Journal of Leukocyte Biology Myeloid cells, immune suppression, tumor immunology
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Originally published online as doi:10.1189/jlb.0304175 on July 18, 2006

Published online before print July 18, 2006
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(Journal of Leukocyte Biology. 2006;80:608-620.)
© 2006 by Society for Leukocyte Biology

Activation of sterol regulatory element-binding proteins (SREBPs) is critical in IL-8-induced angiogenesis

Min Yao*, Rui-Hai Zhou{dagger}, Melissa Petreaca*, Lei Zheng*, John Shyy{dagger} and Manuela Martins-Green*,1

* Department of Cell Biology and Neuroscience and
{dagger} Division of Biomedical Sciences, University of California, Riverside

1 Correspondence: Department of Cell Biology and Neuroscience, Spieth Hall, 900 University Ave., University of California Riverside, Riverside, CA 92521. E-mail: manuela.martins{at}ucr.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Angiogenesis is essential in many physiological and pathological processes and can be stimulated by many different factors. To better understand and to manipulate this process more effectively, it would be beneficial to identify molecules common to the signaling pathways stimulated by different classes of angiogenic factors. Sterol regulatory element-binding proteins (SREBPs) are involved in the metabolism of cholesterol and fatty acids, molecules that are critical in membrane biology, and hence, many of the processes involved in angiogenesis. Here, we show that angiogenic factors of different families, such as basic fibroblast growth factor, thrombin, and interleukin (IL)-8, stimulate SREBP activation, whereas nonangiogenic factors, such as transforming growth factor-ß1, do not. We focused our detailed studies on IL-8 in vitro and in vivo, as this chemokine is also involved in inflammation and hence, has the potential to be critical in inflammation-induced angiogenesis, a process common to many diseases. Using human microvascular endothelial cells, a rabbit skin wound-healing model, and the chorioallantoic membrane assay, we show that IL-8 stimulates the activation of SREBP-1 and -2, and this activation is specific and receptor-mediated. SREBP activation leads to activation of RhoA through 3-hydroxy-3-methylglutaryl CoA reductase. RhoA is a small guanosinetriphosphatase, important in cytoskeletal functions, which in turn, are critical in many of the cellular processes needed for angiogenesis. Given that diverse, angiogenic factors use different cell-surface receptors, identification of this common step in the signal-transduction pathway provides the opportunity for novel approaches for prevention and treatment of diseases involving abnormal angiogenesis.

Key Words: cytokines • VEGF • bFGF • thrombin • neovascularization


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Interleukin 8 (IL-8) is a member of the CXC chemokine family. The biological activities of this chemokine are mediated by binding to two G-protein-coupled seven-transmembrane domain receptors, CXC chemokine receptor 1 (CXCR1) and CXCR2. IL-8 was originally identified as an activator and chemoattractant of leukocytes, in particular, of neutrophils [1 , 2 ], but more recently, it has been shown to be angiogenic [3 4 5 6 7 ]. Because of its inflammatory and angiogenic activities, IL-8 has been recognized as important in physiological or pathological processes, such as normal and pathological healing and a variety of inflammatory diseases as well as tumor growth and metastasis [8 9 10 ].

Angiogenesis, the process whereby new blood vessels are formed from pre-existing microvasculature, is a crucial event in many physiological and pathological processes. The early stages of angiogenesis involve multiple steps, including destabilization of the endothelium, leading to increased blood vessel permeability and flexibility; proliferation of endothelial cells (ECs); and EC migration, sprout elongation, and tube formation. Although IL-8 clearly possesses the ability to stimulate angiogenesis in diverse processes, the molecular mechanisms by which this stimulation occurs remain unclear. In light of the importance of the cell membrane in mediating inside-out and outside-in signaling pathways and the importance of the membrane biophysical properties in the regulation of EC functions, such as proliferation and migration, it is tempting to propose that molecules that regulate cell membrane biosynthesis or homeostasis might be critical for the regulation of angiogenesis. We have previously shown that sterol regulatory element-binding proteins (SREBPs), transcription factors that govern the cellular lipid biosynthesis and homeostasis, are important in vascular endothelial growth factor (VEGF)-induced angiogenesis [11 ]. In the current study, we hypothesize that CXCR1/2-mediated activation of SREBPs is critical in IL-8-induced angiogenesis and that SREBP activation may be a common pathway in angiogenesis in response to multiple types of angiogenic factors.

