-cell proliferation at late gestation. Unlike CTGF null
embryos, CTGF heterozygotes survive past birth and exhibit a range of islet phenotypes, including an
intermingling of islet cell types, increased number of glucagon⫹ cells, and ␤-cell hypertrophy. (Molecular Endocrinology 23: 324 –336, 2009)
he mouse pancreas is formed from dorsal and ventral evaginations of the posterior foregut endoderm at embryonic d
9.5 (e9.5), which fuse later in development to become a single
organ. As organ development proceeds, the pancreas becomes
comprised of two functionally distinct cell populations: the exocrine pancreas, composed of acinar cell clusters and a ductal
network, which produces and secretes digestive enzymes into
the lumen of the small intestine; and the endocrine pancreas,
which consists of microorgans known as islets of Langerhans,
involved in the maintenance of glucose homeostasis. In both
T
mice and humans, disruption of glucose homeostasis can lead to
diabetes.
Pancreatic islets consist of ␣-, ␤-, ␦-, ␧-, and PP cells (which
secrete glucagon, insulin, somatostatin, ghrelin, and pancreatic
polypeptide, respectively). These endocrine cells are organized
into a characteristic architecture in the mouse, with the insulinproducing ␤-cells located in the islet core, and the other cell types
located at the islet periphery. Previous studies have suggested that
maximal ␤-cell to ␤-cell contacts allow for optimal insulin secretion in response to glucose, and that the proper organization of the
ISSN Print 0888-8809 ISSN Online 1944-9917
Printed in U.S.A.
Copyright © 2009 by The Endocrine Society
doi: 10.1210/me.2008-0045 Received February 4, 2008. Accepted December 24, 2008.
First Published Online January 8, 2009
* L.A.C. and M.A.G. contributed equally to this study.
Abbreviations: BAC, Bacterial artificial chromosome; BHR, bacterial homologous recombination; BMP, bone morphogenetic protein; CTGF, connective tissue growth factor; e9.5,
embryonic d 9.5; ECM, extracellular matrix; ES, embryonic stem; ␤-gal, ␤-galactosidase;
HNF, hepatocyte nuclear factor; IP-GTT, ip glucose tolerance test; P1, postnatal d 1;
PECAM, platelet-endothelial cell adhesion molecule; TBS, Tris-buffered saline; VWC, von
Willebrand factor type C; WT, wild type.
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Mol Endocrinol, March 2009, 23(3):324 –336
endocrine cell types may be critical for communication between
endocrine cell types (1–5).
At e17.5 in the mouse, clusters of endocrine cells lie close to
the pancreatic ducts from which these cells originate (6 – 8). As
endocrine cells are specified and differentiate, they must delaminate from the ductal epithelium. Beginning around e18.5 and
continuing after birth, cells destined to populate the mature islet
organize in the acinar parenchyma to form typical islet clusters.
Cell biological processes involved in islet morphogenesis include
cell adhesion, cell migration, cell sorting, and extracellular matrix (ECM) degradation and remodeling (6, 9 –11). Although all
endocrine cells arise from a common neurogenin 3 (ngn3)-expresssing progenitor cell population, approximately 80% of
these cells differentiate as ␤-cells (12). Relatively little is known
about the specific factors that regulate allocation to the different
endocrine lineages or their subsequent organization into islets.
Connective tissue growth factor (CTGF) is a member of the
CCN family of secreted proteins named for its founding members, cysteine rich 61 (CCN1), CTGF (CCN2), and nephroblastoma overexpressed (Nov/CCN3). CTGF is a modular protein
with four domains encoded by separate exons: an N-terminal
region with homology to IGF-binding proteins and Twisted
Gastrulation, which modulates bone morphogenetic protein
(BMP) signaling (13, 14); a von Willebrand factor repeat, with
similarities to the cysteine repeats (CR) in the BMP antagonist
Chordin (15–17); a thrombospondin type 1 repeat (18); and a
C-terminal domain with similarity to the C terminus of the
protein Slit, which is involved in axon guidance and cell migration (19, 20). This last domain contains a cysteine knot structure, which is also present in other growth factors such as
TGF-␤, platelet-derived growth factor, and nerve growth factor.
CTGF is expressed in a variety of cell and tissue types, including fibroblasts, vascular smooth muscle, neurons, endothelial cells, and various epithelial cell types (21). Although a specific CTGF receptor has not yet been identified, it functionally
interacts with integrins, including ␣v␤3 (22), and elicits biological effects specific to the signaling properties of the integrin
combination expressed in a particular tissue. CTGF interacts
with several growth factor signaling pathways, including BMP,
Wnt, and TGF-␤. The most studied of these interactions is that
between CTGF and the TGF-␤ signaling pathway. CTGF is
produced at high levels in human foreskin fibroblasts in response to TGF-␤ treatment (23), and a TGF-␤/Smad response
element has been identified in the CTGF promoter, suggesting
that CTGF expression is directly regulated by TGF-␤ signaling
(24). CTGF interacts directly with TGF-␤ extracellularly
through its von Willebrand factor repeat, resulting in enhancement of TGF-␤ signaling and a positive feedback loop (17),
whereas it inhibits BMP and Wnt signaling through interactions
with its CR and C-terminal domains, respectively. CTGF inhibits Wnt signaling by binding to the Wnt coreceptor LRP6 (25),
whereas its antagonizing effect on BMP signaling is caused by
prevention of BMP binding to its cognate receptors (17).
In other cell types, CTGF mediates several processes known
to occur during normal islet morphogenesis, including cell adhesion and migration (26) and ECM production and degradation (27, 28). CTGF localization and function in the developing
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mouse pancreas have never been described, although CTGF has
been linked to cell survival and proliferation in pancreatic cancer, and is up-regulated in chronic pancreatitis (29, 30). Despite
the strong evidence that CTGF plays a role in disease states and
in cell lines, very little is known about its role in developmental
processes. CTGF is critical for normal chondrogenesis and angiogenesis during skeletal development but its role in other tissues has not yet been examined (31). CTGFneo null mutant mice
die immediately after birth due to respiratory failure. To date,
nothing is known about the normal role of CTGF in the developing pancreas.
We recently found that CTGF expression was down-regulated in a mouse model of islet dysmorphogenesis (32). Here we
examined the expression of CTGF during pancreas development and studied the effects of CTGF inactivation on endocrine
cell development, islet morphogenesis, and function. This study
presents the generation and characterization of a CTGFlacZ allele and is the first to use this mouse to characterize CTGF as a
factor critical for proper endocrine lineage allocation, islet morphogenesis, and embryonic ␤-cell proliferation.
