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J. Biol. Chem., Vol. 283, Issue 7, 3942-3950, February 15, 2008

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KLF2 Transcription Factor Modulates Blood Vessel Maturation through Smooth Muscle Cell Migration^*

From the Department of Molecular Genetics, Biochemistry and Microbiology, University of Cincinnati, College of Medicine, Cincinnati, Ohio 45267

Received for publication, September 20, 2007 , and in revised form, December 5, 2007.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Vasculogenesis, angiogenesis, and maturation are three major phases of the development of blood vessels. Although many receptors required for blood vessel formation have been defined, the intracellular signal transduction pathways involved in vascular maturation remain unclear. KLF2^–/– embryos fail to develop beyond 13.5 days because of a lack of blood vessel stabilization. The molecular mechanism of KLF2 function in embryonic vascular vessels is still largely unknown. Here we show a normal development pattern of endothelial cells in KLF2^–/– embryos but a defect of smooth muscle cells at the dorsal side of the aorta. This phenotype results from arrested vascular maturation characterized by the failure of mural cells to migrate around endothelial cells. This migration defect is also observed when platelet-derived growth factor-B (PDGF) controlled migration is studied in murine embryonic fibroblast (MEF) cells from KLF2^–/– animals. In addition, KLF2^–/– MEFs exhibit a significant growth defect, indicating that KLF2 is required to maintain the viability of MEF cells. The PDGF signal is mediated through the Src signaling pathway, and a downstream target of KLF2 is sphingosine 1-phosphate receptor 1. These studies demonstrate that KLF2 is required for smooth muscle cell migration and elucidate a novel mechanism involving communication between PDGF and KLF2 in vascular maturation.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Krüppel-like transcription factors (KLFs)² constitute a subfamily of zinc finger proteins (1, 2), and more than 15 family members have been identified (3) since first being classified by our laboratory as a family of mammalian zinc finger transcription factors (1). KLFs activate or repress genes involved in a number of cellular processes, many of which appear to be involved in late stages of cellular differentiation and development (4–6). Additional studies by our laboratory and others have demonstrated that KLF2 plays an important role in differentiation and function in a variety of cell types, including T lymphocytes, monocytes, lung cells, and adipocytes (7–10). A massive deficit of peripheral T cells in KLF2-null mice is because of a defect of thymocyte and T cell trafficking rather than T cell apoptosis as was originally thought (7, 11), suggesting an important role of KLF2 for T cell emigration. KLF2-deficient mice also show impaired lung development (6), which occurs at the late canalicular stage. Embryos of mice lacking KLF2 have a reasonably normal blood vessel network, and vasculogenesis as well as angiogenesis appear normal (9, 12). Nevertheless, the KLF2^–/– embryos do not survive beyond 12.5–14.5 days because of severe hemorrhage resulting from defects in the endothelial cell-mediated assembly and/or stabilization of the blood vessel wall (12). However, defects responsible for the loss of vascular integrity are unknown. The bleeding could be due to KLF2 deficiency in endothelial cells and/or mural cells.

The vertebrate vascular wall is composed of endothelial cells and mural cells, which are referred to as either pericytes or vascular smooth muscle cells (SMCs) (13). Vascular development is facilitated by complex patterns of gene expression, which involve a combination of vascular-specific genes and specialized genomic regulatory regions to coordinate expression of both vascular-specific and ubiquitous genes. Many transcription factors critical for vascular development have been identified by targeted disruption, often leading to pronounced vascular defects (14, 15). Angiogenesis is also affected by specific transcription factor ablation and serves as a model that recapitulates many of the molecular events occurring during vascular development (16). After initial endothelial tube formation, vessel maturation requires the subsequent recruitment of surrounding mesenchymal cells and their differentiation into vascular smooth muscle cells. This process involves the interaction of endothelial cells with mural cells and the release of specific growth factors. Several transcription factors have also been shown to be critical for this process. For instance, SMAD5 and MEF2C have been shown to be important in vascular development and in smooth muscle cell differentiation (14, 17). Targeted disruption of SMAD5 leads to vascular formation defects with embryonic lethality at day 10.5 to 11.5. These mice exhibit a prominent defect in vessel morphology, principally enlarged blood vessels with diminished numbers of SMCs. The loss of SMCs results from apoptosis of mesenchymal cells, which subsequently decreases the number of these cells available for maturation into SMCs.