SREBPs include SREBP1 and -2 and are encoded by two genes, srebp1 and srebp2 [12 ]. SREBP1 consists of two isoforms, SREBP1a and SREBP1c, as a result of alternative splicing [13 , 14 ]. SREBP1 is responsible for the biosyntheses of cholesterol and fatty acids, whereas SREBP2 primarily mediates cholesterol biosynthesis [15 , 16 ]. All three SREBP proteins are basic helix-loop-helix-Zip transcription factors normally localized in the endoplasmic reticulum (ER) and are synthesized as inactive precursors. During times of sterol deficiency, each SREBP forms a complex with the SREBP cleavage-activating protein (SCAP), which binds the C-termini of SREBPs and escorts them from the ER to the Golgi apparatus, where two specific proteinases, Site 1 protease (S1P) and S2P, cleave SREBP proteins and release their N-termini [17 ]. The N-terminus, also called the mature form or active form, enters the nucleus and activates transcription of more than 30 genes, including the low-density lipoprotein receptor (LDLR), 3-hydroxy-3-methylglutaryl CoA reductase (HMGCoAR), and fatty acid synthase (FAS), which participate in the synthesis or uptake of cholesterol and fatty acids [17 , 18 ]. In a negative-feedback loop, cholesterol prevents the movement of the SREBP complex to the Golgi, thus blocking the activation of SREBPs and the expression of downstream genes [19 ].

This study focused on the role of the SREBPs in IL-8-induced angiogenesis. We found that IL-8 stimulates RhoA downstream from SREBP via HMGCoAR regulation and that blockage of SREBP activity inhibits angiogenesis, but rescue of RhoA activity restores the process. In addition, we determined SREBP activation is a common pathway in ECs in response to other angiogenic factors, including basic fibroblast growth factor (bFGF) and thrombin. Identification of the signal transduction pathways for these angiogenic factors belonging to different families has important implications for modulation of angiogenesis during wound-healing, inflammation, tumor growth, and metastasis [3 , 9 , 20 21 22 ].


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Materials
Primary human microvascular ECs (hMVECs) and EC growth medium (EGM)2-MV were purchased from Cambrex (San Diego, CA). The following antibodies were obtained from various suppliers: anti-SREBP1 (2A4), anti-SREBP1 (K-10), anti-SREBP2 (N-19), and horseradish peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology, CA); anti-SREBP2 (BD PharMingen, San Diego, CA); anti-platelet-endothelial cell adhesion molecule-1 (PECAM-1) and anti-IL-8 (R&D Systems, Minneapolis, MN); anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH; RDI Research Diagnostics, Flanders, NJ); and rhodamine- or fluorescein isothiocyanate (FITC)-conjugated secondary antibodies (Zymed Laboratories, San Francisco, CA). Anti-rabbit IL-8 was a gift from Dr. Akihiro Matsukawa (Kurume University, Fukuoka, Japan), and anti-hCXCR1 and anti-CXCR2 were gifts from Genentech (S. San Francisco, CA). DF1681B (CXCR1, -2 inhibitor) was a gift from AMSA (Italy). CXCR2 inhibitor was purchased from CalBioChem (San Diego, CA). 25-Hydroxycholesterol (25-HC), thrombin, geranylgeranyl pyrophosphate (GGPP), and farnesyl pyrophosphate (FPP) were purchased from Sigma Chemical Co. (St. Louis, MO). bFGF was purchased from Upstate Biotech. Inc. (Lake Placid, NY) and transforming growth factor-ß (TGF-ß), from Peprotech (Rocky Hill, NJ).

Cell culture
Primary hMVECs derived from human lung microvessels or pooled from human neonatal dermal microvessels were cultured with EGM2-MV medium containing growth supplements. Cells at Passages 4–7 were used in these experiments.

Immunoblot analysis
Cells were lysed using 150 mM NaCl radioimmunoprecipitation assay buffer containing a variety of protease inhibitors. Total protein extracts were analyzed using 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), followed by immunoblotting using various primary antibodies, as indicated, and were reprobed with anti-GAPDH or histone 2B (H2B) antibodies as loading controls. The intensities of the protein bands were quantified by densitometry, using Scion Image for Windows (Scion Corp., Frederick, MD).

Quantitative real-time polymerase chain reaction (PCR)
Total RNA was isolated from hMVECs using the RNeasy kit (Qiagen, Valencia, CA). mRNA expression of SREBPs and their target genes was quantified by real-time PCR using an ABI Prism 7700 sequence detection system (Perkin-Elmer Applied Biosystems, Wellesley, MA) and the reaction reagent SYBR Green II Master Mix kit (Stratagene, La Jolla, CA) according to the manufacturer’s protocol. GAPDH mRNA was used as an internal control to normalize gene-specific mRNA. The levels of mRNA in IL-8-treated cells were normalized to the controls, which were set at 1. Primer sets used for SREBP1a, SREBP1c, SREBP2, LDLR, FAS, HMGCoAR, and GAPDH were described previously [11 ].