Results
CTGF is expressed in the developing mouse pancreas
Previous studies in our laboratory detected a 2-fold decrease in
CTGF expression in a mouse model of islet dysmorphogenesis
using microarray analysis (32). This prompted us to examine the
expression of CTGF in the normal developing pancreas. Antibody labeling of embryonic pancreas at e18.5, a time when islet
morphogenesis is occurring, showed CTGF expression in the
core of the islets where ␤-cells are located. CTGF was not expressed in ␣-cells at this or any stage of endocrine development
examined (Fig. 1A and data not shown). CTGF was still present
in islets at postnatal d 1 (P1), but was not detectable in the
endocrine pancreas at P3 (data not shown), and remained undetectable in adult islets (Fig. 1B). In addition to expression in
␤-cells, CTGF was localized to structures consistent with ducts
and blood vessels in the developing pancreas. The ductal/vascular expression remained detectable in adult pancreas (Fig. 1C).
Generation and characterization of CTGFlacZ allele
To generate a lacZ-tagged CTGF null allele, the coding sequence of CTGF from the start of its von Willebrand factor type
C domain in exon 3 to the end of the coding sequence in exon 5
was replaced with a transmembrane domain-lacZ cassette (Fig.
2; see Materials and Methods for detail). This results in the
production of a CTGF N terminus/␤-galactosidase (␤-gal) fusion protein, the presence of which can be detected at the plasma
membrane with an antibody against ␤-gal (data not shown).
X-gal staining of CTGFlacZ/⫹ embryos was used to determine
when and where CTGF was expressed in the developing pancreas. These data were compared with the CTGF expression
pattern detected by immunofluorescence described above. X-gal
staining revealed CTGF gene activity in the pancreas as early as
e12.5 (Fig. 3A) and further confirmed the CTGF immunofluorescence results (Fig. 1): CTGFlacZ was strongly expressed
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blood vessels [platelet-endothelial cell adhesion molecule (PECAM)], and lower levels of expression in the ␤-cells (insulin) of
the developing pancreas at e18.5 (Fig. 3, G–I). By P3, CTGF
gene expression, as assessed by ␤-gal activity, was no longer
detected in the endocrine cells of the islets, but was still detectable in the microvessels within the islet, again consistent with
our antibody analyses (data not shown). Endocrine-specific
␤-gal activity remained undetectable in adult animals; however,
CTGFlacZ was reexpressed in the islets of pregnant females at
every stage of pregnancy examined [e7.5, e10.5, e12.5, e13.5,
e15.5, e17.5, e18.5 (supplemental Fig. 1 published as supplemental data on The Endocrine Society’s Journals Online web
site at http://mend.endojournals.org and data not shown)]. This
reexpression during pregnancy was also confirmed with the
CTGF antibody (supplemental Fig. 1).
FIG. 1. CTGF is expressed in the mouse pancreas. A, CTGF (red) was expressed
in the islets, ducts (d), and blood vessels (data not shown) at e18.5. CTGF
expression did not overlap with glucagon (gluc; green). Scale bar, 25 ␮m. B,
Endocrine-specific CTGF expression was extinguished in adult islets. C, CTGF
expression in the ductal network was maintained in the adult pancreas.
within the developing ductal epithelium and blood vessels and
within the endocrine cords (Fig. 3, A–C). X-gal staining also
revealed that within the pancreatic mesenchyme, CTGFlacZ was
broadly expressed at e12.5 but declined as embryogenesis progressed (Fig. 3, A–C). Although ␤-gal activity was detected in
the small intralobular ducts of the acinar clusters, it was not
present in the exocrine cells themselves, even from the earliest
stages (Fig. 3, A–F). Colocalization studies using pancreatic cell
type-specific markers on X-gal-stained tissue confirmed high
levels of CTGF expression within the ducts (cytokeratin) and
CTGF mutant embryos have altered proportions of
differentiated endocrine cells and disrupted islet
morphogenesis
The expression of CTGF in the developing pancreas, and during
islet morphogenesis in particular, suggested a role for CTGF in
endocrine cell development and islet morphogenesis. To test this
hypothesis, we used two models of CTGF inactivation: the previously reported CTGFneo line (31) and the CTGFlacZ mice generated for this study (Fig. 2). The phenotype of CTGFlacZ/lacZ
null mice was indistinguishable from the published CTGF null
phenotype from the CTGFneo line. CTGFlacZ/⫹ mice were viable
and fertile. CTGFlacZ/lacZ animals were found at the expected
Mendelian ratio during embryogenesis, but were not found at
P1 or after due to neonatal lethality. For embryonic studies,
therefore, the two CTGF null lines were used interchangeably.
Because CTGF null animals die at birth, studies of endocrine
cell development and islet morphogenesis in these animals was
limited to embryogenesis. At e18.5, null mutant pancreata
showed increased numbers of glucagon-expressing cells compared with controls (Fig. 4B). Morphometric analysis at this
stage revealed a disrupted endocrine cell composition: islets
from CTGF⫺/⫺ animals had approximately 25% of their endocrine area occupied by glucagon-producing cells (Fig. 4B) compared with only 12% in wild type (WT). The increase in glucagon⫹ cells in islets from null embryos was accompanied by a
decrease in insulin area compared with WT (49% in CTGF⫺/⫺
vs. 70% in WT). This corresponds to an ␣- to ␤-cell ratio of 1:4
in WT and 1:2 in mutant animals. We observed no difference in
average islet size or in the distribution of different size islets in
the CTGF null embryos compared with controls (data not
shown). Alterations in endocrine composition could be detected in CTGF mutant animals as early as e13.5, at which
point there was an abundance of large glucagon clusters in
CTGF mutants exceeding those observed in WT controls (Fig. 5).
Morphometric analyses performed at e15.5 confirmed early alterations in endocrine composition in CTGF⫺/⫺ animals (Fig. 4C). At
this stage, nearly 45% of the endocrine area in CTGF⫺/⫺
pancreata was occupied by glucagon⫹ cells, compared with
23% in WT animals. Insulin⫹ cells occupied 37% of endocrine area in CTGF⫺/⫺ embryos at e15.5 compared with 60%
in WT controls.
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FIG. 2. Generation of the CTGFlacZ allele. Map of the CTGF endogenous locus (A), targeting vector (B), and mutant allele (C) produced by homologous recombination.