Previous studies revealed an important role of KLF2 in blood vessel wall integrity (9, 12, 18). However, it is not known whether KLF2 controls SMC coverage of vessels in a cell-autonomous fashion by functioning directly in SMCs or indirectly through its activity in endothelial cells. Therefore, it is important to explore the molecular mechanisms of KLF2 in regulating embryonic vascular vessels. Studies described in this investigation are directed toward elucidating the role of KLF2 regulation in mural cells during embryonic vascular vessel development and maturation.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mice and Cell Culture—KLF2-null mice were produced as described previously (9). Briefly, Hprt-negative ES cells (El4TG2a) were targeted by homologous recombination to generate a 2.2-kb deletion that includes the promoter region, as well as exons 1 and 2 that code for the transcription activation domain and a part of the DNA binding domain, ensuring that no KLF2 mRNA and therefore no KLF2 protein is made in animals homozygous for the targeted allele. The chimeric mice were bred with Black Swiss for germ line competency testing. Resultant heterozygotes have been maintained on a mixed background (N18, C57BL6/129P2OlaHsd/nTac:NIHBS).

Because KLF2^–/– mouse embryos die around 12.5 days, we used KLF2^–/– and KLF2^+/+ embryos that were harvested at day 11.5. The preparation of MEFs was established by a modified protocol (10). In brief, fibroblasts were obtained by overnight trypsin digestion of eviscerated embryos and plating of cells in DMEM with 15% FBS. After first passage, MEFs were incubated with 10% FBS/DMEM. Cells used for this study were from passages 3–6. Jurkat/KLF2 Tet-On cells were cultured in RPMI 1640 medium supplemented with 10% FBS, 100 units/ml penicillin, and 100 µg/ml streptomycin. KLF2 induction was performed as described previously (19).

Histological Analysis and Immunohistochemistry—Embryos (E12.5 days), including yolk sac, were dissected into PBS and observed for gross abnormalities. The yolk sac was carefully removed, and the embryos were then fixed in 10% neutral buffered formalin overnight. The embryos were dehydrated through increasing concentrations of ethanol, transferred into xylene, and embedded in paraffin. The embryos were sectioned at 5 µm, and a series was stained with hematoxylin and eosin or Masson's Tri-Chrome. For immunohistochemistry, alternating sections were taken and baked at 60 °C for 1 h, cleared in xylene two times for 15 min, and hydrated through a descending alcohol series for 3 min each ending in distilled H₂O. The sections were rinsed once in PBS + 0.1% Tween 20 (PBS Tw). Antigen retrieval was accomplished using an antigen retrieval solution (Vector Laboratories) at 95–100 °C for 10 min. The sections were allowed to cool to room temperature in the citrate buffer for 1 h. The sections were then rinsed in PBS Tw. Endogenous peroxidase activity was blocked using 3% H₂O₂ in PBS for 10 min. Sections were rinsed several times in PBS Tw and were blocked for 1 h at room temperature with 2.5% normal goat serum in PBS with 0.3% Triton X-100. Sections were incubated overnight at 4 °C with monoclonal antibodies against smooth muscle -actin (mouse 1:1500; Sigma), CD34 (rat 1:50 antibody; Abcam, Inc.). Sections were rinsed, and appropriate biotinylated 2° antibodies (1:300) were applied for 45 min at room temperature followed by ABC for 45 min using Vector ABC kit (Vector Laboratories). Reactions were visualized using 3,3-diaminobenzidine (Vector Laboratories), counterstained with hematoxylin, and finally coverslipped using Vectamount.