Cell transfection with RNA interference
hMVEC grown to 90% confluence were transfected with small interference RNA (siRNA) duplexes (Dharmacon, Lafayette, CO), specific for SCAP (SCAPi; 5'-AACCUCCUGGCAGUAUGUA-3') or specific for the plasmid GL3-luciferase gene (pGL3i; 5'-CTTACGCTGAGTACTTCGA-3') as a control using Oligofectamine (Invitrogen, Carlsbad, CA), according to the manufacturer’s protocol. Twenty-four hours after transfection, the cells were treated and then analyzed at the indicated times.

Chromatin immunoprecipitation (ChIP) assay and nested PCR
hMVEC were cultured to 95% confluence. Following the treatment of IL-8 at various times, the cells were washed twice with phosphate-buffered saline (PBS) and cross-linked with 1% formaldehyde for 15 min. Cells were rinsed with ice-cold PBS twice, collected by scraping into 100 mM Tris-HCl (pH 9.4) containing 10 mM dithiothreitol, and incubated for 15 min in collection buffer, centrifuged, and washed with 1 ml ice-cold PBS, 1 ml Buffer I (0.25% Triton X-100, 10 mM EDTA, 0.5 mM EGTA, 10 mM HEPES, pH 6.5), and 1 ml Buffer II (200 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 10 mM HEPES, pH 6.5). The cells were resuspended in lysis buffer (1% SDS, 10 mM EDTA, 150 mM Tris-HCl, pH 8.1, 1x protease inhibitor cocktail), sonicated three times, 10 s each, and centrifuged. Supernatants were collected and diluted in IP incubation buffer (1% Triton X-100, 2 mM EDTA, 150 mM NaCl, 20 mM Tris-HCl, pH 8.1). Anti-SREBP1 or anti-SREBP2 antibodies (1 µg) were added into the samples overnight at 4°C to perform IP. After IP, protein A-Sepharose was added and incubated for 1 h at 4°C. Precipitates were washed three times for 10 min each in transmissible spongiform encephalopathy I (TSE I; 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl, pH 8.1, 150 mM NaCl), TSE II (0.1 SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl, pH 8.1, 500 mM NaCl), and Buffer III (0.25 M LiCl, 1% Nonidet P-40, 1% deoxycholate, 1 mM EDTA, 10 mM Tris-HCl, pH 8.1), extracted three times with 1% SDS, 0.1 M NaHCO3, and heated at 65°C overnight to reverse the formaldehyde cross-linking. DNA fragments were purified with a QIAquick gel extract kit (Qiagen). PCR was performed with primers for the human HMGCoAR promoter region –222 to –10 base pairs, which contains the SREBP binding site (forward: 5'-TGCTGGGACTCGAACGGCTAT-3'; reverse: 5'-TTACGCACGCTCGGAGCTGGAC-3') for 25 cycles (melting at 94°C for 30 s, annealing at 57°C for 30 s, and extension at 75°C for 30 s per cycle). Nested PCR was then performed by using 1 µl of the 100 µl PCR reaction as template and by performing an additional 25–30 cycles of amplification. The final PCR products were resolved on a 3% agarose gel, stained with ethidium bromide, and imaged with a charged-coupled device camera. The gel pictures were reversed to show the DNA as dark bands to better illustrate the differences.

Rabbit skin wound-healing model
New Zealand white rabbits of 4 months old (Western Oregon Rabbit Co., Philomath) were used to create a partial-thickness skin wound-healing model. All procedures were approved by the University of California Riverside Institutional Animal Care and Use Committee (AUP #A-0204011-1). Briefly, the rabbits were sedated and anesthetized with acepromazine (1 mg/kg, IM), ketamine (50 mg/kg, IM), and xylazine (5 mg/kg, IM). After shaving the rostral-dorsal hair, two partial-thickness skin wounds in the rostral-dorsal area were made in each rabbit, using a 5-mm Acupunch (Acuderm, Ft. Lauderdale, FL). At different time-points following the initial injury, samples of the wounded areas were collected using a 10-mm Acupunch. Normal, unwounded skin tissue samples were also removed and were used as controls. For inhibition of IL-8 or its receptors, various antibodies or inhibitors were applied to the wounds each day until sample harvest. Half of the specimens were extracted for Western blotting analysis, and the rest were fixed and embedded for frozen sectioning. The sections were used for immunofluorescence detection.

