Deletion of the PDGFR-ß Gene Affects Key Fibroblast ... · phosphorylation of Akt, ... analysis of granulation tissue formation following subcutaneously ... resulting ES clones in

Gao et al.

Deletion of the PDGFR-ß Gene Affects Key Fibroblast Functions

Important for Wound Healing

Zhiyang Gao1, Toshiyasu Sasaoka2, Toshihiko Fujimori3, Takeshi Oya1, Yoko Ishii1,

Hemragul Sabit1, Makoto Kawaguchi4, Yoko Kurotaki3, Maiko Naito2, Tsutomu Wada5,

Shin Ishizawa1, Masashi Kobayashi5, Yo-Ichi Nabeshima3 and Masakiyo Sasahara1, 6*

From the 1 Department of Pathology, 2 Department of Clinical Pharmacology and 5 First

Department of Internal Medicine, Toyama Medical and Pharmaceutical University,

2630 Sugitani, Toyama 930-0194, Japan.

3 Department of Pathology and Tumor Biology, Graduate School of Medicine, Kyoto

University, Kyoto 606-8501, Japan.

4 Department of Pathology, Niigata Rosai Hospital, Labor Health and Welfare

Organization 1-7-12 Tooun-cho, Jhoetsu, Niigata 942-8502, Japan.

6 Core Research for Evolutional Science and Technology, Japan Science and

Technology Agency (CREST, JST), 4-1-8 Honcho, Kawaguchi-shi, Saitama 332-0012,

Japan.

*Corresponding author: Department of Pathology, Toyama Medical and Pharmaceutical

University, 2630 Sugitani, Toyama 930-0194, Japan.

Tel.: 81-76-434-7238

Fax: 81-76-434-5016

E-mail: [email protected]

Running title: Roles of PDGFR-ß in Dermal Fibroblasts

JBC Papers in Press. Published on December 6, 2004 as Manuscript M413081200

Copyright 2004 by The American Society for Biochemistry and Molecular Biology, Inc.

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Summary

This study provides new perspectives of unique aspects of platelet-derived growth

factor beta-receptor (PDGFR-β) signaling and biological responses through the

establishment of a mutant mouse strain in which two loxP sequences were inserted into

the introns of PDGFR-ß genome sequences. Isolation of skin fibroblasts from the

mutant mice and Cre-recombinase transfection in vitro induced PDGFR-ß gene deletion

(PDGFR-ß∆/∆). The resultant depletion of PDGFR-ß protein significantly attenuated

PDGF-BB-induced cell migration, proliferation, and protection from H2O2-induced

apoptosis of the cultured PDGFR-ß∆/∆ dermal fibroblasts. PDGF-AA and fetal bovine

serum were mitogenic and anti-apoptotic but were unable to induce the migration in

PDGFR-ß∆/∆ fibroblasts. Concerning the PDGF signaling, PDGF-BB-induced

phosphorylation of Akt, ERK1/2 and JNK, but not p38, decreased in PDGFR-ß∆/∆

fibroblasts, but PDGF-AA-induced signaling was not altered. Overexpression of

phospholipid phosphatases, SHIP2 and/or PTEN, inhibited PDGF-BB-induced

phosphorylation of Akt and ERK1/2 in PDGFR-ß∆/∆ fibroblasts, but did not affect that

of JNK and p38. These results indicate that disruption of distinct PDGFR-ß signaling

pathways in PDGFR-ß∆/∆ dermal fibroblasts impairs their proliferation and survival, but

completely inhibits migratory response, and that PDGF-BB-induced phosphorylation of

Akt and ERK1/2 possibly mediated by PDGFR-α is regulated, at least in part, by the

lipid phosphatases SHIP2 and/or PTEN. Thus, the PDGFR-ß function on dermal

fibroblasts appears to be critical in PDGF-BB action for skin wound healing, and is

clearly distinctive from that of PDGFR-α in the ligand- induced biological responses and

the underlying properties of cellular signaling.


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Introduction

Platelet-derived growth factors (PDGFs) are major mitogens for connective tissue

cells that are involved in diverse biological processes including physiological

development, tissue repair, tumorigenesis, and atherosclerosis (1). PDGF family

members, PDGF-A, -B, -C and -D, are assembled as disulfide- linked homo- or

heterodimers, and exert their activity by binding to and activating specific high-affinity

cell surface receptors. Two receptor subtypes with protein tyrosine kinase activity

have been identified that can form homo- and hetero-dimeric receptor complexes: the

α-subunit, which can bind to the A-,B- and C-chains of PDGF and the ß-subunit,

specific for the B-, C- and D-chains (1, 2, 3, 4).

Dermal fibroblasts are one of the major target cells of PDGF in the initiation and

propagation of wound healing in the skin (5). Levels of PDGF receptor (PDGFR)-α

expression are high in fibroblasts during early embryogenesis, and disruption of

PDGFR-α results in a reduction in fibroblasts throughout the embryo (6, 7). In

contrast, targeted deletion of PDGFR-β and analysis of blastocyst chimeras

demonstrated no effect of PDGFR-β on fibroblast development (8, 9). However, in

mice prepared from the blastocyst chimeras that contain a combination of wild-type and

PDGFR-ß-/- cells, analysis of granulation tissue formation following subcutaneously

implantation of sponges demonstrated that PDGFR-ß-/- cells were depleted in the

granulation tissue (10). These reports suggest that the two subtypes of PDGFR play

distinct roles in development, and that PDGFR-ß is important in wound healing by

dermal fibroblasts. Analysis of mutant mice in which the cytoplasmic signaling

domain of the PDGFR-ß was used to replace the PDGFR-α cytoplasmic domain


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showed no obvious defects in any of the PDGFR-α -dependent cell types (11). On the

other hand, when the PDGFR-ß was dependent upon PDGFR-α cytoplasmic domain,

multiple abnormalities occurred in vascular smooth muscle cell development (11).

These data suggest that PDGFR-ß has unique signaling capacities comparing to

PDGFR-α.

PDGF binding to the receptors activates a variety of intracellular signaling

molecules (12). One of these is phosphatidylinositol 3-kinase (PI3K), which results in

the local accumulation of PI(3,4,5)-triphosphate (PI(3,4,5)P3) at the plasma membrane.

Synthesized PI(3,4,5)P3 recruits the pleckstrin homology domain-containing signaling

molecule Akt/PKB to the membrane (12, 13). PDGFs are also known to activate the

mitogen-activated protein kinase (MAPK) families including the extracellular

signal-regulated kinase (ERK1/2), c-Jun N-terminal kinase (JNK, also known as SAPK),

and p38 MAPK. ERK1/2 is predominantly activated by receptor tyrosine kinases of

growth factors, while JNK and p38 are preferentiallyactivated by stress- inducing

stimuli such as UV light, heat shock, and pro- inflammatory cytokines, and by mitogenic

stimuli as well (14). The PI3K/Akt and MAPKs pathways play important roles in the

regulation of cell growth, migration, and survival during the wound healing process (13,

15, 16), although the relative importance of these kinases in various cellular phenomena

differ depending on the cell type (14). SHIP2 and PTEN are recently identified

phospholipid phosphatases that are thought to negatively regulate the PDGF activation

of PI3K/Akt pathway by removing 5’- and 3’-phosphates, respectively, from

PI(3,4,5)P3 . In addition, PTEN and SHIP2 also appear to inhibit PDGF activation of

ERK1/2 (17, 18). However, the specific role of PDGFR-ß in the activation of Akt and


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MAPKs, and the implication of the phospholipid phosphatases in PDGFR-ß-mediated

signaling are still poorly understood.

To elucidate the biological properties and signaling pathways of PDGFR-ß, it is

critical to analyze its function in non-transformed cells that physiologically express the

receptor. Because homozygous disruption of PDGF-B or PDGFR-ß results in

perinatal death in mouse embryos (19, 8), it has been difficult to evaluate the role of the

PDGF-B/PDGFR-ß system in repair of adult tissues, such as wound healing. To

address this problem, we established a new mouse strain in which the PDGFR-ß exons

are flanked by two loxP sequences. The Cre-mediated recombination by

adenovirus-gene transfer markedly reduced the expression of PDGFR-ß in the dermal

fibroblasts derived from the mutant mice without affecting the expression of PDGFR-α.