Cell Growth Curve and Thymidine Uptake Assay—KLF2^+/+, KLF2^+/–, and KLF2^–/– MEFs were seeded in 24-well plates at 1 x 10⁴ cells per well, incubated at 37 °C, and treated with standard medium that was replaced every 2 days. The number of cells/ml was determined daily by removing cells from triplicate wells and counting in a hemocytometer. Dead cells were determined by the exclusion of trypan blue. KLF2^–/– ES cells and the parental cell line E14TG2a were recovered from frozen stocks onto mitomycin-treated MEFs and then passaged once onto gelatinized plates to eliminate feeders. All media contained leukemia inhibitory factor (Chemicon), 15% FBS, L-glutamine, β-mercaptoethanol, and nonessential amino acids in DMEM. Cells were trypsinized, counted, and plated at 2000/well in triplicate for each time point. Measurement of cellular proliferation was also determined by estimating [³H]thymidine incorporation into DNA. MEFs (1 x 10⁴) in 12-well dishes were starved for 24 h in serum-free medium and pulsed with 1 µCi/ml [³H]thymidine and 20 ng/ml PDGF 6 h before harvest. After 6 h, cells were harvested on glass fiber filters using a cell harvester. Disintegrations per min were determined by liquid scintillation counting.

Cell Migration Assay—Migration of mouse embryonic fibroblasts toward a PDGF-B gradient was examined according to the procedure described previously (20). Briefly, 3 x 10⁵ cells were made quiescent by incubation with DMEM and 0.4% fetal bovine serum for 48 h before PDGF-B treatment. A 0.1-ml aliquot of the cell suspension was added to the top chamber of tissue culture-treated Transwell polycarbonate membrane with 8-µm pores in 24-well plates. The lower Transwell compartment contained 0.6 ml of DMEM, 0.4% fetal bovine serum, and 0.2% bovine serum albumin with or without 20 ng/ml PDGF-B. After incubating for 4 h at 37°C, the upper surface of the filters was washed with phosphate-buffered saline. The cells were then fixed with methanol for 10 min at 4 °C followed by hematoxylin staining. The number of cells that migrated to the lower surface of each filter was counted in different high power fields at a magnification of 320. Experiments were performed in triplicate.

RNA Preparation, Northern Blot, and RT-PCR—Total RNA was isolated from cultured cells using TRIzol (Invitrogen) and treated with DNase I (Invitrogen). Northern blot analysis was performed as described previously (19). Briefly, an aliquot of total RNA (5 µg/lane) was size-fractionated by 1% agarose gel electrophoresis and transferred to a nylon membrane. Blots were immobilized by UV cross-linking and hybridized to [-³²P]dCTP-labeled cDNA probe from the 3'-untranslated region of mouse KLF2. The bands were detected using a Storm 860 PhosphorImager (GE Healthcare). For RT-PCR, total RNA (3.5 µg) was reverse-transcribed and the PCR conditions were as follows: initial denaturation at 95 °C for 10 min followed by up to 25 cycles of denaturation at 95 °C, annealing at 55 °C, and extension at 72 °C. PCR primer pairs were as follows: KLF2 (forward, 5'-GCACGGATGAGGACCTAAAC-3'; reverse, 5'-GTAGCTGCAAGTATGTGTGG-3'), GAPDH (forward, 5'-GGAGATTGTTGCCATCAACG-3'; reverse, 5'-GATGCAGGGATGATGTTCTG-3'), S1P1 (forward, 5'-CAGCTCAGTCTCTGACTATG-3'; reverse, 5'-CCTTGTTGGTCAGAGTGTAG-3'), S1P3 (forward, 5'-CCGGGAACATTACGATTACG-3'; reverse, 5'-GCCTCATCTTGATCATGGTC-3'), and KLF4 (forward, 5'-GGAACTCTCTCACATGAAGC-3'; reverse, 5'-CTGTGGCGGAATGTATACTG-3').