Cell infection with SREBP2 N-terminal [SREBP2(N)]-containing adenovirus (Ad)
Recombinant Ad-HA-SREBP2(N), encoding functional human SREBP2 N-terminal, was prepared as described previously [23 ]. To infect the cells, 30 µl media containing virus was added to confluent hMVECs and incubated for 12 h. The infected cells were then incubated with fresh growth medium for another 12 h and replated into 24-well plates for tube formation assay. Ad-ß-galactosidase (ß-gal) was used as a control for infection.

Cell tranfection with a vector expressing C3
Plasmid containing enhanced green fluorescent protein (pEGFP)-C1, encoding Clostridium botulinum C3 exoenzyme, was a gift from Dr. Miguel Angle Del Pozo (University Autonoma Madrid, Spain) [24 ]. The exoenzyme C3 was originally made by Dr. Andrew Hall (University College, London, UK) and cloned into pEGFP-C1 (BD Clontech, Inc., Palo Alto, CA). Lipofectin® transfection reagent (Invitrogen) was used for endothelial cell transfection with pEGFP-C1-C3 DNA or with pEGFP-C1 (as a control) according to the manufacturer’s protocol. Twenty-four hours after transfection, the cells were replated and treated for tube formation assay.

Endothelial cells tube formation assay
Matrigel (Becton Dickinson, Bedford, MA) was added to a 24-well plate for 1 h at 37°C. hMVECs were transfected with SCAPi or GL3i or preincubated with 25-HC (5 µg/ml) for 24 h. For the rescue experiments, we also added FPP (10 µmol/L) and GGPP (10 µmol/L) at the same time. The cells were plated in 24-well plates at a density of 3 x 104 cells per well and treated with IL-8 (50 ng/ml) or without IL-8 and were then incubated for 24 h. Tube formation was observed under an inverted-phase contrast microscope (Nikon, Tokyo, Japan), and the images were captured with a digital camera system after 24 h of culture.

Angiogenesis assay in chicken chorioallantoic membranes (CAM)
This assay was performed in the CAM as described previously [25 ]. Briefly, fertilized eggs purchased from Hy-line Inc. (Lakeview, CA) were incubated at 37°C. Four days after fertilization, a window was opened on the eggshell to drop the embryo away from the egg-shell membrane; this window was covered with transparent tape until Day 10, allowing blood vessel development. Pellets were prepared using 20 µl 1% methylcellulose containing the following treatments: dimethyl sulfoxide (DMSO; control) alone, DMSO + IL-8, DMSO + IL-8 + 25-HC, or 25-HC + IL-8 + GGPP + FPP; they were applied onto the CAM at Day 10. After incubation for an additional 4 days, the CAMs were fixed in situ and then excised from the eggs. Images were captured, and the numbers of blood vessel branches in the pellet area were counted. In separate experiments, areas of the CAMs, which were covered by the pellet, were harvested at Day 14 and were pooled for protein extraction and immunoblotting.

Immunolabeling and fluorescence microscopy
hMVECs on transwell membranes or frozen skin-tissue sections were washed with PBS, fixed with 4% paraformaldehyde for 20 min, permeablilized with 0.1% Triton X-100, and blocked with serum of the species for the secondary antibodies for 30 min. The specimens were incubated with primary antibodies for 2 h at room temperature and then washed three times with 0.1% bovine serum albumin in PBS. FITC or rhodamine-conjugated secondary antibodies were applied to the specimens for 1 h in the dark. After washing three times, the specimens were mounted with Vectashield (Vector Laboratories, Burlingame, CA); the images were viewed, and pictures were taken using a Nikon TE300 fluorescence microscope.

Permeability assay
hMVECs (1x105) in 100 µl EGM2-MV media were plated onto the upper chamber of a transwell system containing 3 µm pore size membranes coated with a thin layer of Matrigel. The cells were then incubated for 24 h, at which point, more cells were added into the upper chamber and incubated for another 24 h to allow the formation of a tight monolayer before further treatments and analysis. FITC-Dextran (3 kD) and IL-8, at a respective concentration of 10 µg/ml and 50 ng/ml, were added to the lower chamber of the transwell system. Aliquots of 10 µl media were removed from the upper chamber of the transwell system at the indicated time-points, and the fluorescence intensity was quantified using a fluorimeter (VictorTM 1420, Perkin-Elmer Life Sciences, Boston, MA) with excitation at 485 nm and emission at 535 nm. The average and standard deviation were calculated from triplicate transwells.