Only part of the CTGF gene is shown: exons 1–5 (gray arrows) with 100 bp flanking each end. Blue arrows indicate coding sequence. Red boxes indicate the different
protein domains as predicted by Pfam. The sequence being replaced by TM-lacZ is marked and encompasses the coding sequence of the VWC domain (constituting the
greater part of exon 3) to the Stop codon (in exon 5). The donor fragment used for BHR is shown below the map of the CTGF gene. HB5⬘ and HB3⬘ denote the
homology boxes (50 bp each) used for BHR. Expression of NeoR is driven by two promoters: PGK (for expression in mammalian cells) and EM7 (for expression in E. coli)
and is flanked by loxP sites (blue boxes) on either side. TM-lacZ is followed by an simian virus 40 polyadenylation signal and simian virus 40-derived associated sequence
(65), whereas the NeoR open reading frame is followed by the mouse PGK polyadenylation signal and associated sequence (66). All of these elements are the standard
elements used by Velocigene (56). The replacement afforded into the CTGF BAC by BHR is also translated in an identical manner into the mouse genome during
targeting. Therefore, all the features shown above are those present in the modified CTGF locus in the targeted ES cells. SS, Signal sequence; IB, IGF binding; TSP,
thrombospondin; CT, C terminal; pA, poly A; Neo, Rneomycin resistance. PGK, Phosphoglycerate kinase.
We also examined islet cell composition and morphology in
CTGF heterozygous embryos. Similar to CTGF⫺/⫺ embryos,
morphometric analysis of pancreata from e18.5 animals confirmed that endocrine cell composition was altered in CTGF⫹/⫺
animals compared with WT littermates. Whereas WT endocrine
clusters contained about 12% glucagon⫹ cells, islets from
CTGF⫹/⫺ animals had nearly 30% of their endocrine area occupied by glucagon-producing cells (Fig. 4B). This increase in
glucagon area was accompanied by a decrease in insulin area:
70% in WT islets vs. 51% in CTGF⫹/⫺ islets. The alteration in
insulin and glucagon area did not affect the average islet size in
CTGF⫹/⫺ animals, and histogram analysis showed no difference in the distribution of islet sizes between the two genotypes
(data not shown). At e18.5, both homozygous and heterozygous
CTGF mutants showed similar abnormalities in islet organization, including a mixing of ␣- and ␤-cells (Fig. 6B), and the
presence of small glucagon⫹ clusters lacking insulin altogether
(inset, Fig. 6B). In addition, large glucagon clusters were found
associated with islets (Fig. 6C). Morphometric analysis at e15.5
revealed that CTGF⫹/⫺ islets comprised 37% glucagon-expressing cells compared with 23% in WT controls (Fig. 4C), whereas
insulin-producing cells represented 42% of endocrine area in
CTGF⫹/⫺ in contrast with up to 60% in WT pancreata. Insulin/
glucagon double-positive cells were not observed at any stage
analyzed in CTGF⫹/⫺ or CTGF⫺/⫺ animals. Thus, the phenotypes of heterozygous and homozygous CTGF inactivation were
very similar, suggesting haploinsufficiency for CTGF during islet development.
CTGF null animals displayed additional defects in pancreas
development and islet morphogenesis. Cytokeratin (a pan-ker-
atin marker of ductal structures) and synaptophysin (a general
marker of endocrine tissue) colabeling of WT and mutant pancreata revealed that endocrine clusters in CTGF⫺/⫺ animals
were very closely apposed to the ductal epithelium at e18.5 (Fig.
7, compare panels A and C). In addition, the ductal tissue with
which the endocrine clusters were associated in CTGF⫺/⫺ animals had a morphology suggestive of a less mature organ: the
lumen and cross-sectional area were larger, resembling the main
pancreatic duct from which endocrine progenitors normally
arise at an earlier stage in pancreas development, rather than the
smaller ducts detected near islets of the same stage in WT (Fig. 7,
compare panels B and D).
To ensure that transcriptional squelching from the promoter
used to drive the NeoR cassette still present in the CTGFlacZ/⫹
construct was not responsible for the observed phenotype we
observed in these mice, we crossed them to transgenic mice
expressing Cre recombinase under control of a 4.3-kb promoter
fragment of the pancreatic gene, pdx1 (33). This allowed for the
removal of the loxP-flanked NeoR cassette in pancreatic progenitor cells very early in development. The phenotype of pdx1-Cre;
CTGFlacZ/⫹ mice was indistinguishable from the CTGFlacZ/⫹
alone, confirming that the alterations seen in these animals were
due to the loss of CTGF.
CTGF mutant animals have decreased ␤ cell-proliferation
during embryogenesis
To determine the mechanism leading to the altered endocrine
cell distribution, we used morphometric analysis to determine
the percentage of ␣- and ␤-cells undergoing active proliferation
at e13.5, e14.5, e15.5, e16.5, e17.5, and e18.5 in WT and CTGF
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less than 0.1%, respectively, of ␤-cells undergoing
proliferation. We were unable to detect any ␤-cells
undergoing apoptosis in WT or CTGF mutants at
any stage examined. Studies have shown that ␤-cell
proliferation is relatively high at early postnatal
stages and contributes significantly to overall ␤-cell
mass (35). Therefore, we also examined ␤-cell proliferation at P2. Because CTGF⫺/⫺ animals die at
birth, this analysis was restricted to CTGF⫹/⫺ animals. The number of ␤-cells proliferating tended to
be less in CTGF⫹/⫺ animals at P2 compared with
WT animals but did not reach significance (Fig. 8B).
We next examined the expression of the cell cycle
inhibitor p27Kip1. Previous studies have shown an
increase in ␤-cell proliferation in p27 null mutant
pancreata, thus leading to ␤-cell hyperplasia (36).
Furthermore, several studies have shown that enhanced proliferation in response to exogenous
CTGF is due to a reduction in p27. For example,
stimulation of human lung fibroblasts with CTGF
leads to an increase in proliferation concomitant
with a decrease in p27 levels (37). In addition, we
have shown increased p27 protein in ␤-cells in
Foxm1 null mutant pancreata in which postnatal
␤-cell proliferation is dramatically reduced (38).
Western blot analysis revealed an increase in p27 in
both CTGF⫹/⫺ and CTGF⫺/⫺ animals compared
with WT (Fig. 9). These data suggest that reduced
CTGF expression leads to a decrease in ␤-cell proliferation, at least in part, via up-regulation/stabilization of p27.