Transient Transfection and Promoter Luciferase Assay—A –287 S1P1 promoter fragment was generated by PCR, designated as S1P1luc. A site-directed mutagenesis generated mutation of a consensus KLF2-binding site was designated as S1P1mt. For S1P1 promoter activity, transient transfections of MEFs were plated in 6-well dishes at a density of 3 x 10⁵ cells/well, and transient transfections were performed the following day using FuGENE 6 (Roche Applied Science). The total amount of DNA used for the transfection assay per well was always held constant at 1 µg. The KLF2 expression plasmid was generated by subcloning KLF2 cDNA into pCDNA3.1. Cell lysates were prepared 48 h later, and luciferase activity was measured using a Monolight 3010 luminometer (Pharmingen) and expressed as relative light units using a luciferase assay kit (Promega). β-Galactosidase activity was measured with a commercially available kit (Promega). Promoter activity of each construct was expressed as the ratio of luciferase/β-galactoside activity. All transfections were performed in triplicate from three independent experiments.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Normal Endothelium in the Vasculature of KLF2^–/– Embryos—Previous studies (9, 12, 18) have shown that the loss of KLF2 is embryonic lethal because of internal hemorrhaging and deletion of endothelial KLF2, but the molecular mechanism(s) through which KLF2 regulates blood vessel wall stabilization is unknown. To address this issue, we first examined histological sections from KLF2^–/– mice. Up to day E11.5, KLF2^–/– embryos appeared phenotypically normal (data not shown). At E12.5, the KLF2^–/– embryos could be identified by their abnormal yolk sacs with reduced blood in the vasculature (data not shown). After removing the yolk sac, intraembryonic bleeding was evident in the abdomen of KLF2^–/– embryos (data not shown; as described previously by us (9) and Leiden and co-workers (12)). The morphology of the vasculature was characterized by immunohistochemical staining using CD34 (a marker for endothelial cells). Consistent with previous reports (12), a substantially normal vascular plexus was observed both in KLF2^–/– animals and age-matched control embryos (data not shown). Histological sections of hematoxylin and eosinstained E12.5 KLF2^–/– embryos revealed a normal capillary structure with both small and large vessels as well as normal capillary splitting (data not shown). In summary, these data confirmed that KLF2 is not essential for the formation of tubular endothelial structures (vasculogenesis) or for sprouting and splitting of existing vessels (angiogenesis) or remodeling of tubular endothelial structures into arteries and veins.

To further assess blood vessel wall morphology and stabilization at later stages of development, we analyzed the aorta of KLF2^–/– mice. Hematoxylin and eosin-stained saggital and transverse sections of aorta in KLF2^–/– embryos showed a normal pattern of the endothelial cell layer (Fig. 1) compared with wild type littermates. Wild type aorta contained an intact layer of CD34⁺ endothelial cells (Fig. 1 A, C and E), surrounded by well organized layers of differentiating smooth muscle cells (SMCs), which is confirmed by -actin (SMA) immunohistochemical staining (Fig. 3A). Since a previous study (12) reported endothelial cell necrosis in KLF2^–/– mice mediated the assembly and stabilization of the vessel wall, we closely examined the endothelial layer in small and large vessels in KLF2^–/– embryos. The KLF2^–/– aorta (Fig. 1, D and F) and other arteries as well as veins (data not shown) clearly contained CD34⁺ endothelial cells, confirming that KLF2 is not required for the specification of endothelial cell lineage. Moreover, the CD34+ staining patterns in KLF2^–/– embryos are similar to those in wild type embryos (Fig. 1, D and F), indicating a normal endothelium pattern in KLF2 knock-out mice. High magnification microscopy of hematoxylin and eosin staining in KLF2^–/– embryos showed no evidence of necrosis or degeneration of the endothelial cell layer (Fig. 1H and Fig. 2D), arguing against defects of the vascular wall being caused by endothelial cell necrosis.

View larger version (93K): [in this window] [in a new window]   FIGURE 1. Normal pattern of vasculature in the aorta of KLF2–/– embryos. A, hematoxylin and eosin-stained sagittal sections of wild type aorta; B, hematoxylin and eosin (H&E)-stained sagittal sections of KLF2–/– embryo aorta (E12.5); C, CD34-stained sagittal sections of wild type aorta; D, CD34-stained sagittal sections of KLF2–/– aorta; E (wild type) and F (knock-out), CD34-stained transverse sections of aortic arch. G (wild type) and H (knock-out), collagen type IV-stained transverse sections of aortic arch. Arrows indicate the dorsal side of aorta. Scale bar represents 5 µm.

View larger version (131K): [in this window] [in a new window]   FIGURE 2. Vascular smooth muscle defects in the KLF2–/– embryos. The hematoxylin and eosin staining pattern of the mice aorta. A (wild type) and B (knock-out), sagittal sections of aorta; C (wild type) and D (knock-out), transverse sections of aortic arch. Arrows indicate the dorsal side of aorta. Scale bar represents 5 µm.