Cell proliferation assay
The bromodeoxyuridine (BrdU) incorporation assay was used for the hMVEC proliferation studies according to the manufacturer’s protocol. Briefly, hMVECs were transfected with siRNA or GL3i pretreated with 25-HC (5 µg/ml) for 24 h, replated into 96-well plates, and then treated with IL-8 (100 ng/ml) and BrdU for 24 h. BrdU incorporation was measured by enzyme-linked immunosorbent assay.

Cell migration assay
The cloning ring assay was used to create a model for outward migration of endothelial cells [26 ]. hMVECs (1x105) in 100 µl medium were plated within a 6-mm diameter cell-cloning ring, which was set inside a 35-mm cell culture dish. Four hours after plating, the cylinder was removed, the edge of cells was marked, and migration was measured at the indicated times by determining the distance between the migrating front of the endothelial cells and the marked areas. Cells were treated with IL-8 every 24 h.

Chemotaxis assays
Chemotaxis assays were also performed with the hMVECs, according to methods described previously [27 ]. Briefly, transwell polycarbonate membranes of 8 µm pore size (BD Biosciences, San Jose, CA) were coated on both sides with 50 ng/ml type I collagen (Sigma Chemical Co.). hMVECs (1x104) in 100 µl medium were seeded on the underside of the transwell membrane, by inverting the transwell insert, and were allowed to adhere for 30 min. The transwells were then mounted into the wells of a 24-well plate, such that the seeded cells were facing the lower chamber. Media were then added to the upper and lower chambers of the transwell system. IL-8, at a concentration of 50 ng/ml, was added to the upper chamber for 3 h at 37°C. The cells remaining on the underside of the membrane after 3 h were removed using a cotton swab, and the membranes were fixed and stained with 2% toludine blue in paraformaldehyde. The number of cells on the upper side of the membrane was counted at 10x magnification, and the cell numbers in 10 fields were counted and averaged.

Rho-guanosine 5'-triphosphate (GTP) pull-down assay
To determine the levels of active RhoA, hMVECs subjected to various treatments were lysed with cold lysis buffer (50 mM Tris, pH 7.2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 500 mM NaCl, 10 µg/ml each leupeptin and aprotinin, and 1 mM phenylmethylsulfonyl fluoride). Whole cell lysates were centrifuged, and the supernatants were transferred into tubes containing glutathione S-transferase (GST)-T cell receptor ß diversity (TRBD; 20–30 µg) beads (kindly provided by Dr. Kathryn DeFea) and incubated for 60 min with gentle shaking. The beads were washed and boiled in loading buffer, and the lysates were subjected to SDS-PAGE followed by immunoblotting using the RhoA antibody to determine levels of the active RhoA precipitated by the GST-TRBD beads. Crude cell lysates were also analyzed using SDS-PAGE and immunoblotting with RhoA antibody to ensure equal input of total RhoA.

Statistical analyses
Data were analyzed by two-tail Student’s t-test. The results were shown as mean ± SD from at least three independent experiments. Values of P < 0.05 were considered to be statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
SREBPs are activated by multiple angiogenic factors
We used primary hMVECs to examine whether SREBPs are activated by different types of angiogenic factors. The precursor forms of SREBP1 and -2 and their cleavage products (active forms) were increased by bFGF (Fig. 1A ), IL-8 (Fig. 1B) , and thrombin (Fig. 1C) . In contrast, TGF-ß1, which does not stimulate angiogenesis directly, did not cause any changes of SREBPs (Fig. 1D) . Overexposure of the immunoblots resulted in a variety of nonspecific bands, but SREBPs are not changed. For detection of SREBP1, we used an antibody recognizing the SREBP1 N-terminus, which can detect the precursor and the active forms of SREBP1. An antibody to the SREBP2 C-terminus was used to detect the precursor and cleaved C-terminus of this molecule, as the antibody to the N-terminus does not label this protein effectively in immunoblot analysis. The modulation of SREBPs was demonstrated by increased levels of the precursor protein and the active forms of these transcription factors. Our data suggest that SREBPs may play an important role in mediating the angiogenic effects of different classes of angiogenic factors.