Using these dermal fibroblasts, we studied PDGF-induced biological effects, including

proliferation, migration, and apoptosis, which are central processes in wound healing.

In addition, we examined the involvement of PDGFR-ß in the activation of Akt and

MAPKs to clarify the intracellular signals activated by the PDGFs. Finally, we

examined the effect of the phospholipid phosphatases, SHIP2 and PTEN, in the

regulation of PDGF-BB signaling in dermal fibroblasts lacking PDGFR-ß.

Experimental Procedures

Materials

Human recombinant PDGF-AA was provided by Strathmann Biotech AG

(Hamburg, Germany), and human recombinant PDGF-BB was purchased from Life

Technologies, Inc. (Grand Island, NY). Polyclonal anti-PDGFR-ß antibody was from


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Upstate Biotechnology (Lake Placid, NY). Polyclonal anti-PDGFR-α antibody,

polyclonal anti-Tyr857 phosphospecific PDGFR-ß antibody, monoclonal

anti-phosphotyrosine antibody (PY99), monoclonal anti-Akt1 antibody, and monoclonal

anti-PTEN antibody were from Santa Cruz Biotechnology (Santa Cruz, CA).

Polyclonal anti-Thr308 phosphospecific Akt antibody, polyclonal ERK1/2 antibody,

polyclonal anti-Thr202/Tyr204 phosphospecific ERK1/2 antibody, polyclonal anti-JNK

antibody, polyclonal anti-Thr183/Tyr185 phosphospecific JNK antibody, polyclonal

anti-p38 antibody, and polyclonal anti-Thr180/Tyr182 phosphospecific p38 antibody were

from Cell Signaling Technology, Inc. (Beverly, MA). Polyclonal anti-SHIP2 antibody

was generated as described previously (20). Fetal bovine serum (FBS) was obtained

from Bio Whittaker A Cambrex Inc. (Walkersville, ML). Dulbecco’s modified

Eagle’s medium (DMEM) was from Nissui Pharmaceutical Co., Ltd. (Tokyo, Japan).

All other reagents were of analytical grade and were purchased from Sigma (St. Louis,

MO) or Wako Pure Chemical Industries Ltd. (Osaka, Japan).

Generation of PDGFR-ßflox/flox mutant mice

For the construction of the targeting vector, a 13.5 kb BamHI/SpeI genomic

fragment of the PDGFR-ß gene from the 129X1/SvJ mouse was cloned into

p-Bluescript SK- (Stratagene, La Jolla, CA). We inserted a neomycin-thymidine

kinase (neo-TK) selection cassette and one-loxP sequence at the 5’- and 3’-ends,

respectively, of the 3.3 kb FspI/HpaI genomic fragment that includes exons 4 to 7

encoding the extracellular domain of the PDGFR-ß (Fig. 1A). The neo-TK selection

cassette consisted of MC1-neo and MC1-TK selection cassette flanked by two loxP

sites. A diphtheria toxin selection cassette (DT-A) was inserted at the 3’-end of the


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vector (Fig. 1A). After linearization, the vector was electroporated into 129SV mouse

embryonic stem (ES) cells. The obtained recombinant ES clones were transfected with

the pCre-Pac plasmid by electroporation to delete the neo-TK selection genes. The

resulting ES clones in which the exons 4 to 7 of the PDGFR-ß gene were flanked by

two loxP sequences (PDGFR-ßflox/+), were then injected into blastocysts to obtain

chimeric mice. The chimeras were bred with C57BL/6 mice (Sankyo Lab. Tokyo,

Japan) to obtain the F1 progenies containing the recombinant allele (PDGFR-ßflox/+).

These F1 progenies were crossbred to obtain PDGFR-ßflox/flox homozygous mice.

Genotyping was performed by both Southern blotting and PCR-based analyses. The

probes for Southern blotting are shown in Figure 1A. The primers for genomic PCR

were as follows: primer 1, 5’-TAGCCATGGAGTCATCTCTTCAGCCCTAAA-3’;

primer 2, 5’-CCTGCATCAAGTAGCTCACAACTGCCTGTA-3’; primer 3,

5’-TTCTTGTCTGAGAGCCTGTTGTGTGATGGA-3’; primer 4,

5’-TGTCTGCAGATCTCTAGCCTTGGGGAAATC-3’; primer 5,

5’-AGCAAGGTCGCGCAAGGGATAACAGC-3’.

Adenovirus

We used adenovirus expression vectors with an E1-deleted replication deficiency.

Two adenoviral vectors expressing Cre recombinase and LacZ tagged with a nuclear

localization signal under the control of the CAG promoter (21) were kindly provided by

Dr. Izumu Saito (Tokyo University, Tokyo, Japan) (22). The recombinant adenovirus

expressing PTEN (23) was a generous gift of Dr. Tomoichiro Asano (Tokyo University,

Tokyo, Japan). The SHIP2-expressing adenovirus vector was constructed as described

previously (18).


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Cell culture and infection with adenovirus

Mouse dermal fibroblast cells were prepared from 8- to 12-week-old male control

C57BL/6 mice and PDGFR-ßflox/flox mice. Briefly, mice were anesthetized with

pentobarbital (50 mg/kg body weight), and a full thickness of the back skin was cut out

by scissors. The skin tissues were cut into small pieces and were implanted into

plastic tissue culture dishes containing DMEM with 10% FBS. The fibroblast cultures

were used after three to seven passages. Cre recombinase, PTEN, and SHIP2 were

transiently expressed in cultured cells by adenovirus-mediated gene transfer. Cells

were infected in DMEM containing 5% FBS at a multiplicity of infection (m.o.i.) of 10

pfu/cell. After 16 h, the virus was removed by replacing the medium. Experiments

were conducted 24 to 48 h after the initial addition of the virus.

DNA synthesis assay

The cultured fibroblasts obtained from wild-type or PDGFR-ßflox/flox mice were

infected with Cre-adenovirus as described above. The cells were cultured in DMEM

containing 10% FBS for an additional 24 h. The cells were then plated into 96-well

plates at 1.5 x 104 cells/well in DMEM containing 10% FBS. After 24 h, the cells

were serum-starved for 24 h and then were incubated for 18 h with 1 nM PDGF-AA, 1

nM PDGF-BB, or 10% FBS in DMEM containing 10 mM 5-bromo-2’-deoxyuridine

(BrdU). BrdU incorporation into DNA was determined with a

5-Bromo-2’-deoxy-uridine Labeling and Detection Kit (Roche, Mannheim, Germany)

according to the manufacturer’s instructions and detected with an ELISA plate reader

(Nippon InterMed. Ltd., Tokyo, Japan).


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Wound scratch assay

Confluent cultured fibroblasts obtained from wild-type or PDGFR-ßflox/flox mice

were grown in 6-well-plates and were infected with Cre-adenovirus as described above.

The cells were cultured in DMEM containing 10% FBS for an additional 24 h. After

the cells were serum-starved for 24 h in DMEM, they were scratched with a 1-ml plastic

pipette tip and washed twice with PBS to remove the floating cells. The cells were

then cultured in DMEM supplemented with 1 nM PDGF-AA, 1 nM PDGF-BB, or 10%

FBS. After 48 h, the cells were viewed with an inverted phase contrast microscope

(Olympus CK30, Tokyo, Japan) and photographed.

Apoptosis assay

Apoptosis was detected using an APOPercentageTM kit (Biocolor Ltd., Belfast,

NorthernIreland). Briefly, confluent wild-type or PDGFR-ß flox/flox fibroblasts grown

in 6-well plates were infected with the Cre-adenovirus as described above. After

infection, the cells were replated into 12-well plates and cultured for further 24 h in

DMEM containing 10% FBS. The apoptosis were induced by 1 µM H2O2 in 1 ml

DMEM containing 1 nM PDGF-AA, 1nM PDGF-BB, or 10% FBS, and 50 µl of

APOPercentage Dye. After 60 min, the medium was drained from the wells, and the

cells washed twice with PBS. The cells were then observed with an inverted phase

contrast microscope.