Vascular Smooth Muscle Cell Defects in KLF2^–/– Embryos—We observe some smooth muscle cells in the vessels of KLF2^–/– embryos (Fig. 1B, Fig. 2, B and D, and Fig. 3, B and D), confirming that KLF2 is unlikely necessary for the specification of smooth muscle cell lineage. However, the smooth muscle layer in the KLF2 knock-out embryo is poorly developed although it shows no evidence of the necrosis or degeneration (Fig. 1B and Fig. 2, B and D). We did not observe any cuboidal smooth muscle cells in KLF2^–/– aorta. In contrast, KLF2^–/– SMCs displayed a sparse patchy morphology of smaller size and condensed nuclei compared with their wild type controls. In longitudinal sections of E12.5 control embryos stained with anti-SMA, four to five layers of smooth muscle cells evenly enveloped the entire dorsal aorta (Fig. 3A). In contrast, the aorta of KLF2^–/– mice was strikingly different. SMCs were deficient along the entire length of the dorsal surface examined (Fig. 3, B and D). Most of SMA-positive SMCs were present on the ventral side; few of them were on the dorsal side. As has been shown before (24), after the initial formation of the vascular plexus, the vessel maturation is stabilized through recruitment and differentiation of mural cells in vascular walls. SMCs initially appear on the ventral surface of the aorta in E10.5 embryos (25), followed by migration to the dorsal surface. By E12.5, the aorta is completely surrounded by SMCs. Any defect of migration or proliferation in this stage may result in the poor distribution of smooth muscle cells. These deficits could weaken the integrity of the blood vessel wall; therefore, the vessel wall cannot maintain blood flow and results in the rupture of the blood vessel. Thus, the deficiency of SMCs along the entire length of the dorsal surface in KLF2^–/– mice indicates a possible defect of SMC migration. The deficient SMCs at the dorsal side of KLF2^–/– embryos were also observed in the hematoxylin and eosin-stained sagittal and transverse sections of the aorta as shown in Fig. 2.

View larger version (136K): [in this window] [in a new window]   FIGURE 3. Deficient smooth muscle cells and extracellular matrix in KLF2–/– embryos. A, SM-actin-stained sagittal section of aorta in wild type embryos; B, SM-actin-stained sagittal section of aorta in KLF2 knock-out embryos; C, SM-actin-stained transverse section of aortic arch (ascending aorta) in wild type embryos; D, SM-actin-stained transverse sections of aortic arch (ascending aorta) in KLF2 knock-out embryos; E, trichrome-stained transverse section of aortic arch (ascending aorta) in wild type embryos; and F, trichrome-stained transverse section of aortic arch (ascending aorta) in KLF2 knock-out embryos. Arrows indicate the dorsal side of aorta. Scale bar represents 5 µm.

View larger version (22K): [in this window] [in a new window]   FIGURE 4. Defects of migration and proliferation in KLF2–/– MEFs. A, MEFs were derived from mouse embryos as described under "Experimental Procedures." The chemotactic response of wild type and knock-out primary MEFs to PDGF-B was examined under conditions containing (+PDGF) or lacking (–PDGF) PDGF-B at 20 ng/ml. B, growth curve of wild type and knock-out primary MEFs. KLF2+/+ MEFs and KLF2–/– MEFs were seeded in 12-well plates at 1 x 104 cells per well, incubated at 37 °C, and treated with complete DMEM. The medium was replaced every 2 days to maintain cell growth. C, growth curve of wild type and knock-out ES cells in the presence of 15% serum. D, thymidine uptake of wild type and knock-out MEFs following by PDGF stimulation. KLF2+/+ MEFs and KLF2–/– MEFs were starved in FBS-free medium for 24 h following by PDGF-B for 6 h. Each group was studied in triplicate (three wells).