Figure 1
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  		Figure 1. Different types of angiogenic factors activate SREBPs in hMVECs, which were cultured and treated with cytokines and growth factors; the cell lysates were harvested at indicated time-points and analyzed by immunoblotting for SREBP1 or SREBP2. P = Precursor forms of the SREBPs; N = N-terminus of SREBP1 (active form); and C = C-terminus of SREBP2 (indicates release of the active N-terminus). hMVECs were treated with (A) bFGF {15 ng/ml in 0.5% 3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate with PBS}, (B) IL-8 (50 ng/mL), (C) thrombin (2 U/ml), and (D) TGF-ß1 (15 ng/ml). Bar graphs show the average band density of the N-terminus of SREBP1 and the C-terminus of SREBP2. Mean ± SD, n = 4. *, P < 0.05. GAPDH was used to detect equal loading. Results are representative of three independent experiments. M.W., Molecular weight.

 
IL-8 stimulates expression of SREBPs and of genes downstream of these transcription factors
We showed previously that VEGF-induced angiogenesis involves activation of SREBPs. As VEGF acts through a tyrosine kinase receptor, we found it important to study in detail one of the angiogenic factors, which activates a different type of receptor (i.e., a seven-transmembrane receptor), to determine whether the processes of activation of SREBPs are the same or similar. We chose to study IL-8, a chemokine that is important in inflammation-induced angiogenesis. Analysis of hMVECs showed that the mRNA levels of SREBP1 and SREBP2 were increased after IL-8 treatment (Fig. 2A ). SREBP1c was strongly expressed, whereas SREBP1a was barely detected (data not shown). Therefore, we focused on SREBP1c, and henceforth, in this paper, SREBP1 refers to the 1c isoform only. In addition to SREBP gene expression, we examined the expression of several target genes of SREBPs and found that the levels of mRNAs encoding HMGCoAR, LDLR, and FAS are increased significantly upon stimulation of hMVECs by IL-8 and that this increase occurs in a time-dependent manner (Fig. 2B) . Furthermore, to strengthen these results, we performed ChIP on IL-8-treated hMVECs using antibodies specific for the N-terminus of SREBP1 or SREBP2. These assays showed that the chromatin fragments immunoprecipitated by the SREBP antibodies contain the promoter regions of HMGCoA reductase genes, to which these transcription factors bind (Fig. 2C) .


Figure 2
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  		Figure 2. IL-8 stimulates SREBP1 and -2 gene expression and genes downstream form SREBP. (A, B) hMVECs were treated with IL-8 (50 ng/mL) for different times as indicated, and total RNA was extracted. Expression of SREBP1, SREBP2, HMGCoAR, LDLR, and FAS was quantified by real-time PCR. Data show the mean level of mRNA encoding these genes relative to that of GAPDH mRNA from triplicate experiments and normalized to the nontreated cells (Control). **, P < 0.001. (C) ChIP assays: Chromatin was prepared from hMVECs treated with IL-8 (50 ng/mL) for indicated times and immunoprecipitated with antibodies specific for the N-terminus of SREBP1 or -2. The immunoprecipitated DNA was amplified using pairs of primers that cover the regions of the HMGCoAR gene promoter. Input is control. Images were reversed to show the DNA as dark bands to better illustrate the differences.

 
IL-8 stimulates activation of SREBPs in endothelial cells in a receptor-dependent manner in vitro and in vivo
To examine whether these observations are specific for IL-8, we detected the precursor and cleaved forms of SREBP1 after preincubation of the endothelial cells with anti-CXCR1 alone, with anti-CXCR2 alone, or with both antibodies simultaneously, followed by treatment with IL-8. Pretreatment with anti-CXCR1 or anti-CXCR2 alone does not fully inhibit the IL-8-induced SREBP1 activation, although the anti-CXCR2 antibody exerts a more potent inhibitory effect. In contrast, SREBP1 activation was abolished when the cells were preincubated with both antibodies simultaneously. Taken together, these data suggest that the IL-8-mediated SREBP1 activation process is dependent on CXCR1 and CXCR2, although CXCR2 may play a larger role in SREBP1 activation (Fig. 3A ). One possible explanation for the more prominent role of CXCR2 in this process may lie in the fact that these microvascular endothelial cells express much higher levels of CXCR2 than CXCR1. We also preincubated the hMVECs with or without antibodies to IL-8 or its receptors, CXCR1 and CXCR2, or with an inhibitor for both receptors prior to IL-8 exposure. Treatment with both antibodies simultaneously or the inhibitor eliminated IL-8-induced SREBP cleavage (Fig. 3B) , indicating that IL-8-induced SREBP activation is specific to this chemokine and that the signaling pathways from CXCR1 and CXCR2 involve SREBP.