Immunoprecipitation and Western blotting

Confluent wild-type and PDGFR-ßflox/flox fibroblasts grown in 6-well plates were


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infected for 16 h with the Cre-adenovirus at a m.o.i. of 10 pfu/cell. The cells were

then incubated for 24 h in DMEM containing 10% FBS. Next, the cells were

serum-starved for 48 h and then treated with various concentrations of PDGF-AA or

PDGF-BB at 37oC for the indicated times. The cells were lysed for 15 min at 4ºC in a

buffer consisting of 50 mM Tris, 150 mM NaCl, 10 mM EDTA, 0.5% sodium

deoxycholate, 1% Triton X-100, 2 mM Na3VO4, 150 mM sodium fluoride, 2 mM

phenylmethylsulfonylfluoride, 10 µg/ml aprotinin, and 10 µg/ml leupeptin (pH 7.4).

Lysates obtained from the same number of cells were centrifuged to remove insoluble

materials. The supernatants were incubated with the indicated antibodies for 4 h at

4ºC and then immune complexes were mixed with glutathione-Sepharose for 2 h at 4ºC.

The immune complexes were precipitated by centrifugation. Immune complexes or

whole cell lysates were separated by SDS-PAGE and then electrophoretically

transferred to polyvinylidene difluoride membranes. The membranes were incubated

for 1 h at 20ºC in a buffer containing 50 mM Tris, 150 mM NaCl, 0.1% Tween 20, and

5% non-fat dry milk. The membranes were then probed with specified antibodies for

16 h at 4ºC. After the membranes had been washed in a buffer containing 50 mM Tris,

150 mM NaCl, and 0.1% Tween 20, the blots were incubated with an appropriate

horseradish peroxidase-conjugated secondary antibody. Immunoreactive bands were

detected using enhanced chemiluminescence reagents (Amersham Biosciences,

Buckinghamshire, UK) according to the manufacturer’s instructions.

Immunofluorescence

Cultured wild-type or PDGFR-ßflox/flox fibroblasts were infected with the

Cre-adenovirus as described above and were replated in chamber slides. All staining


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procedures were performed at room temperature. After growth for 24 h, the cells were

fixed with 1% formalin in PBS for 10 min. Non-specific immunoreactions were

blocked by incubation for 20 min with 10% goat serum in PBS. Cells were then

incubated for 2 h with 1:100 anti-PDGFR-ß, washed three times for 5 min with PBS,

and then incubated for 1 h with 1:1,000 Alexa Fluor 488-conjugated goat anti-rabbit

IgG (Molecular Probes, Eugene, OR). The cells were mounted with Vectashield

mounting medium containing DAPI (Vector Laboratories, Burlingame, CA) and

observed with an Olympus AX80 microscope.

Statistical analysis

All data are presented as means ± S.D. Student’s t-test was used to determine

the p values, and p < 0.05 was considered statistically significant.

Results

PDGFR-ß flox/flox mutant mice are healthy and show no gross abnormalities

After transfection of the targeting vector (Fig. 1A) and selection with neomycin,

we obtained 4 embryonic stem cell (ES cell) clones with homologous recombination out

of 319 clones examined. In addition to a 12.8 kb fragment, clones that had undergone

recombination generated a 7.3 kb fragment in Southern blotting using a 5’ -probe after

digestion of the genomic DNA with XbaI (Fig. 1B). Generation of a 5.7 kb fragment

after SacI digestion and 8.8 kb fragment after XbaI digestion was detected by Southern

blotting using a 3’ -probe and neo probe, respectively (Fig. 1B).

ES cell clones that had undergone homologous recombination were transfected

with Cre-expressing plasmid (pCre-Pac; gift from Dr. Takashi Yagi, Osaka University,


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Japan) to delete the neo-TK selection cassette. Deletion of the neo-TK selection gene

was confirmed by Southern blotting with an internal probe (data not shown). The

resulting ES clones (PDGFR-ßflox/+) were used for the generation of chimeric mice.

The PDGFR-ßflox/+ ES clones were also transfected with the pCre-Pac plasmid to

confirm that the Cre recombinase can induce recombination and eliminate the floxed

PDGFR-ß allele. After transfection with the pCre-Pac plasmid, genomic PCR with

primers 1, 2, and 5 generated bands of 271, 329 and 410 bp, which corresponded to the

PDGFR-ß genes for wild-type , floxed, and deleted alleles (PDGFR-ß? ), respectively

(Fig. 1C). These results confirmed that the Cre recombinase caused PDGFR-ß gene

deletion in the PDGFR-ßflox/+ ES cells.

The chimeras generated from PDGFR-ßflox/+ ES cells were bred with C57BL/6

mice, generating F1 progenies containing the recombinant allele (PDGFR-ßflox/+).

These mice were crossed with each other to obtain homologous PDGFR-ßflox/flox mice.

The PDGFR-ßflox/flox mice were healthy, and no gross abnormalities were detected.

The genotype of the mice was confirmed by PCR with primers 1 and 2 (Fig. 1D) and

primers 3 and 4 (data not shown). PCR with primers 1 and 2 generated a 271 bp band

from the wild-type PDGFR-ß gene and a 329 bp band from the floxed allele.

Adenovirus-mediated Cre expression eliminates the expression of PDGFR-ß in

PDGFR-ßflox/flox dermal fibroblasts

We first investigated whether Cre-adenovirus infection could abrogate the

expression of PDGFR-ß in PDGFR-ßflox/flox dermal fibroblasts. Similar levels of

PDGFR-ß were expressed in both wild-type and PDGFR-ßflox/flox fibroblasts in the

absence of Cre-adenovirus infection (Fig. 2A). Treatment with 1 nM PDGF-BB


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induced similar levels of tyrosine phosphorylation of PDGFR-ß in both types of

fibroblast (Fig. 2A). Infection with the Cre-adenovirus caused a marked reduction in

the expression and tyrosine phosphorylation of PDGFR-ß in PDGFR-ßflox/flox but not

wild-type fibroblasts. However, Cre transfection had no effect on the expression of

PDGFR-α in wild-type or PDGFR-ßflox/flox fibroblasts (Fig. 2A). Immunofluorescent

studies revealed that the cytoplasmic immunoreactivity for the PDGFR-ß was

eliminated in greater than 90% of PDGFR-ßflox/flox fibroblasts after Cre transfection

(PDGFR-ß∆/∆) (Fig. 2B).

The effects of Cre transfection were further assessed by examining PDGF-AA-

and PDGF-BB-induced tyrosine phosphorylation of PDGFR-α and PDGFR-ß.

Stimulation by either PDGF-AA or PDGF-BB induced equivalent and dose-dependent

tyrosine phosphorylation of PDGFR-α in both Cre-transfected wild-type and

PDGFR-ßflox/flox fibroblasts (Figs. 2C-a and 2C-b). In contrast, PDGF-BB-induced

tyrosine phosphorylation of PDGFR-ß was not detected in PDGFR-ß∆/∆ fibroblasts,

although it was clearly observed in wild-type fibroblasts following Cre transfection (Fig.

2C-c). Thus, Cre transfection of PDGFR-ßflox/flox fibroblasts selectively abrogated the

expression and function of PDGFR-ß but not PDGFR-α.

PDGF-BB- and fetal bovine serum-induced BrdU incorporation is decreased after

PDGFR-ß gene deletion

PDGFR is known to play an important role in cell proliferation (12). We

therefore examined the effect of Cre transfection on PDGF-AA-, PDGF-BB- and fetal

bovine serum (FBS)- induced BrdU incorporation in wild-type and PDGFR-ßflox/flox


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fibroblasts. As shown in Figure 3, 1 nM PDGF-AA induced similar levels of BrdU

incorporation in wild-type and PDGFR-ß∆/∆ cells. In contrast, 1 nM PDGF-BB- and

10 % FBS-induced BrdU incorporation were significantly decreased in PDGFR-ß∆/∆

fibroblasts compared to wild-type fibroblasts, 48.4 ± 1.2% and 34.2 ± 1.9%, respectively.

These results demonstrate a significant contribution of PDGFR-ß to cell proliferation in

response to PDGF-BB and FBS, but not PDGF-AA.