Essential Role of KLF2 in Cell Migration—Progress has been made in defining the signaling pathways necessary for vessel maturation, including the recruitment and differentiation of mural cells. Peri-endothelial support cells, pericytes for capillaries, SMCs for small and large vessels, and myocardial cells in the heart, are recruited to cover the endothelial tubes. This provides both stability and modulatory functions. One essential molecule for the recruitment of mesenchymally derived mural cell precursors is PDGF-B. PDGF-B recruits cells expressing PDGF receptor-β to vessel walls (26). Disruption of the PDGF-B and/or PDGF receptor-β genes in mice leads to microvasculature aneurysms, lethal hemorrhaging, and edema in the perinatal stage because of a lack of pericytes (26, 27). Interestingly, PDGF-B knock-out mice display a similar hemorrhagic phenotype. Therefore, we hypothesized that PDGF may be activating a signal transduction pathway which in turn activates KLF2 transcription required for these mural cell movements.

To understand the potential involvement of KLF2 in PDGF-B-induced migration, MEFs from embryos were cultured and used to analyze mural cell migration. This system is widely used as a model to study vascular smooth muscle biology (28, 29). MEFs were obtained from wild type, KLF2^+/–, and KLF2^–/– embryos as described previously (10). As expected, PDGF-B induced a significant increase in the chemotaxis of wild type fibroblasts as well as heterozygous fibroblasts (Fig. 4A). But the KLF2^–/– fibroblasts displayed a significantly reduced migratory response. This demonstrates that KLF2 is required for PDGF-B-induced migration in these cells.

Involvement of KLF2 in Cell Proliferation—Because mural cell proliferation is also important for the maturation and stabilization of blood vessels, we further examined the role of KLF2 in MEF proliferation. Analyses of MEFs isolated from E12.5 KLF2 knock-out embryos indicated a striking reduction in cell proliferation when compared with KLF2^+/+ MEFs (Fig. 4B). As a control, we cultured KLF2^–/– ES cells, and these cells grew normally compared with wild type ES cells (Fig. 4C). We conclude that, although KLF2 is not required for the proliferation and survival of ES cells, it is required for the normal proliferation of MEFs. To study the kinetics of S phase entry of total asynchronous MEF populations, wild type MEFs and KLF2^–/– MEFs were labeled with [³H]thymidine following the stimulation of PDGF. During 6 h of incubation, the KLF2^–/– MEFs consistently displayed markedly less thymidine incorporation in S phase than WT MEFs (Fig. 4D). We conclude that the reduced proliferation of KLF2 knock-out MEFs is associated primarily with slower cell cycle progression.

PDGF Induces KLF2 through the Src Signal Pathway—To better understand how PDGF-B signaling regulates KLF2 expression, we measured KLF2 gene expression following PDGF-B stimulation (Fig. 5). As shown in Fig. 5, A and B, KLF2 is dramatically induced 30 min after PDGF-B stimulation in serum-starved MEFs, and the induction peaks at 1 h, indicating KLF2 is an early responsive transcription factor, which is consistent with its expression pattern in T cell activation and epithelial cell stimulation (30, 31). On the other hand, KLF2 signal is absent in knock-out MEF cells even after PDGF treatment. We also found that herbimycin A (32, 33), a potent protein-tyrosine kinase inhibitor that inhibits PDGF-B-induced Src/PLD activation, abrogated KLF2 transcription induced by PDGF-B (data not shown). Other inhibitors such as LY294002, SB202190, PD98059, and SB203580 (data not shown) did not suppress the KLF2 induction by PDGF-B, indicating that PDGF-B regulates KLF2 expression through the Src/PLD signaling pathway. Further PP1 treatment (a selective Src inhibitor) (28, 33, 34) also showed significant inhibition of KLF2 expression in a dose-dependent manner (Fig. 5C). These data suggest that PDGF regulates KLF2 expression through the Src signaling pathway.

View larger version (63K): [in this window] [in a new window]   FIGURE 5. PDGF induces KLF2 expression via the Src signaling pathway. A, Northern blot analysis of KLF2 gene expression. Early confluent KLF2+/+ MEFs, KLF2+/– MEFs, and KLF2–/– MEFs were incubated with 0.5% FBS/DMEM for 24 h followed by 30 ng/ml PDGF treatment. Total RNA of MEFs at different time points following stimulation with PDGF was isolated with TRIzol reagent and subjected to Northern blot analysis. An aliquot of total RNA (10 µg/lane) was hybridized to labeled KLF2 cDNA probe. Equal loading was confirmed by the fluorescence imaging (data not shown). B, multiplex PCR analysis of KLF2 gene expression. Early confluent KLF2+/+ MEFs were incubated with 0.5% FBS/DMEM for 24 h followed by 30 ng/ml PDGF treatment. A multiplex RT-PCR was carried out with primers of KLF2 and a house-keeping gene GAPDH simultaneously. C, Src-selective inhibitor PP1 specifically inhibits KLF2 induction in a dose-dependent manner. Synchronized MEFs were pretreated with Me2SO (DMSO) or specific Src inhibitor PP1 for 1 h, followed by treatment of 30 ng/ml PDGF for 1 h. Multiplex PCR was carried out with KLF2 and GAPDH primers.