Figure 3
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  		Figure 3. IL-8-induced SREBP activation is abrogated by inhibition of IL-8 or CXCR1/2. (A) hMVECs were preincubated for 1 h with 3 µg/ml CXCR1 antibody, 3 µg/ml CXCR2 antibody, or 3 µg/ml CXCR1 and anti-CXCR2 antibodies, followed by treatment with 50 ng/ml IL-8. Lysates were analyzed by immunoblotting for SREBP1. GAPDH was used to verify equal loading. (B) hMVECs were pretreated for 1 h with 3 µg/ml IL-8 antibody, 3 µg/ml CXCR1, -2 antibodies, or 100 nM DF1681B (CXCR1/2 inhibitor), followed by treatment with 50 ng/ml IL-8. Lysates were analyzed by immunoblotting for SREBP1 or SREBP2. GAPDH was used to verify equal loading. (C) Partial excision wounds were made in the skin of rabbits. Skin samples were removed at indicated time-points after wounding, extracted for immunoblot analysis for IL-8. H2B was used to verify equal loading. (D) Skin samples were removed at Days 0 and 4 post-wounding, fixed, and sectioned for immunofluorescence with anti-IL-8 (shown in green). The arrows indicate positive staining in the epidermis (Day 0) or dermis of the skin (Day 4). Original scale bars = 100 µm. (E) Sections of the tissues were immunolabeled with anti-PECAM to delineate endothelial cells in the microvessels. Vessel number was counted by microscopy (40x objective). Mean ± SD, n = 4. *, P < 0.05. (F) Wounds were treated with PBS or anti-CXCR1/2 in PBS, and samples were collected at Day 4. Tissues were stained with antibodies against PECAM and SREBP1, and secondary antibodies were conjugated to CY3 or FITC, respectively. PECAM is shown in red and SREBP1, in green. Original scale bars = 100 µm.

 
To determine whether our observations in hMVEC cultures correlate with SREBP expression in vivo, we observed microvessels of the skin during wound-healing. We found that IL-8 protein levels increase rapidly and remain elevated up to 5 days after wounding (Fig. 3C) . The increase in IL-8 occurs in the connective tissue and in the ECs, whereas in nonwounded skin, IL-8 is primarily found in the epidermis (Fig. 3D) . Figure 3E shows that the number of microvessels on Day 4 after wounding is increased and that application of antibodies to IL-8 or CXCR1 and -2 during the first 4 days of the healing process greatly decreased the numbers of new blood vessels. In addition, SREBP1 levels increased in the ECs by Day 4 after wounding, and treatment during the healing process with antibodies against CXCR1 and -2 inhibited this increase in SREBP1 (Fig. 3F) . Similar results were observed with antibodies to IL-8 (not shown). These data suggest that IL-8 and its receptors are key factors in SREBP activation during angiogenesis in vivo.

Activation of SREBPs by IL-8 is SCAP-dependent and is critical for angiogenesis
As SCAP is critical for SREBP activation during lipid metabolism, we investigated the involvement of SCAP in IL-8-induced SREBP activation. To inhibit SCAP activity, we transfected hMVECs with SCAPi and found that this approach reduces the SCAP protein level by 50–60% [11 ]. The results showed that the increase of the precursor and the cleaved forms of SREBP1 and SREBP2 stimulated by IL-8 is abrogated by SCAPi, whereas pGL3i has no effect (Fig. 4A and 4B ). IL-8 stimulation of SREBP cleavage is also blocked by treatment with 25-HC, a steroid SREBP inhibitor that prevents the transport of the SCAP-SREBP complex from the ER to the Golgi, thereby blocking SREBP cleavage (activation) [28 , 29 ]. Furthermore, to show that activation of SREBPs is involved in IL-8-induced angiogenesis, we performed in vivo and in vitro experiments with the CAM and tube formation assays, respectively. We showed previously that local application of IL-8 on 10-day-old CAMs stimulates angiogenesis [25 , 30 , 31 ]. This effect of IL-8 was abolished by application of 25-HC with IL-8; counting the number of blood vessel branches in all treatments revealed that 25-HC reduces the number of branches to basal levels (Fig. 4C and 4D) . To confirm these results, we extracted protein from treated CAMs, detected the SREBP protein level by immunoblot analysis using antibodies to SREBP1 (chicken antibodies to SREBP2 are not available), and showed that 25-HC decreased the levels of the precursor form of SREBP and completely inhibited the cleaved form of this transcription factor (Fig. 4E) . We do not present SCAPi transfection assays in the CAM, as neither SREBPs nor SCAP have been isolated in chickens, and the human probes do not function in the CAM (data not shown). In the tube formation assay, we showed that SCAPi and 25-HC markedly inhibited IL-8-induced tubulogenesis. To further prove the critical functions of SREBPs in angiogenesis, we infected hMVECs with a recombinant Ad, which encodes for the HA-SREBP2(N), active form, or as control, an Ad-ß-gal and used these cells for the tube formation assay (Fig. 4F) . These data show that sustaining expression of the SREBP2(N) induced tube formation, whereas Ad-ß-gal did not affect tube formation, underscoring the importance of SREBP in IL-8-induced angiogenesis.