In vitro repair of a scratch wound is inhibited after PDGFR-ß gene deletion

Because PDGFR is also known to be implicated in cell migration (12), we

examined the effect of the deletion of the PDGFR-ß gene on the combined migratory

and proliferative response to scratch wounding that mimics aspects of wound healing.

A wound was formed in confluent cell monolayers by scratching with a plastic pipette

tip. In wild-type fibroblasts cultured for 48 h with 1 nM PDGF-BB, the wound was

completely closed (Figs. 4A and 4B). Similarly, substantial numbers of wild-type

fibroblasts had filled in the scratched area following treatment with 10% FBS for 48 h

(Figs. 4E and 4F). In contrast, the scratched wound remained unclosed at 48 h after

culture with 1 nM PDGF-BB in PDGFR-ß∆/∆ fibroblasts (Figs. 4C and 4D). Wound

closure was mostly suppressed in PDGFR-ß∆/∆ cells after culture with 10% FBS (Figs.

4G and 4H). These results indicate that the PDGF-BB/PDGFR-ß system is essential

for the combined migration and proliferation required to fill in a scratch wound with

PDGF-BB- and 10% FBS as stimulants, and suggests that the role of PDGF-α is

minimal. This concept was further supported by the fact that PDGF-AA did not

induce apparent wound closure in either wild-type or PDGFR-ß∆/∆ cells (Figs. 4I-4L).


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Apoptosis induced by H2O2 is enhanced by elimination of the PDGFR-ß gene

We next examined the effect of the deletion of the PDGFR-ß gene on

H2O2-induced apoptosis in dermal fibroblasts. To detect apoptosis, we utilized the

APOPercentageTM kit, which stains apoptotic cells, but not necrotic cells, with a

purple-red color (24, 25). Few apoptotic cells were observed in wild-type or

PDGFR-ß∆/∆ fibroblasts when the cells were cultured with 10% FBS (data not shown).

In wild-type cells cultured with DMEM supplemented only with 0.2% bovine serum

albumin, the addition of H2O2 increased the number of apoptotic cells by 78.3 ± 3.2%.

The addition of 1 nM PDGF-BB protected wild-type cells from apoptosis. However,

PDGFR-ß∆/∆ fibroblasts showed an increase in the number of apoptotic cells from 23.3

± 3.7% in wild-type to 56.9 ± 6.5% in PDGFR-ß∆/∆ fibroblasts treated with 1 nM

PDGF-BB (Figs. 5A, 5B and 5G). These data indicate that PDGF-BB mediates

protection from H2O2-induced apoptosis through PDGFR-ß. In contrast, H2O2 induced

only a low level of apoptosis in wild-type and PDGFR-ß∆/∆ fibroblasts cultured in the

presence of 10% FBS or 1 nM PDGF-AA, and there was no significant difference in the

degree of apoptosis among the four treatments (apoptosis for wild-type cells in 10%

FBS was 23.8 ± 3.6%; for wild-type cells in PDGF-AA, 21.3 ± 2.9%; for PDGFR-ß∆/∆

cells in 10% FBS, 24.3 ± 1.9%; and for PDGFR-ß∆/∆ cells in PDGF-AA, 23.2 ± 1.6%;

Figs. 5C-5F and 5G). These data provide further data that PDGFR-α signaling by

PDGF-AA is not perturbed by conditional deletion of PDGFR-β , and that it is sufficient

to inhibit H2O2-induced apoptosis in fibroblasts.


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PDGF-BB-induced phosphorylation of Akt is decreased after PDGFR-ß gene deletion

Because various biological actions were impaired in PDGFR-ß∆/∆ fibroblasts, we

next investigated the effect of PDGFR-ß deletion on PDGF-induced intracellular

signaling. Akt is one of the important signaling molecules involved in the biological

effects of PDGF (12). As shown in Figure 6A, stimulation with PDGF-BB for 5 min

dose-dependently increased the phosphorylation of Akt in wild-type fibroblasts, and the

levels of phosphorylation reached a maximum at 1 nM PDGF-BB. Compared to the

wild-type cells, the levels of phosphorylation following treatment with PDGF-BB were

significantly lower in PDGFR-ß∆/∆ fibroblasts. At 0.2 nM PDGF-BB, Akt

phosphorylation was decreased by 50.8 ± 2.1% in PDGFR-ß∆/∆ cells compared

wild-type cells.

We further conducted time course studies of PDGF-BB-induced phosphorylation

of Akt. In wild-type cells, PDGF-BB induced the phosphorylation of Akt in a

time-dependent manner for up to 60 min, and the phosphorylation was sustained at high

levels until 120 min after stimulation (Fig. 6C). Akt phosphorylation was significantly

reduced at all time points in PDGFR-ß∆/∆ fibroblasts, while the dose (Fig. 6B) and time

dependency (Fig. 6D) of PDGF-AA-induced phosphorylation of Akt were comparable

in wild-type and PDGFR-ß∆/∆ fibroblasts.

PDGF-BB-induced phosphorylation of ERK1/2 and JNK, but not p38, are decreased

after deletion of the PDGFR-ß gene

In addition to Akt, MAPKs, including ERK1/2, JNK, and p38, are known to be

key molecules in the biological actions of PDGF (26). Therefore, we investigated the

effect of PDGFR-ß gene deletion on PDGF-induced phosphorylation of ERK1/2, JNK,


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and p38. In wild-type fibroblasts, PDGF-BB dose-dependently induced the

phosphorylation of ERK1/2 at up to 1 nM (Fig. 7A). In the presence of 1nM

PDGF-BB, the levels of phosphorylation reached a maximum after 5 min with a gradual

decrease thereafter (Fig. 7C). PDGF-BB-induced ERK1/2 phosphorylation was

significantly decreased at all doses and time points in PDGFR-ß∆/∆ cells (Figs. 7A and

7C). Phosphorylation of JNK was induced by PDGF-BB in a similar dose- and

time-dependent manner as that observed in ERK1/2. JNK phosphorylation was also

decreased in PDGFR-ß∆/∆ cells compared to wild-type cells (Figs. 8A and 8C). At 0.4

nM PDGF-BB, the levels of ERK1/2 and JNK phosphorylation were reduced by 42.7 ±

3.8% and 53.2 ± 2.7%, respectively, in PDGFR-ß∆/∆ cells compared to wild-type cells.

The phosphorylation of ERK1/2 and JNK induced by PDGF-AA was essentially

identical between wild-type and PDGFR-ß∆/∆ cells at all doses and time points examined

(Figs. 7B, 7D, 8B, and 8D).

PDGF-BB and PDGF-AA dose-dependently induced the phosphorylation of p38

up to a concentration of 2 nM (Figs. 9A and 9B). In addition, both 1 nM PDGF-BB

and 1 nM PDGF-AA caused a transient phosphorylation that lasted for 15 min,

decreasing thereafter and returning to a basal level at 120 min (Figs. 9C and 9D).

Interestingly, both PDGF-BB- and PDGF-AA-induced phosphorylation of p38 was

nearly identical at all dose and time points in wild-type and PDGFR-ß∆/∆ cells.

Effect of PTEN and SHIP2 expression on the phosphorylation of Akt and MAPKs in

wild-type and PDGFR-ß∆/∆ fibroblasts

PTEN and SHIP2 are phospholipid phosphatases that may act as negative


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regulators of PDGF signaling (17, 18). To understand their roles in

PDGF-BB-induced phosphorylation of Akt and MAPKs, we overexpressed PTEN

and/or SHIP2 in wild-type and PDGFR-ß∆/∆ fibroblasts. As shown in Figure 10A, we

were able to enhance the expression of PTEN and SHIP2 by 10-folds of endogenous

expression. PDGF-BB-induced phosphorylation of Akt was only modestly decreased

by the expression of either PTEN or SHIP2, but was significantly inhibited when both

phosphatases were overexpressed in wild-type cells. In agreement with the results in

Fig. 6, the levels of the Akt phosphorylation were decreased by the depletion of

PDGFR-ß, and expression of either PTEN or SHIP2 effectively inhibited

PDGF-BB-induced phosphorylation of Akt in PDGFR-ß∆/∆ cells (64.2 ± 4.1% and 65.5

± 3.1% reduction, respectively; Fig. 10B).