CTGF heterozygous adults show defects in
islet morphogenesis and proportions of
differentiated endocrine cells
Studies of adult pancreas structure and function
lacZ
lacZ
FIG. 3. Characterization of CTGF
expression. A–F, X-gal staining of CTGF
pancreata at
were limited to CTGF heterozygous animals (CTe12.5 (panel A), e14.5 (panel B), e16.5 (panel C), and e18.5 (panels D–I). CTGFlacZ was
expressed in the developing ductal epithelium (de), and mesenchyme (m), but not in acinar
GFlacZ/⫹ were used exclusively). The size and gross
tissue (a) at e12.5– e16.5. D, At e18.5 CTGFlacZ was expressed in intralobular ducts (d) of the
morphology of CTGF heterozygous pancreata were
exocrine pancreas but not in the majority of acinar cells (a) themselves. E, CTGFlacZ expression in
normal at all ages examined; however, analysis of
pancreatic ducts (d) and adjacent endocrine clusters (e). F, CTGFlacZ was expressed in the
vasculature [artery (art) and vein (v)] of the developing pancreas. G–I, Colocalization of CTGFlacZ
pancreatic tissue from adult CTGFlacZ/⫹ animals rewith cell type-specific markers. CTGF expression was detected by X-gal staining shown in bright
vealed alterations in islet architecture. Some islets
field. Insulin (G*), PECAM (H*), and cytokeratin (I*) immunofluorescence was performed on the
exhibited a morphology indistinguishable from WT
same sections shown in D, E, and F, respectively. Arrows indicate cells expressing CTGF and the
particular marker. Asterisks indicate lacZ⫹ structures that do not express the cell type-specific
islets (Fig. 10, compare panels A and B); however,
marker. Magnification: B and C, ⫻200; A, and D–H, ⫻400; I, ⫻1000.
56% of the islets within the CTGFlacZ/⫹ pancreata
had a mixed-islet phenotype, with ␣-cells found
within the islet core (Fig. 10C). This was not observed in WT
mutant embryos. We observed no difference in the proportion
animals at this age. Additionally, a number of islets were irregof ␣-cells undergoing proliferation between WT or CTGF muularly shaped and appeared to be composed of multiple fused
tant (⫹/⫺ or ⫺/⫺) at any stage analyzed. We also detected no
islets that had not separated during islet morphogenesis (Fig.
differences in ␤-cell proliferation between WT and CTGF mu10D). Morphometric analysis of pancreata from 3-month-old
tants at e13.5, e14.5, e15.5, e16.5, or e17.5; however, there was
WT and CTGF heterozygotes revealed a disrupted endocrine
a significant decrease in ␤-cell proliferation in both CTGF⫹/⫺
cell composition in CTGFlacZ/⫹ animals. In WT animals, only
and CTGF⫺/⫺ embryos at e18.5 compared with WT (Fig. 8A).
We previously showed that ␤-cell proliferation is normally oc5% of the islet area was occupied by ␣-cells, whereas in
curring at this particular time point (34). Whereas in WT panCTGFlacZ/⫹ animals, ␣-cells make up nearly 15% of islet area
creas nearly 1.0% of ␤-cells were proliferating at e18.5,
(Fig. 4A). Despite the increase in ␣-cell area, the area occupied
CTGF⫹/⫺ and CTGF⫺/⫺ animals showed less than 0.2% and
by ␤-cells was not different between WT and CTGFlacZ/⫹ adult
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329
and a slight increase in the number of PP in the CTGF⫹/⫺ animals compared with WT (data not shown).
FIG. 4. CTGF mutant animals have an altered islet composition. Morphometric
analysis of adult (A), e18.5 (B), and e15.5 (C) insulin and glucagon area. A,
Insulin area was unchanged in CTGF⫹/⫺ adults, whereas glucagon area was
increased. At e18.5 (B) and e15.5 (C), insulin area was decreased in CTGF⫹/⫺ and
CTGF⫺/⫺ pancreata, whereas glucagon area was increased. Animals (n ⫽ 5) for
each e18.5 genotype, and animals for each e15.5 and adult genotype (n ⫽ 3)
(see Materials and Methods for full details). Error bars determined by SEM. *, P ⬍
0.01 compared with WT. ␺, P ⬍ 0.05 compared with CTGF⫹/⫺ (Student’s t test).
animals; thus CTGFlacZ/⫹ animals showed an overall larger average islet size compared with WT (WT: 5660 ␮m2 vs. HET:
8609 ␮m2; P ⫽ 0.04). Analysis of the other endocrine cell types
revealed no detectable difference in the proportion of ␦-cells,
CTGF heterozygous adults show compensatory ␤-cell
hypertrophy but no decrease in ␤-cell function
Due to the fact that insulin area was decreased in e18.5 CTGF
heterozygous pancreas, but was not different from WT animals
during adulthood, we hypothesized that there was some compensatory mechanism occurring between e18.5 and adulthood
acting to equalize insulin area in CTGF heterozygous animals.
Analysis of ␤-cell proliferation at 3 months of age revealed no
difference between WT and heterozygous animals (data not
shown). We therefore examined whether individual ␤-cell hypertrophy might contribute to the restoration in insulin⫹ area in adult
islets. Using two different methods to assess cell size (see Materials
and Methods), the ␤-cells from CTGF heterozygotes were statistically larger than ␤-cells in WT animals (Fig. 11), suggesting that
␤-cell hypertrophy contributes to the increase in ␤-cell area that
occurs between e18.5 and 3 months of age. These results do not
exclude the possibility that ␤-cell proliferation also contributes to
the restoration of ␤-cell mass after birth in CTGF heterozygous
animals. Despite the fact that CTGFlacZ/⫹ heterozygous adults had
larger individual ␤-cells, they had a lower total pancreatic insulin
content compared with WT littermates of the same age (83.67
ng/mg pancreas vs. 67.44 ng/mg pancreas; P ⬍ 0.05 at 8 wk of age),
suggesting that that these larger ␤-cells do not have the ability to
produce more insulin than WT ␤-cells.
To determine whether heterozygosity for CTGF resulted in
susceptibility to diabetes under certain conditions, CTGFlacZ/⫹
animals were challenged with 12 wk on high-fat diet. CTGFlacZ/⫹ animals showed normal fasting blood glucose levels and
glucose clearance (data not shown). Thus, by this assay, the
decrease in total pancreatic insulin content in CTGF heterozygous animals does not translate into defects in ␤-cell function.