We also used the tetracycline-inducible system (19) to examine whether KLF2 expression induces S1P1 gene expression. As shown in Fig. 6D, where KLF2 is induced by doxycycline in Jurkat/KLF2 cells, an increase in S1P1 expression is observed, suggesting KLF2 as a regulator of S1P1 gene expression. To examine the mechanisms regulating the expression of S1P1, we also analyzed the effect of KLF2 on S1P1 promoter activity via transient transfections in MEF cells. The S1P1 luciferase reporter construct was transfected into MEF cells in the presence or absence of KLF2 (Fig. 6E). In agreement with the mRNA data shown in Fig. 6, B and C, the S1P1 promoter activity was strongly induced, up to 4–5-fold by KLF2. Site-directed mutagenesis of the consensus KLF2-binding site resulted in a marked reduction in promoter activity, indicating this site is responsible for KLF2 gene regulation. Taken together, our data indicate that KLF2 is a positive regulator of the S1P1 gene, which is consistent with the role of KLF2 in T cell migration (11).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Our analysis of KLF2-null embryos demonstrated an incomplete vascular maturation resulting from a failure of mural cells (vascular smooth muscle cells and pericytes) to migrate to arterial walls and thus potentially stabilize them. Because disruption of the PDGF-B or PDGF receptor-β genes (27, 35, 36) in mice resulted in a similar phenotype and many polypeptide growth factors regulate SMC migration, especially PDGF-B, we speculated that interplay between PDGF-B and KLF2 might be required for cell migratory responses. Our in vitro studies demonstrate that KLF2 is required for PDGF-B induced migration in mouse embryonic fibroblasts, suggesting an important biological role of KLF2 in vascular remodeling. However, complete loss of migration toward PDGF-B was not observed in these mutant fibroblasts, indicating that KLF2 deletion does not disrupt all essential mechanisms of directed cell movement.

KLF2 induction by PDGF is strikingly dependent on the Src signaling pathway that is also critical for vascular formation (37–39), providing additional insight related to the upstream regulation of KLF2 expression. The finding that KLF2 in mural cells is involved in blood vessel maturation is very interesting as KLF2 is also critical for maintaining endothelial cell function.

In vitro and in vivo experiments with human, murine, and chicken endothelial cells indicate that KLF2 exhibits sustained induction under physiological levels of pulsatile and laminar shear stress (40–42). Interestingly, high levels of KLF2 expression in cultured human umbilical vein endothelial cells in response to fluid flow is down-regulated by treating cells with inflammatory cytokines such as tumor necrosis factor or interleukin-1β (43, 44). KLF2 overexpression in static cultures prevents the up-regulation of vascular cell adhesion molecule-1 and the endothelial adhesion molecule E-selectin in response to inflammatory cytokine treatment (44). Based on these findings, KLF2 is thought to be necessary and sufficient to coordinate the entire flow-activated transcriptional pathways involved in promoting vasodilatory, anti-coagulant, and anti-inflammatory phenotypes (45). Microarray analyses indicated that flow-mediated KLF2 induction is required for the regulation of a number of genes controlled by fluid shear stress. These findings implicate KLF2 as an important regulator of endothelial cell homeostasis and function. Thus, KLF2 plays roles in both mural cells and endothelial cells but has different functions in each cell. One is stabilization of vascular development in terms of mural cells; another is homeostasis in endothelial cells.