In wild-type cells, PDGF-BB-induced phosphorylation of the ERK1/2 did not

appear to be affected by the overexpression of SHIP2 or PTEN. However, in

PDGFR-ß∆/∆ cells, overexpression of SHIP2 but not PTEN inhibited PDGF-BB-induced

phosphorylation of ERK1/2 (30.6 ± 1.2% reduction; Fig. 10C). In contrast to the

results with Akt and ERK1/2, expression of PTEN, SHIP2, or both did not affect

PDGF-BB-induced phosphorylation of JNK (Fig. 10D) or p38 (Fig. 10E) in either

wild-type or PDGFR-ß∆/∆ fibroblasts.

Discussion

In the current studies, we developed a new mouse line in which a portion of

PDGFR-ß exons were flanked by two loxP sequences that did not affect development of

the mutant mouse. The expression of PDGFR-α and -ß and the level of ligand- induced

phosphorylation were comparable between dermal fibroblasts isolated from wild-type


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and mutant mice without Cre transfection. The adenovirus-mediated Cre transfection

in PDGFR-ßflox/flox cells efficiently abrogated PDGFR-ß expression without affecting

PDGFR-α expression and its signaling pathways. Therefore, this is a powerful

approach for unambiguously determining the role of the two PDGFR subtypes in the

biological effects of and signaling pathways stimulated by PDGF.

In wild-type fibroblasts, PDGF-AA and PDGF-BB induced the incorporation of

BrdU, but PDGF-BB- and 10% FBS-induced BrdU incorporation were significantly

lower in PDGFR-ß∆/∆ fibroblasts than in wild-type fibroblasts. These results indicate

that PDGFR-ß plays an important role in mediating PDGF-BB- and 10% FBS-induced

DNA synthesis. However, because the depletion of PDGFR-ß caused only a partial

decrease of BrdU incorporation, PDGFR-α appears to partly mediate the effects of

PDGF-BB in dermal fibroblasts. In addition, PDGF-AA-induced DNA synthesis was

unaffected by depletion of PDGFR-ß, consistent with all of the effects of PDGF-AA

being mediated by PDGFR-α.

In the wound scratch assay, PDGF-BB and 10% FBS, but not PDGF-AA, induced

wound closure in wild-type fibroblasts. The wound closure induced by PDGF-BB and

10% FBS was significantly inhibited by depletion of the PDGFR-ß. Wound closure is

known to be dependent on both cell motility and proliferation. Because PDGF-AA

clearly induced cell proliferation in both wild-type and PDGFR-ß∆/∆ cells, the lack of

wound closure by PDGF-AA suggests that PDGFR-α signaling is insufficient to

stimulate cell motility. Similarly, studies with aortic endothelial cells transfected with

equal numbers of PDGFR-α or -ß showed that chemotactic migration was mediated

only by PDGFR-ß (27). Also, in dermal fibroblasts, PDGF-BB is known to be the


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major inducer of chemotaxis on type I collagen, although the subtype of PDGFR that

was involved was not determined (28). Furthermore, contribution of PDGFR-ß -/-

fibroblasts to granulation formation was reduced by 85% after subcutaneous

implantation of sponges in the blastocyst chimeric mice expressing mixtures of

wild-type and PDGFR-ß -/- cells (10). Taken together, our results clearly indicate that,

in dermal fibroblasts, PDGFR-ß, but not PDGFR-α, plays a crucial role particularly in

the mediation of PDGF-induced cellular motility.

PDGF-AA, PDGF-BB, and 10% FBS protected wild-type fibroblasts from

H2O2-induced apoptosis to similar extents. In PDGFR-ß∆/∆ fibroblasts, the effect of

PFGF-BB was mostly lost, whereas the protective effect of PDGF-AA and 10% FBS

were essentially unchanged. These results indicate that the PDGFR-ß plays an

important role in the ability of PDGF-BB to protect dermal fibroblasts from

H2O2-induced apoptosis, and that the remaining PDGFR-α appears to be sufficient for

the protection by PDGF-AA and 10% FBS.

Based on these results, the functional importance of PDGFR-ß and PDGFR-α

appears to differ depending on the specific biological effects triggered by PDGF-AA,

PDGF-BB, and 10% FBS. Our studies have further clarified the functional

significance of PDGFR-ß signaling in mediating its biological effects by examining the

activation of two of its downstream pathways, Akt and MAPKs. Compared to

wild-type fibroblasts, there was a decrease in PDGF-BB-induced phosphorylation of

Akt, ERK1/2, and JNK in PDGFR-ß∆/∆ fibroblasts. Interestingly, PDGF-BB-induced

phosphorylation of p38 MAPK was unaffected by depletion of PDGFR-ß. On the

other hand, PDGF-AA-induced phosphorylation of Akt, ERK1/2, JNK, and p38 was


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comparable between wild-type and PDGFR-ß∆/∆ fibroblasts. These results indicate that

PDGFR-ß is involved in PDGF-BB-induced phosphorylation of Akt, ERK1/2, and JNK.

Because the inhibition of these pathways was partial, we presume that PDGFR-α

mediates part of the signal triggered by PDGF-BB. In addition, our data suggest that

PDGFR-α mediates PDGF-AA-induced phosphorylation of Akt, ERK1/2, JNK, and

p38, and, unexpectedly, PDGF-BB-induced phosphorylation of p38 as shown in our

dose- and time-course studies.

In NIH3T3 fibroblasts, PDGFR-α, but not PDGFR-ß, mediates PDGF signals

leading to the activation of JNK, although both PDGFR-α and PDGFR-ß activate

ERK1/2 (29). This is different from our current finding that ERK1/2 and JNK are

possible downstream signaling molecules for both PDGFR-α and PDGFR-ß. The

apparent discrepancy may be due to the specific cell types examined and/or the

experimental procedures employed. A recent study reported that mouse embryonic

fibroblasts lacking both JNK1 and JNK2 exhibit much slower proliferation than

wild-type fibroblasts (30). In addition, studies with pharmacological inhibitors,

antisense oligonucleotides, and c-Jun-/- fibroblasts suggest that JNK plays a key role in

cell cycle progression (31). Furthermore, using dominant-negative mutants, Kawano

et al. (26) showed that ERK and JNK, but not p38, are involved in PDGF-BB-induced

mesangial cell proliferation. In the current studies, we found that PDGFR-ß∆/∆ cells

have reduced PDGF-BB-induced cell proliferation and decreased activation of ERK1/2

and JNK, but there was no effect on the activation of p38. Taken together, these

findings support the idea that, in dermal fibroblasts, PDGFR-ß mediates

PDGF-BB-induced proliferation through ERK1/2 and JNK but not through p38.


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The involvement of p38 and JNK in cell migration is controversial, possibly

because of different findings in various cell types and different methods to evaluate cell

migration. Ras-mediated activation of p38, but not JNK, has been reported to be

crucial for the migration of PDGFR-transfected porcine aortic endothelial cells (32).

In contrast, Javelaud et al. (33) showed that, in JNK-/- mouse embryonic fibroblasts and

human dermal fibroblasts, TGF-ß stimulates cell motility via the activation of JNK and

c-Jun. Consistent with these findings, our data showed that, in PDGFR-ß∆/∆ cells, the

phosphorylation of JNK was decreased, whereas that of p38 was unaffected.

Collectively, our results suggest that the decrease in PDGF-BB-induced

phosphorylation of JNK in PDGFR-ß∆/∆ fibroblasts may contribute to the lack of cell

movement and that normal activationof p38 is not sufficient to induce cell motility.

Ligand binding- or stress-induced activation of PDGFR-ß is known to mediate

Akt activation resulting in the protection of cells from apoptosis (34, 35). After

PDGF-BB stimulation, association of the activated Akt with IκB appears to be a

mechanism for protection of human skin fibroblasts from apoptosis (36). In PDGF-BB

stimulated wild-type cells, the phosphorylation of Akt sustained at high levels until 2 h

after stimulation, while the phosphorylation of MAPKs showed transient peak and

tended to decrease within 15-30 minutes after stimulation. The decrease of

phosphorylation of Akt was marked until 2 h in PDGF-BB stimulated PDGFR-ß∆/∆

fibroblasts. These features of Akt activation may imply that the Akt, rather than

MAPKs, contributed to the protection of PDGF-BB stimulated wild-type cells from

H2O2-induced apoptosis.