FIG. 5. Altered lineage allocation in CTGF null pancreata. When compared with controls (A), CTGF null mutant embryos showed no difference in glucagon area (red)
at e12.5 (D). However, an increase in glucagon⫹ cells was seen in CTGF mutant embryos beginning at e13.5 (compare E with B) and continuing throughout embryonic
development. By e15.5. there was clearly a decrease in insulin⫹ cells (green) in the mutant pancreata (compare C and F). Magnification: ⫻ 200 (A, B, D, and E); ⫻ 400
(C and F).
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FIG. 6. CTGF mutant embryos have altered islet architecture. Insulin (green) and
glucagon (red) labeling of islet clusters in WT (A), CTGF⫹/⫺ (B), and CTGF⫺/⫺ (C)
animals at e18.5. B, Increased glucagon area in CTGF⫹/⫺ was due to increased
glucagon clusters associated with insulin-positive cells and with clusters
containing only ␣-cells (inset). C, Increased glucagon area in CTGF⫺/⫺ was due to
large glucagon clusters located within islets. Gluc, Glucagon; ins, insulin.
Discussion
The present study represents the first report on the expression pattern and function of CTGF in the developing pancreas. CTGF
Mol Endocrinol, March 2009, 23(3):324 –336
expression is elevated in pancreatic diseases such as cancer and
pancreatitis and contributes to pancreatic tumor growth and metastasis (30, 39), but before this study, nothing was known about
its potential role in pancreas development and function. Here we
demonstrate that CTGF is expressed in multiple cell types in the
developing pancreas and is involved in endocrine lineage allocation, islet morphogenesis, and embryonic ␤-cell proliferation.
We first became interested in the role of CTGF in the developing pancreas after a microarray analysis revealed a 2-fold
decrease in CTGF expression in a transgenic model of islet dysmorphogenesis and diabetes studied in our laboratory
(pdx1PBHNF6) (32, 40, 41). Recently, the CTGF promoter was
shown to be directly repressed by the proendocrine transcription factor, Ngn3 (42). Ngn3 expression is increased in
pdx1PBHNF6 animals, which may partially account for the decrease in CTGF expression in these transgenic mice. Interestingly, our analysis of CTGF mutant animals showed an islet
phenotype strikingly similar to the hepatocyte nuclear factor
(HNF)6 transgenic phenotype. The increase in ␣-cell number
and mixed islets observed in the CTGF mutant animals at e18.5
is nearly identical to what was described in HNF6 transgenic
animals at the same developmental stage (supplemental Fig. 2
and Ref.39). In addition, HNF6 transgenic pancreata have endocrine tissue closely apposed to ducts, and larger, misshapen
islets, some of which appear to be fused, characteristics also
observed in CTGF null animals (Figs. 7 and 10D). These phenotypic similarities, in conjunction with the fact that CTGF is
down-regulated in HNF6 transgenic pancreata, suggest that the
lack of CTGF may be at least partially responsible for the defects
in islet morphogenesis observed in this mouse model of diabetes.
Based on reports in the literature describing the role of CTGF in
many of the biological processes involved in islet morphogenesis
(cell adhesion, cell migration, and ECM remodeling), and the decrease in CTGF expression observed in a mouse model of islet
dysmorphogenesis, we chose to investigate CTGF as a potential
factor involved in normal islet development. The novel CTGFlacZ
null allele described in this study facilitated studies of CTGF gene
expression and provided important information about the dynamics of CTGF gene activity during normal mouse development.
Morphometric analysis revealed alterations in endocrine cell
composition at e13.5, but not before, in both CTGF heterozygous and homozygous null animals without a concomitant increase in islet size. These data suggest that CTGF functions to
affect endocrine diversification specifically during the secondary
transition. The secondary transition is a wave of differentiation
that begins around e13.5 and continues postnatally (6, 43). The
endocrine cells that arise during this period are thought to contribute to the mature, functional endocrine mass of the animal.
To understand the mechanism(s) leading to this alteration in
endocrine cell distribution, ␣- and ␤-cell proliferation and apoptosis were quantitated at multiple embryonic stages. These
analyses revealed a decrease in ␤-cell proliferation in CTGF
mutants at e18.5, but showed no alterations in ␣-cell proliferation or ␤-cell death at any stage examined. Although a decrease
in embryonic ␤-cell proliferation may explain the decrease in
Mol Endocrinol, March 2009, 23(3):324 –336
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331
The modular nature of the CTGF protein allows it to interact with multiple
growth factor signaling pathways. It is currently unclear whether loss of CTGF in the
developing pancreas primarily impairs
TGF-␤ signaling, results in enhanced BMP
or Wnt signaling, or a combination of all
three. Preliminary studies in our laboratory
indicate that loss of CTGF causes a dose-dependent decrease in membrane-localized
␤-catenin (supplemental Fig. 3A), although
there is no change in total ␤-catenin protein
(supplemental Fig. 3, B and C), providing a
link with the Wnt signaling pathway. Both
chemical and genetic inhibition of TGF-␤ signaling in the pancreas affect islet morphogenesis and pancreatic function (10, 46). The
finding, that inhibition of TGF-␤ signaling
and inactivation of CTGF lead to similar abnormalities in islet development, suggests that
some aspects of the phenotype we observed in
CTGF mutant animals may be due to decreased TGF-␤ signaling. Future studies will
further explore the undoubtedly complex interactions between CTGF, TGF-␤, BMP, and
Wnt signaling in the establishment of normal
FIG. 7. CTGF⫺/⫺ animals have immature duct morphology at e18.5. Cytokeratin (red) and synaptophysin
(green) labeling of e18.5 WT (A and B) and CTGF null (C and D) pancreata. A and B, WT endocrine tissue
islet morphogenesis.
was organized into spherical islet clusters, not directly apposed to ductal structures. C, CTGF null endocrine
Our findings show that CTGF is exclusters remained closely apposed to ducts and did not organize into spherical clusters as in WT (compare
pressed in developing ␤-cells and suggest
C with A). D, CTGF null ducts had a larger lumen and cross-sectional area than WT ducts at e18.5
(compare D with B; also seen in C). synap, Synaptophysin.