View larger version (58K): [in this window] [in a new window]   FIGURE 6. S1P1 is positively regulated by KLF2 in MEF cells. A, multiplex PCRs were carried out to analyze gene expression of KLF2, KLF4, S1P1, and S1P5 in MEFs. KLF2+/+, KLF2+/– and KLF2–/– MEFs were incubated in 10% FBS/DMEM to early confluent stage. RNAs were isolated and subjected to RT-PCR. The graph represents quantitative results of S1P1 expression levels via ImageQuant5.1. B, PDGF-B first induces KLF2 followed by S1P1 expression. KLF2+/+ MEFs were placed in the FBS-free medium following the treatment of PDGF-B for different time points. RT-PCR of KLF2, S1P1, and Vav3 was performed after cDNA was synthesized. C, expression pattern of KLF2 and S1P1 in primary SMCs followed by PDGF-BB treatment. Primary human smooth muscle cells were incubated without serum for 24 h followed by the PDGF-BB treatment for different times. RT-PCR of KLF2, S1P1, and GAPDH was carried out following cDNA synthesis. D, S1P1 expression is up-regulated by doxycycline (Dox)-induced KLF2 in the Jurkat Tet-On/KLF2 system. RNAs of J/KLF2 cells were isolated after treatment of control medium and medium containing doxycycline. Multiplex PT-PCR of S1P1 and GAPDH was performed. E, KLF2 transactivates the S1P1 promoter. MEFs were transfected with KLF2 plasmid or control plasmid (pcDNA3.1), S1P1 promoter luciferase reporter plasmid, and S1P1 mutation plasmid or pGL3 luciferase control, and β-galactosidase internal control.

In addition, because the control of SMC migration and its maintenance is brought about through changes in gene expression patterns, which in turn are critical for altering cellular phenotypes during vascular development and migration, delineation of transcriptional networks controlled by transcription factors becomes a major goal. An obvious question is what genes are regulated by KLF2. Therefore, the identification of "downstream targets" is important in understanding the functional role of this transcription factor. Among a number of potential candidate target genes, S1P1 is the most intriguing one. Sphingolipid signaling pathways have been implicated in many critical vascular events (54). S1P, a bio-active lipid found in high concentrations in platelets and blood, stimulates the endothelial differentiation gene family of G protein-coupled receptors. S1P stimulation triggers diverse effects, such as cell growth, survival, migration, and morphogenesis (55). S1P1 knock-out mice, which have a similar phenotype as KLF2 knock-out mice, exhibit blood vessel malformation, i.e. hemorrhage leading to embryonic lethality between E12.5 and E14.5 (56). Vasculogenesis and angiogenesis appear normal in the mutant embryos. Interestingly, however, vascular maturation is incomplete because of a deficiency of vascular smooth muscle cells/pericytes. The mechanism behind this deficiency is a failure of S1P1-mediated/S1P-induced migration (29, 56). Consistent with the role of KLF2 in T cell development, our data indicate that KLF2 is a positive regulator of S1P1, which is an important molecule for vessel formation and maturation.

Taken together, KLF2 plays crucial and unique roles in the vascular wall by functioning as an important transcriptional integrator of upstream and downstream signaling necessary for blood vessel formation. This may also underlie the defect in blood vessel stabilization that results in embryonic lethality of KLF2 homozygous null mice. Studying the impact of KLF2 upon the vasculature has the potential to benefit regenerative medicine and treatment of various vascular diseases, including atherosclerosis and hypertension.

	FOOTNOTES

 
^* This work was supported by National Institutes of Health Grant RO1 HL 57281. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ To whom correspondence should be addressed: Dept. of Molecular Genetics, Biochemistry and Microbiology, University of Cincinnati, College of Medicine, 231 Albert Sabin Way, Cincinnati, OH 45267. E-mail: jerry.lingrel{at}uc.edu.

² The abbreviations used are: KLF, Krüppel-like transcription factor; PDGF, platelet-derived growth factor-B; MEF, murine embryonic fibroblast; SMC, smooth muscle cell; ECM, extracellular matrix; SMA, smooth muscle -actin; RT, reverse transcription; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; S1P1, sphingosine 1-phosphate receptor 1; PBS, phosphate-buffered saline; ES, embryonic stem cell; S1P, sphingosine 1-phosphate.

	ACKNOWLEDGMENTS

 
We thank Dr. David Witte and Lisa McMillin for their helpful insights and technical support regarding histological analysis.

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