The phospholipid phosphatases SHIP2 and PTEN have been reported to be

negative regulators of PDGF signaling. These phosphatases lower the level of


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PI(3,4,5)P3, which results in a reduction in Akt activation (37, 17, 18). We found that

the overexpression of either SHIP2 or PTEN effectively inhibited PDGF-BB induced

phosphorylation of Akt in PDGFR-ß∆/∆ fibroblasts, whereas Akt phosphorylation was

only partially inhibited by expression of both SHIP2 and PTEN in wild-type fibroblasts.

In addition, SHIP2 is known to competitively inhibit Shc/Grb2 binding, which is

required for the activation of p21Ras and ERK1/2 (20). Consistent with this,

PDGF-BB-induced phosphorylation of ERK1/2 was inhibited by the expression of

SHIP2 but not PTEN in PDGFR-ß∆/∆ fibroblasts. These results indicate that the

PDGF-induced phosphorylation of Akt and ERK1/2 mediated by PDGFR-α, and not

PDGFR-ß, is preferentially regulated by phospholipid phosphatases. However, we can

not rule out the possibility that SHIP2 or PTEN alone are insufficient for the negative

regulation of PDGF-BB induced Akt phosphorylation when it is mediated by both

PDGFR-α and PDGFR-ß. In contrast to the involvement of SHIP2 and/or PTEN in

Akt and ERK1/2 activation, expression of SHIP2 and PTEN did not affect

PDGF-BB-induced phosphorylation of JNK or p38 MAPK in either wild-type or

PDGFR-ß∆/∆ fibroblasts. These results indicate that the phospholipid phosphatases are

a negative regulator of certain, but not all, PDGF-BB-induced intracellular signaling

events. Further studies are needed to determine whether phospholipid phosphatases

play a direct role in the negative regulation of PDGFR-α or PDGFR-ß mediated

biological effects.

In summary, we examined the biological effects and signaling stimulated by

PDGF in dermal fibroblasts in which PDGFR-ß was depleted by the

adenovirus-mediated Cre/loxP system. This system allowed us to assess the functional


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importance of PDGFR-ß and examine the role of its specific intracellular signaling

pathways. Our results indicate that 1) PDGFR-ß plays a crucial role in

PDGF-BB-induced cell proliferation, migration, and protection from apoptosis; 2)

PDGFR-α activation can not mediate fibroblast migratory response to fill in a scratch

wound although it supports a proliferative response; 3) PDGF-BB-induced

phosphorylation of Akt, ERK1/2, and JNK, but not p38, is involved in the biological

effects of PDGF-BB; 4) although PDGFR-α alone can mediate PDGF-BB-induced

activation of p38, it is not enough to mediate the cellular responses in fibroblasts,

including cell proliferation, migration, and protection from apoptosis ; and 5) SHIP2,

and not PTEN, preferentially plays a role as a negative regulator of PDGFR-α-mediated

phosphorylation of Akt and ERK1/2, but not of JNK or p38.


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Footnotes

We thank Dr. Kazuhito Fukui, Dr. Hiroshi Tsuneki, Mr. Yoichi Kurashige, Ms.

Takako Matsushima, and Ms. Sayaka Takakuwa (Toyama Medical and Pharmaceutical

University, Japan) for their technicalassistance. We also thank Prof. Elaine W.

Raines (University of Washington, U.S.A.) for critical reading of the manuscript. We

are also grateful to Dr. Izumu Saito (Tokyo University, Japan) for providing

AxCANCre (Cre) and AxCALNLZ (LacZ) adenovirus vectors, Dr. Junichi Miyazaki

(Osaka University Graduate School of Medicine, Japan) for the CAG promoter, and Dr.

Tomoichiro Asano (Tokyo University, Japan) for providing the recombinant adenovirus

expressing PTEN. This study was supported in part by a Grant-in-Aid for Scientific

Research from the Ministry of Education, Science, and Culture of Japan (16390114 and

12470053).

The abbreviations used are: BrdU, 5-bromo-2’-deoxyuridine; DMEM,

Dulbecco’s modified Eagle’s medium; ERK, extracellular signal- related kinase; ES

cells, embryonic stem cells; FBS, fetal bovine serum; JNK/SAPK, c-jun NH2-terminal

kinase/stress-activated protein kinase; MAPK(s), mitogen-activated protein kinase(s);

m.o.i, multiplicity of infection; neo-TK, neomycin-thymidine kinase; PDGF,

platelet-derived growth factor; PDGFR, platelet-derived growth factor receptor;

PI(3,4,5)P3, phosphatidylinositol (3,4,5)-triphosphate; PI3K, phosphatidylinositol

3-kinase; PTEN, phosphatase and tensin homolog deleted on chromosome 10; SHIP2,

Src homology domain 2-containing inositol phosphatase 2; WT, wild-type.


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Figure legends

Fig. 1. Construction of PDGFR-ß gene targeting vector and genotyping. A, A

diagram of the targeting vector. Three loxP sequences and neo-TK and diphtheria

toxin selection cassettes were inserted into the PDGFR-ß locus, and it was subcloned

into the p-Bluescript SK- plasmid. XbaI and SacI sites were artificially introduced in

the indicated positions. B, Southern blotting of ES cells after homologous

recombination using three different probes. Genomic DNA prepared from isolated ES

cells was digested with XbaI or SacI. C, Cre-mediated gene depletion in ES cells.

PDGFR-ßflox/+ ES cells were treated with Cre recombinase, and genomic PCR was

performed. The 410-bp band corresponds to the PDGFR-ß deleted allele (PDGFR-ß∆)

after Cre treatment. D, PCR-based genotyping of mice. Genomic PCR products

obtained from mouse tail DNA show distinct band patterns in PDGFR-ßflox/flox,

PDGFR-ß+/+, and PDGFR-ßflox/+ mice.

Fig. 2. Depletion of PDGFR-ß protein expression after Cre-adenovirus infection.

A, Wild-type and PDGFR-ßflox/flox fibroblasts were infected with the Cre-adenovirus at a

m.o.i. of 10 pfu/cell and were stimulated with 1 nM PDGF-BB for 5 min. Whole cell

lysates were immunoblotted with anti-PDGFR-α, anti-PDGFR-ß, or anti-Tyr857

phosphospecific PDGFR-ß antibody. B, The Cre-adenovirus- infected wild-type or

PDGFR-ßflox/flox fibroblasts were immunostained with anti-PDGFR-ß antibody (green).

Representative immunofluorescent staining for PDGF-ß is shown. Fibroblasts were

unambiguously identified by DAPI staining of nuclei (blue). C, PDGF-induced

phosphorylation of PDGFR after deletion of the PDGFR-ß gene. Wild-type and

PDGFR-ßflox/flox fibroblasts were infected with the Cre-adenovirus at a m.o.i. of 10

pfu/cell and serum-starved for 24 h. a, The cells were stimulated with various


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concentrations of PDGF-AA for 5 min. The cell lysates were immunoprecipitated

with anti-PDGFR-α antibody and immunoblotted with anti-phosphotyrosine antibody

(PY99) or anti-PDGFR-α antibody. b, Cells were stimulated with various

concentrations of PDGF-BB for 5 min. Proteins were immunoprecipitated from cell

lysates using anti-PDGFR-α antibody and then immunoblotting was performed with

PY99 or anti-PDGFR-α antibody. c, Cells were stimulated with various

concentrations of PDGF-BB for 5 min. Proteins were immunoprecipitated from cell

lysates using anti-PDGFR-ß antibody, and immunoblotting was performed with PY99

or anti-PDGFR-ß antibody.

Fig. 3. Effect of PDGFR-ß depletion on BrdU incorporation. Wild-type and

PDGFR-ßflox/flox fibroblasts were infected with Cre-adenovirus at a m.o.i. of 10 pfu/cell.

The cells were serum-starved for 24 h and then treated with 1 nM PDGF-AA, 1 nM

PDGF-BB, or 10% FBS. After 18 h, BrdU incorporation was assessed. Results

represent means ± S.D. of six separate experiments. ∗, p<0.05 vs. BrdU incorporation

in wild-type fibroblasts.