that CTGF is important for ␤-cell proliferation during development. This is consistent with reports that CTGF plays a role in cell proliferation in
islet ␤-cell area, the mechanism for the increased number of
other systems (47–50). Until recently, ␤-cells were viewed as a
␣-cells has not yet been uncovered. We hypothesize that the
expanded ␣-cell population may be due to an increased allocanonproliferating cell population. Recent data, however, demonstrate that ␤-cells actually undergo slow turnover in adults and
tion of endocrine progenitors to the ␣-cell fate. Interestingly, the
insulin⫹ cell area normalizes in CTGFlacZ/⫹ islets by 3 months
that under normal circumstances, the majority of new ␤-cells
arise via proliferation of existing ␤-cells (51, 52). There have
of age due to ␤-cell hypertrophy. The hypertrophy seen in CTGF
heterozygous animals in the face of an absolute decrease in
been several factors described as having a specific role in postnatal ␤-cell proliferation, including known cell cycle regulators
␤-cell number is consistent with what we have observed in another mouse model of decreased ␤-cell proliferation (38).
the inactivation of which has no effect on embryonic ␤-cell
During development, CTGF is expressed in pancreatic ducts
proliferation (reviewed in Refs. 43, 53, and 54). Currently, there
and vasculature in addition to ␤-cells. Global inactivation of
are few genes known to affect the proliferation of embryonic
CTGF results in a loss of CTGF from all of the cell types within
␤-cells (34, 55). Efforts in many laboratories are focused on
the pancreas from the very earliest developmental stages. Thereidentifying genes involved in ␤-cell replication, with the hopes of
fore, at this point in our analysis, we are unable to determine
altering/increasing ␤-cell mass in vivo or proliferation of in
which cell type(s) is responsible for the defects in endocrine
vitro-derived ␤-cells to increase insulin production and potenlineage allocation, ␤-cell proliferation, and islet morphogenesis
tially treat diabetes. Because many of these studies aim to direct
the differentiation and proliferation of a precursor or progenitor
displayed in CTGF mutant animals. It remains possible that the
population, it is critical to identify factors that can cause an
endocrine phenotype is secondary to the loss of CTGF from the
expansion of an immature cell population. Because CTGF can
ducts or the vasculature of the developing pancreas. Early panaffect embryonic ␤-cell proliferation, we propose that it is an
creatic endocrine cells delaminate from the ductal epithelium in the
pancreas, and previous studies have shown that blood vessel endointeresting candidate to investigate for its potential in studies
thelial cells are a source of developmental signals promoting endoinvolving expansion of the ␤-cell population. Although hetcrine differentiation during pancreas development (44, 45). Cell
erozygosity for CTGF did not result in high-fat diet-induced
type-specific inactivation of CTGF will ultimately reveal which
diabetes, conditional deletion of CTGF in adult ␤-cells will alcellular source of CTGF functions to promote proper lineage allolow us to explore whether decreased CTGF expression may be a
cation, ␤-cell proliferation, and/or islet morphogenesis.
contributing factor to type 2 diabetes or gestational diabetes in
332
Crawford et al.
CTGF in Pancreatic Islet Development
Mol Endocrinol, March 2009, 23(3):324 –336
allele (LOA) assay. Of 96 clones, eight were targeted, indicating a
targeting frequency of 8.3%. Two of these clones (286A-B8 and
286A-F9) were used to generate mice. CTGF⫺/⫺ mice resulting from
F1 CTGF⫹/⫺ intercrosses (from mice generated from both of these
clones) displayed a phenotype identical to that published elsewhere
(31) (data not shown).
Genotyping was performed by PCR amplification of a portion of CTGF
exon 4. This exon is missing in the null allele. Amplification of a 193-bp
fragment from the endogenous allele was performed using the following
primers: CTGFlacZfor, 5⬘-aagacacatttggcccagac-3⬘; and CTGFlacZrev,
5⬘-ttttcctccaggtcagcttc-3⬘. CTGFlacZ/⫹ animals were further distinguished
from WT using X-gal staining of ear punches (adult) or ribcage (embryo).
The pdx1-Cre mice were generously provided by Guoqiang Gu
(Vanderbilt University). The generation and genotyping of these mice
have been previously described (33, 38).
Tissue preparation, X-gal staining, and histology
FIG. 8. CTGF mutant animals have reduced ␤-cell proliferation. A, ␤-Cell
proliferation at e18.5 in WT (dark bar, n ⫽ 3), CTGF heterozygotes (light bar, n ⫽
3), and CTGF null (stippled bar, n ⫽ 3) animals. B, ␤-Cell proliferation at P2 in WT
(dark bar, n ⫽ 3) and CTGF heterozygotes (light bar, n ⫽ 3). Proliferation was
assessed using double immunofluorescence for insulin and phosphorylated
histone H3 as described in Materials and Methods. Error bars determined by SEM.
*, P ⬍ 0.05 compared with WT (determined by Student’s t test).
combination with metabolic and/or physiological stresses on the
␤-cell (i.e. increased body mass, increased insulin resistance).
Materials and Methods
Animals
Generation of a CTGF null allele (CTGFneo) and genotyping of mice
were previously described by Ivkovic et al. (31). The CTGFlacZ null
allele (Fig. 2) was generated as follows. Targeted embryonic stem (ES)
cells harboring a lacZ-targeted null allele of CTGF were generated using
Velocigene technology, essentially as described (56). Briefly, a bacterial
artificial chromosome (BAC) containing mouse genomic DNA encompassing CTGF was selected by PCR-screen from a BAC library of 129/
Svj mouse genomic DNA (Incyte). The CTGF BAC (identification no.
460d11) contains approximately 170 kb of genomic DNA. To generate
the targeting vector, the coding sequence of CTGF contained within
exon 3 [from the start of its von Willebrand factor type C (VWC)
domain] to the end of the coding sequence in exon 5 was replaced
with a transmembrane domain-lacZ/Neomycin phosphotransferase
(TM-lacZ/ NeoR) cassette using bacterial homologous recombination (BHR) (57– 60). The NeoR selection minigene was flanked with
LoxP sites to allow its removal using Cre (61). Using restriction mapping, it was determined that the resulting modified BAC had homology arms of approximately 120 and 40 kb flanking the TM-lacZ/
NeoR cassette. This modified BAC was used as a targeting vector to
target the CTGF gene in an F1 (C57BL/6-129SvJ) hybrid ES line.
Ninety six ES cell clones were picked and genotyped using a loss of
Digestive organs or pancreata from embryonic and adult stages were
dissected in PBS and fixed immediately in 4% paraformaldehyde (3–5
h). Tissues were dehydrated using an increasing ethanol series, followed
by two xylene washes, infiltrated with xylene-paraffin (1:1; vol/vol) and
two changes of paraffin at 56 C, and embedded in paraffin for sectioning. Pancreata for frozen sectioning were dissected and fixed as described above and then placed in a 30% sucrose solution overnight.
Tissue was then embedded and frozen in Tissue-Tek O.C.T. compound
and sectioned at 5 ␮m.
␤-gal activity was detected using X-gal as previously described (62).
Tissues were postfixed (4% paraformaldehyde in PBS; 1 h at 4 C) and
then dehydrated for embedding as above, with isopropanol replacing
xylene to minimize leaching of the blue precipitate. For colabeling on
X-gal-stained tissue, antibodies were added directly to sections of pancreata that had been stained with X-gal in whole mount.
Serial 5-␮m sections were deparaffinized and rehydrated using an
increasing ethanol series to distilled water. Indirect protein localization
was obtained by incubation of tissue with the following primary antibodies: guinea pig antiinsulin (1:1000; Linco Research, Inc., St. Charles,
MO), rabbit anti-CTGF raised against residues 81–94 (1:500; this antibody did not react with tissue from CTGF null animals, demonstrating
its specificity; data not shown) (63); guinea pig antiglucagon (1:1000;
Linco); rabbit antiphosphorylated histone H3 (pH3; Upstate Biotechnology, Inc., Lake Placid, NY); rabbit anticytokeratin (1:1000; DAKO
Corp., Carpinteria, CA), mouse anti-PECAM (PharMingen, San Diego,
CA; 1:200), mouse antisynaptophysin (Upstate, 1:500), mouse anti-␤catenin (BD Transduction Laboratories, Lexington, KY; 1:50). All primary antibodies were incubated overnight at 4 C in a humid chamber.
Detection of CTGF, and pH3, and synaptophysin required antigen retrieval. For CTGF, sections were microwaved in TEG (1 mM Tris, 0.5
mM EGTA; pH 9.0) buffer on 10% power until lightly boiling. For pH3,
synaptophysin, and ␤-catenin, slides were microwaved in 10 mM sodium citrate on full power for 3 min, followed by a 1 min cool and
another 1 min on full power. Chicken anti-␤-gal antibody (1:1000;
Abcam, Inc., Cambridge, MA) was used on cryosections to detect ␤-gal
protein expression.
Primary antibodies were detected by species-specific donkey secondary antibodies conjugated to Cy2 or Cy3 fluorophores (1:500; Jackson
ImmunoResearch Laboratories, Inc., West Grove, PA). In the case of
CTGF, amplification was needed such that antirabbit biotin was added
for 1 h at room temperature, followed by fluorescent-conjugated
streptavidin (1:1000; Vector Laboratories, Inc., Burlingame, CA). Fluorophores were excited using either an Olympus BX41 research microscope or a Zeiss LSM 510 confocal microscope (Carl Zeiss, Thornwood,
NY). Tiff images were captured using either MagnaFire software
(Optronics Engineering, Goleta, CA) or LSM Viewer (Zeiss), and total
range and brightness were equivalently adjusted by histogram using
Adobe Photoshop (Adobe Systems, Inc., San Jose, CA).
Mol Endocrinol, March 2009, 23(3):324 –336
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333
FIG. 9. CTGF mutant animals have an increase in p27Kip1 levels. A, Western blot showing four individual pancreatic samples at e18.5 from each genotype (WT,
CTGF⫹/⫺ and CTGF⫺/⫺). B, Densitometric quantitation of p27 levels illustrated as a ratio of p27:␤ actin, where WT levels were assigned a value of 1.0. Error bars
represent SEM. P values were calculated using a Student’s t test. Arrowhead indicates band quantitated for densitometry. HET, Heterozygous; KO, knockout.
Insulin and glucagon area
␤-Cell apoptosis
Entire paraffin blocks containing e18.5 digestive organs or adult pancreas were serially sectioned at 5 ␮m. Every tenth slide of pancreas tissue
was immmunolabeled for insulin and glucagon (described above). Every
islet from one section on each slide was photographed using an Olympus
BX41 microscope and MagnaFire software. Insulin and glucagon area
in each islet was calculated using Metamorph 6.1 morphometric analysis software. Islets were considered to be mixed in adult animals if
glucagon-expressing cells were found beyond the fourth cell layer from
the outermost edge.
␤-Cell apoptosis was quantitated using the ApoAlert DNA fragmentation assay kit (CLONTECH Laboratories, Inc.) according to the
manufacturer’s instructions. Slides were colabeled with insulin (described above). Using Metamorph 6.1 software, the total number of
apoptotic cells was counted, as were the number of cells colabeled for
both insulin and apoptosis. The percentage of ␤-cells undergoing
apoptosis (colabeled for Apo and insulin) was determined.
␣-Cell and ␤-cell proliferation
Entire paraffin blocks containing e15.5-P2 digestive organs were serially
sectioned at 5 ␮m. Every sixth (for e14.5 and e16.5) or tenth (for e17.5e18.5) slide was immunolabeled for pH3 and insulin or glucagon (described above). For adult tissue, blocks containing pancreata was serially sectioned at 5 ␮m. One hundred slides containing five sections per
slide were made. From these, every twentieth slide was immunolabeled
for pH3 and insulin as previously described. Every insulin or glucagon
cell on one section per slide was photographed using an Olympus
BX41 microscope and MagnaFire software. Using Metamorph 6.1
software, the total number of cells positive for insulin or glucagon
only was counted, as were the number of cells colabeled for both
insulin or glucagon and pH3. The percentage of ␤- or ␣-cells proliferating (colabeled for pH3 and insulin or glucagon) was determined.
␤-Cell size
Entire paraffin blocks containing adult pancreas were serially sectioned
at 5 ␮m. Every thirtieth slide of pancreas tissue was immmunolabeled
for insulin (described above). Every islet from one section on each slide
was photographed using an Olympus BX41 microscope. Using Metamorph 6.1 software, the total ␤-cell area for each islet was determined.
The number of ␤-cells in each islet was counted, and average ␤-cell size
for each islet was determined by dividing the total ␤-cell area by the
number of ␤-cell nuclei. ␤-Cells from at least 125 islets from three
animals per genotype were used to determine the average ␤-cell size. As
an alternative method to measure ␤-cell size, rabbit antimouse Glut2
(from Bernard Thorens, 1:500) was used to visualize the plasma
membrane of ␤-cells. Using Metamorph 6.1 software, the area of 100
individual ␤-cells was determined in at least five different islets per
mouse. Three mice per genotype were analyzed to determine the
average ␤-cell size.