Fig. 4. Effect of PDGFR-ß deletion on wound scratch assay. Confluent wild-type

and PDGFR-ßflox/flox fibroblasts were infected with the Cre-adenovirus. The cells were

subjected to injury by scratching with a plastic pipette tip. The cells were then treated

for 48 h with 1 nM PDGF-BB (A-D), 10% FBS (E-H), or 1 nM PDGF-AA (I-L).

Large black dots marked the bottom surface of the culture dish to specify the site of

observation. Representative images are shown from three separate experiments.

Fig. 5. Effect of PDGFR-ß deletion on H2O2-induced apoptosis. Wild-type and

PDGFR-ßflox/flox fibroblasts were infected with the Cre-adenovirus at a m.o.i. of 10

pfu/cell and then were cultured in 12-well plates. After 24 h, 1 µM H2O2 and dye from


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the APOPercentageTM kit were added along with 1 nM PDGF-BB (A, B), 10% FBS (C,

D), or 1 nM PDGF-AA (E, F). After 1 h, the cells were washed twice with PBS and

then photographed. G, Results represent the means ± S.D. of three separate

experiments. ∗, p<0.05 vs. PDGF-BB incubation in wild-type fibroblasts.

Fig. 6. Effect of PDGFR-ß depletion on PDGF-induced phosphorylation of Akt.

Wild-type and PDGFR-ßflox/flox fibroblasts were infected with Cre-adenovirus at a m.o.i.

of 10 pfu/cell, and the cells were serum-starved for 48 h. The cells were then treated at

37ºC for 5 min with the indicated concentrations of PDGF-BB (A) or PDGF-AA (B), or

they were treated at 37ºC for the indicated times with 1 nM PDGF-BB (C) or 1 nM

PDGF-AA (D). Cell lysates were separated by 7.5% SDS-PAGE and immunoblotted

with anti-Thr308 phosphospecific Akt antibody or anti-Akt1 antibody. The relative

amount of phosphorylated Akt was determined by densitometry. Results represent

means ± S.D. of three separate experiments. ∗, p< 0.05 vs. Akt phosphorylation in

wild-type fibroblasts.

Fig. 7. Effect of PDGFR-ß depletion on PDGF-induced phosphorylation of ERK1/2.

Wild-type and PDGFR-ßflox/flox fibroblasts were infected with the Cre-adenovirus at a

m.o.i. of 10 pfu/cell, and the cells were serum-starved for 48 h. The cells were treated

at 37ºC for 5 min with the indicated concentrations of PDGF-BB (A) or PDGF-AA (B),

or they were treated at 37ºC for the indicated times with 1 nM PDGF-BB (C) or 1 nM


with anti-Thr202/Tyr204 phosphospecific ERK1/2 antibody or anti-ERK1/2 antibody.

The relative amount of phosphorylated ERK1/2 was determined by densitometry.

Results represent the means ± S.D. of three separate experiments. ∗, p< 0.05 vs.

ERK1/2 phosphorylation in wild-type fibroblasts.


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Fig. 8. Effect of PDGFR-ß depletion on PDGF-induced phosphorylation of JNK.






with anti-Thr183/Tyr185 phosphospecific JNK antibody or anti-JNK antibody. The

relative amount of phosphorylated JNK was determined by densitometry. Results

represent means ± S.D. of three separate experiments. ∗, p< 0.05 vs. JNK

phosphorylation in wild-type fibroblasts.

Fig. 9. Effect of PDGFR-ß depletion on PDGF-induced phosphorylation of p38.





PDGF-AA (D). The cell lysates were separated by 7.5% SDS-PAGE and

immunoblotted with anti-Thr180/Tyr182 phosphospecific p38 antibody or anti-p38

antibody. The relative amount of phosphorylated p38 was determined by densitometry.

Results represent means ± S.D. of three separate experiments.

Fig. 10. Effect of co-infection with PTEN and SHIP2 on PDGF-induced

phosphorylation of Akt, ERK1/2, JNK, and p38 in wild-type and PDGFR-ß∆ /∆

fibroblasts. Wild-type and PDGFR-ßflox/flox fibroblasts were co- infected with the

Cre-adenovirus, and LacZ-, PTEN-, or SHIP2-adenovirus at a m.o.i. of 10 pfu/cell.

The cells were serum-starved for 48 h and then treated at 37ºC for 5 min with 1 nM


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Gao et al.

PDGF-BB. The cell lysates were separated by 7.5% SDS-PAGE and immunoblotted

with PTEN or SHIP2 antibody (A), anti-Thr308 phosphospecific Akt and anti-Akt1

antibodies (B), anti-Thr202/Tyr204 phosphospecific ERK1/2 and anti-ERK1/2 antibodies

(C), anti-Thr183/Tyr185 phosphospecific JNK and anti-JNK antibodies (D), or

anti-Thr180/Tyr182 phosphospecific p38 and anti-p38 antibodies (E). The relative

amounts of phosphorylated Akt, ERK1/2, JNK, and p38 were determined by

densitometry. Results represent means ± S.D. of three separate experiments. ∗, p<

0.05 vs. Akt phosphorylation of Cre- and LacZ-infected wild-type fibroblasts in (B) and

vs. ERK1/2 phosphorylation of Cre- and LacZ-infected PDGFR-ßflox/flox fibroblasts in

(C). +, p< 0.05 vs. Akt phosphorylation in Cre- and LacZ-infected PDGFR-ßflox/flox

fibroblasts in (B).


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Figure1:

A:

C: D:

B:

12.8kb

7.3kb

5’ probe

(XbaI cut)

3’ probe

(SacI cut)

9.7kb

5.7kb

neo probe

(XbaI cut)

WT R.E WT R.E WT R.E

WT, wild type ES cells;

R.E, recombinated ES cells)

8.8kb

flox/flox +/+ flox/+

271bp329bp

271bp329bp

410bp500bp

400bp

300bp

M.W. flox/+

Transfect with Cre

to remove neo-TK

S

XS

S XX

neo-TK

loxP loxP loxP

E2 E3 E4 E5E6 E7 E8 E9 E10

5’probe 3’probe

neo probe

DT-A

SS XX SX

7.3kb 5.5kb

5.7kb4.0kb

p1 p2

p3 p4Internal

probe

WT locus

Targeting

vector

Floxed

locus

(X,XbaI site;S,SacI site;F,FspI site;H,HpaI site;p1-5,primers1-5)

F H

p5


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Figure 2

A: B:

WT PDGFR- ∆/∆

C:

a:

b:

c:

WT PDGFR- ∆/∆

∝ PDGFR I.P.

∝ PDGFR I.P.

∝ PY99 blot

∝ PDGFR blot

PDGF-AA (nM)

phospho-PDGFR-

PDGFR-

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

WT PDGFR- ∆/∆

∝ PDGFR I.P.

∝ PDGFR I.P.

∝ PY99 blot

∝ PDGFR blot

PDGF-BB (nM)

phospho-PDGFR-

PDGFR-

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

WT PDGFR- ∆/∆

∝ PDGFR I.P.

∝ PDGFR I.P.

∝ PY99 blot

∝ PDGFR blot

PDGF-BB (nM)

phospho-PDGFR-

PDGFR-

∝ PDGFR- blot

∝ PDGFR- blot

∝ Tyr857 phospho-PDGFR- blot

PDGFR-

phospho-PDGFR-

PDGFR-

- + - +Cre

Cell WT PDGFR- flox/flox


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Figure 3

∗∗

∗∗WT

∆/∆

Brd

UIn

corp

ora

tio

n

(Absorb

ance O

.D.)