FIG. 10. CTGF⫹/⫺ adult islets exhibit a range of phenotypes. Insulin (green) and glucagon (red) labeling of WT (A) and CTGF⫹/⫺ heterozygous (B–D) islets. B, CTGF⫹/⫺
islet with normal islet architecture. C, CTGF⫹/⫺ islet with mixing of ␣-cells into the ␤-cell core. D, Two apparently fused islets that failed to separate during islet
morphogenesis. gluc, Glucagon; ins, insulin.
334
Crawford et al.
CTGF in Pancreatic Islet Development
Mol Endocrinol, March 2009, 23(3):324 –336
FIG. 11. ␤-Cell hypertrophy in CTGF⫹/⫺ adult islets. ␤-Cell size was measured by labeling for Glut2 (red), which specifically outlines the membranes of ␤-cells (insulin,
green) in both wild type (A) and CTGF heterozygous (B) adults. C, There was a statistically significant increase in average ␤-cell size in the CTGF heterozygous animals.
See Materials and Methods for details regarding measurement of ␤-cell size. Het, Heterozygous.
In vivo analysis of glucose homeostasis
Intraperitoneal glucose tolerance tests (IP-GTTs) were performed as
previously described (39). Briefly, 4- to 12-wk-old mice fasted for 16 h
were given ip injection of filter-sterilized glucose in PBS (2.0 mg dextrose/g body weight). Glucose concentrations were measured in tail vein
blood using the Freestyle glucose meter and test strips (TheraSense, Inc.,
Alameda, CA) before injection (time 0) and 15, 30, 60, 90, and 120 min
after injection.
Pancreatic extracts and measurement of insulin content
Pancreata from e17.5 embryos and 4-wk-old mice were dissected,
weighed, and homogenized in acid alcohol for extraction of insulin (64).
Insulin content from acid alcohol-extracted pancreas was measured by
solid-phase RIA (125I-insulin, Diagnostic Products Corp., Los Angeles,
CA) for mouse antiinsulin (MP Biomedicals, Irvine, CA). Average insulin concentration was calculated as a function of total pancreatic wet
weight.
Protein extraction and Western blotting
Pancreata from e18.5 embryos were dissected in ice-cold PBS and immediately placed into extraction buffer containing protease inhibitors
(0.5 mg/liter N-␣-tosyl-L-phenylalanylchloromethylketone, 0.5 mg/liter
N-␣-tosyl-L-lysine chloromethyl kefone hydrochloride (TLCK), 0.6 ␮M
leupeptin, and 2 ␮M pepstatin), 0.5 M dithiothreitol, and 0.1 M phenylmethylsulfonyl fluoride. Samples were homogenized and centrifuged
briefly, and the supernatant was frozen at ⫺80 C. Protein was quantitated using the Bio-Rad DC protein assay according to manufacturer’s
instructions (Bio-Rad Laboratories, Hercules, CA). Western blot analysis was performed on individual pancreatic protein extracts. Protein
was electrophoresed on 4 –12% Bis Tris gels under denaturing conditions, and blotted to polyvinylidenedifluoride membrane using the NuPAGE Western blotting system (Invitrogen, Carlsbad, CA). Blots were
blocked in 5% nonfat milk in Tris-buffered saline (TBS) (pH 7.6) for 1 h
at room temperature and probed with primary antibodies diluted in 3%
nonfat milk in TBS overnight at 4 C: mouse anti-␤ actin (1:5000; Santa
Cruz Biotechnology, Inc., Santa Cruz, CA), mouse anti-p27
(1:2500; BD Biosciences, Palo Alto, CA), mouse anti-␤-catenin (1:500;
BD Biosciences), and washed in 0.05% Tween 20 in TBS for 30 min at
room temperature with three changes of buffer. Horseradish peroxidase-conjugated species-specific secondary antibodies were diluted to
1:3000 (antimouse IgG; Promega Corp., Madison, WI) in 1% nonfat
milk in TBS and incubated for 1 h at room temperature. After washes,
protein detection was performed using the ECL detection system (Amersham Pharmacia Biotech, Arlington, IL) per manufacturer’s instructions using Kodak X-Omat Blue film (Eastman Kodak, Rochester, NY).
Protein levels in individual pancreata were quantitated on a Molecular
Imager FX densitometer (Bio-Rad) using Quantity One 4.6 software
(Bio-Rad) and normalized to the quantity of ␤-actin obtained for
each sample. Protein levels were represented as a ratio of p27:␤-actin
or ␤-catenin:␤-actin, where WT levels were assigned a value of 1.0.
Error bars were determined using the SEM. P values were calculated
using Student’s t test.
High-fat diet feeding
At weaning (21 d after birth) animals were assayed by IP-GTT, and
subsequently placed on high-fat diet (58.7% fat, 26.7% carbohydrate,
14.8% protein; Bio-Serv) or normal (control) mouse chow (24% fat,
57% carbohydrates, 19% protein; TestDiet) and fed ad libitum. Every 4
wk for 3 months, animals were subjected to IP-GTT as above.
Acknowledgments
We thank Dr. Elizabeth Tweedie Ables and other members of the Gannon laboratory for helpful discussions and suggestions. A special thanks
to Christine Pope and David Lowe for technical assistance. We thank
Mol Endocrinol, March 2009, 23(3):324 –336
Jami Day and Andre Boustani for help with adult morphometric analyses, and Dr. Shubhada Jagasia for helpful discussions. Thanks also to
Dr. Guoqiang Gu for generously providing the pdx1-Cre mice and Dr.
Bernard Thorens for the Glut2 antibody. Imaging was performed, in
part, through the use of the Vanderbilt University Medical Center Cell
Imaging Shared Resource.
Address all correspondence and requests for reprints to: Maureen
Gannon, Ph.D., Department of Medicine, Division of Diabetes, Endocrinology, and Metabolism, Vanderbilt University Medical Center, 2213 Garland Avenue, 7425C MRBIV, Nashville, Tennessee 37232-0475. E-mail:
[email protected].
This work was supported by National Institutes of Health Grants
CA68485, DK20593, DK58404, HD15052, DK59637, and EY08126
(to the VUMC Cell Imaging Shared Resource), NIH Grant DK065131,
and JDRF Career Development Award, Grant 2-2002-583 (to M.G.),
and the Vanderbilt Molecular Endocrinology Training Program, Grant
5-T32-DK07563 (to L.C. and M.A.G.).
Disclosure Statement: L.A.C., M.A.G., Y.A.O., A.D., K.M.L.,
D.R.B., and M.G. have nothing to declare. D.M.V., A.J.M., G.D.Y., and
A.E. are employees of Regeneron Pharmaceuticals.
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