0 PDGF-BB

1nM

PDGF-AA

1nM

10%FBS


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Figure 4A: WT 0h B: WT 48h

C: PDGFR- ∆/∆ 0h D: PDGFR- ∆/∆ 48h

PDGF-BB 1nM

I: WT 0h J: WT 48h

K: PDGFR- ∆/∆ 0h L: PDGFR- ∆/∆ 48h

PDGF-AA 1nM

E: WT 0h F: WT 48

G: PDGFR- ∆/∆ 0h H: PDGFR- ∆/∆ 48h

10% FBS


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Figure 6

A: B:

C: D:

PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

PDGFR- ∆/∆WT

∝ Akt blot

Akt

PDGFR- ∆/∆WT

p-Akt

∝ phospho-Akt blot

PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

Akt

Pho

sph

ory

lation

Arb

itra

ry U

nit

WT∆/∆

∗

∗∗

∗

0 0.04 0.2 0.4 1 2

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

Akt

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

p-Akt


Akt

Pho

sph

ory

latio

n

Arb

itra

ry U

nit

WT∆/∆

WT∆/∆

0min 5min 15min 30min 60min 120min

∝ Akt blot

PDGF-AA (nM)

PDGFR- ∆/∆WT

∝ Akt blot

Akt

PDGF-AA (nM)

PDGFR- ∆/∆WT

p-Akt


0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

Akt

ph

ospho

ryla

tion

Arb

itra

ry U

nit

WT∆/∆

WT∆/∆

0 0.04 0.2 0.4 1 2

PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ Akt blot

Akt


PDGF-BB (1nM) - + + + + + - + + + + +

p-Akt

0 5 15 30 60 120 0 5 15 30 60 120(min)

WT PDGFR- ∆/∆


WT∆/∆

Akt

Pho

sph

ory

lation

Arb

itra

ry U

nit

∗∗∗∗

∗∗ ∗∗∗∗


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Figure 7

B:

C: D:

A:

PDGF-AA (nM)

PDGFR- ∆/∆WT

∝ ERK1/2 blot

ERK1/2

PDGF-AA (nM)

p-ERK1/2

∝ phospho-ERK1/2 blot

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

ER

K1

/2 P

ho

sp

ho

ryla

tio

n

Arb

itra

ry U

nit

WT∆/∆

WT∆/∆

0 0.04 0.2 0.4 1 2

WT PDGFR- ∆/∆

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ ERK1/2 blot

ERK1/2

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

- + + + + + - + + + + +

(min)

p-ERK1/2


WT∆/∆

WT∆/∆

ER

K1

/2 P

ho

sp

ho

ryla

tio

n

Arb

itra

ry U

nit

WT PDGFR- ∆/∆


PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ ERK1/2 blot

ERK1/2


PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

- + + + + + - + + + + +

(min)

p-ERK1/2

ER

K1

/2 P

ho

spho

ryla

tion

Arb

itra

ry U

nit

WT∆/∆

WT PDGFR- ∆/∆

∗∗

∗∗∗∗

∗∗∗∗


PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

PDGFR- ∆/∆WT

∝ ERK1/2 blot

ERK1/2

p-ERK1/2


PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

WT PDGFR- ∆/∆

0 0.04 0.2 0.4 1 2

WT∆/∆

ER

K1

/2 P

ho

sph

ory

lation

Arb

itra

ry U

nit ∗∗

∗∗

∗∗ ∗∗∗∗


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Figure 8

A: B:

C: D:

PDGF-AA (nM)

PDGFR- ∆/∆WT

∝ JNK blot

PDGF-AA (nM)

∝ phospho-JNK blot

JNK

p46-JNK

p54-JNK

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

JN

K P

ho

sp

hory

lation

Arb

itra

ry U

nit

0 0.04 0.2 0.4 1 2

PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

PDGFR- ∆/∆WT

∝ JNK blot


PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

p46-JNK

p54-JNK

JNK

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

∗∗ ∗∗

∗∗ ∗∗

∗∗

0 0.04 0.2 0.4 1 2

JN

K P

ho

sp

hory

lation

Arb

itra

ry U

nit

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ JNK blot

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

- + + + + + - + + + + +

(min)


p46-JNK

p54-JNK

JNK

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

JN

K P

ho

sp

hory

lation

Arb

itra

ry U

nit


PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ JNK blot


PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

- + + + + + - + + + + +

(min)

p46-JNK

p54-JNK

JNK

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆


∗∗∗∗

∗∗

∗∗

∗∗

JNK

Ph

osp

hory

latio

n

Arb

itra

ry U

nit


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Figure 9

A: B:

C: D:

PDGF-AA (nM)

PDGFR- ∆/∆WT

∝ p38 blot

p38

PDGF-AA (nM)

p-p38

∝ phospho-p38 blot

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

p3

8 P

ho

spho

ryla

tion

Arb

itra

ry U

nit

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

0 0.04 0.2 0.4 1 2

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ p38 blot

p38

PDGF-AA (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

- + + + + + - + + + + +

(min)

p-p38


WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

p3

8 P

ho

spho

ryla

tion

Arb

itra

ry U

nit


PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

PDGFR- ∆/∆WT

∝ p-38 blot

p-38

PDGFR- ∆/∆WT

p-p38


PDGF-BB (nM) 0 0.04 0.2 0.4 1 2 0 0.04 0.2 0.4 1 2

p3

8 P

ho

spho

ryla

tion

Arb

itra

ry U

nit

WT∆/∆

WT∆/∆

0 0.04 0.2 0.4 1 2

PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

PDGFR- ∆/∆WT

- + + + + + - + + + + +

(min)

∝ p38 blot

p38


PDGF-BB (1nM)

0 5 15 30 60 120 0 5 15 30 60 120

- + + + + + - + + + + +

(min)

p-p38

p3

8 P

ho

spho

ryla

tion

Arb

itra

ry U

nit

WT∆/∆

WT PDGFR- ∆/∆



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Figure 10

A:

B: C:

D: E:

SHIP2

∝SHIP2 blot

PTEN

∝PTEN blot

PDGF-BB - + - + - + - + - + - + - + - +

LacZ PTEN SHIP2 P+S LacZ PTEN SHIP2 P+S

WT PDGFR- ∆/∆

p-p38

∝phospho-p38 blot

PDGF-BB - + - + - + - + - + - + - + - +


p38

∝p38 blot

PDGF-BB - + - + - + - + - + - + - + - +


WT PDGFR- ∆/∆

WT∆/∆

- + - + - + - +PDGF-BB

Cre+LacZ Cre+PTEN Cre+SHIP2 Cre+PTEN+SHIP2

PDGFR- ∆/∆WT

p3

8 P

ho

sp

ho

ryla

tio

n

Arb

itra

ry U

nit

p-54JNK

∝phospho-JNK blot

PDGF-BB - + - + - + - + - + - + - + - +


∝JNK blot

PDGF-BB - + - + - + - + - + - + - + - +


WT PDGFR- ∆/∆

p-46JNK

JNK

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

JN

K P

ho

sp

hory

lation

Arb

itra

ry U

nit

- + - + - + - +PDGF-BB


p-Akt

∝phospho-Akt blot

PDGF-BB - + - + - + - + - + - + - + - +


Akt

∝Akt blot

PDGF-BB - + - + - + - + - + - + - + - +


WT PDGFR- ∆/∆

WT∆/∆

WT∆/∆

WT PDGFR- ∆/∆

Akt

Pho

sph

ory

lation

Arb

itra

ry U

nit

- + - + - + - +PDGF-BB


+++

∗

p-ERK1/2

∝phospho-ERK1/2 blot

PDGF-BB - + - + - + - + - + - + - + - +


ERK1/2

∝ERK1/2 blot

PDGF-BB - + - + - + - + - + - + - + - +


WT PDGFR- ∆/∆

WT∆/∆

∗∗

WT PDGFR- ∆/∆

ER

K1

/2 P

ho

sph

ory

lation

Arb

itra

ry U

nit

- + - + - + - +PDGF-BB



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Masashi Kobayashi, Yo-Ichi Nabeshima and Masakiyo SasaharaSabit, Makoto Kawaguchi, Yoko Kurotaki, Maiko Naito, Tsutomu Wada, Shin Ishizawa,

Zhiyang Gao, Toshiyasu Sasaoka, Toshihiko Fujimori, Takeshi Oya, Yoko Ishii, Hemragulhealing

Deletion of the PDGFR-ß gene affects key fibroblast functions important for wound

published online December 6, 2004J. Biol. Chem.

10.1074/jbc.M413081200Access the most updated version of this article at doi:

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Documents

Deletion of the PDGFR-ß Gene Affects Key Fibroblast ... · phosphorylation of Akt, ... analysis of granulation tissue formation following subcutaneously ... resulting ES clones in