Figure 4
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  		Figure 4. IL-8-induced angiogenesis is decreased by SREBP inhibition in vitro and in vivo. (A, B) hMVECs were transfected with siRNA (150 nmol/L) specific for SCAP (SCAPi) or for GL3 (GL3i; control) for 24 h or were pretreated with 25-HC (5 µg/µL) or DMSO (vehicle for 25-HC) for 24 h. After transfection or pretreatment, cells were treated with IL-8 (50 ng/ml) and analyzed by immunoblotting for SREBP1 and SREBP2. (C) Ten-day-old CAMs were treated with DMSO, DMSO + IL-8, or 25-HC + IL-8, and samples were collected at 14 days. Micrographs show the pellets and the surrounding areas. (D) Numbers of vessel branches from 12 pellets with each treatment were counted and are presented as mean ± SD. *, P < 0.05; **, P < 0.001. (E) The CAM areas covered by pellet were harvested and pooled for protein extraction (n=5), followed by immunoblot analysis using a SREBP-1 antibody. GAPDH was used to verify equal loading. (F) hMVECs were transfected with SCAPi or GL3i for 24 h or preincubated with 25-HC (5 µg/µL) or DMSO. These cells were then replated on Matrigel-coated 24-well plates and treated with or without IL-8 (50 ng/ml) for another 24 h. To investigate the effect of sustaining SREBP expression on tube formation, hMVECs were infected with Ad-HA-SREBP2(N) or Ad-ß-gal (control) for 24 h. These cells were used to perform a tube formation assay. Representative images come from three independent experiments.

 
SREBPs mediate IL-8-induced permeability, proliferation, and migration of hMVEC
Increased permeability of the endothelium, EC migration, and proliferation is a key process, which occurs during the early stages of angiogenesis, and IL-8 is actively involved in this process [7 , 32 ]. Figure 5A shows that IL-8 stimulates permeability of a hMVEC tight monolayer under conditions that mimic endothelium in vivo and that the IL-8-induced permeability was suppressed when SREBP activation was inhibited by preincubation of the cells with SCAPi or 25-HC. We also visualized the IL-8-induced permeability by examining the formation of intercellular gaps in the hMVEC monolayer by immunolabeling with an antibody to PECAM, a marker for ECs (Fig. 5B) . hMVEC transfected with SCAPi or treated with 25-HC for 24 h prior to IL-8 treatment showed a significantly lower number of gaps compared with that observed under control conditions in which cells were transfected with pGL3i or pretreated with DMSO.


Figure 5
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  		Figure 5. Inhibition of SREBP activation suppresses IL-8-induced permeability and proliferation in hMVEC, which (A) were transfected with SCAPi or GL3i (150 nmol/L) or pretreated with 25-HC (5 µg/µL) or DMSO for 24 h and plated on the membrane (3 µm) of a transwell insert to form a tight endothelial monolayer. Assays for permeability are described in Materials and Methods. (B) hMVECs were plated as above, treated with IL-8 for 30 min, and immunolabeled using an antibody to PECAM to show the formation of intercellular gaps. Arrows point to gaps. (C) hMVECs were transfected or pretreated as in A and then were incubated with IL-8 (50 ng/mL) and BrdU (10 µmol/L) for 12 h before fixation. BrdU incorporation was measured using a cell proliferation assay (Oncogene, San Diego, CA). Mean ± SD from three separate experiments. *, P < 0.05; **, P < 0.001. Original scale bar is 100 µm. OD, Optical density.

 
In proliferation assays, inhibition of SREBP by SCAPi or 25-HC significantly reduces DNA synthesis in hMVEC in response to IL-8 treatment (Fig. 5C) . Likewise, using a variety of approaches, we have shown that IL-8 promotes hMVEC migration. Using the cloning ring migration assay [26 ], we found that inhibition of SREBPs by SCAPi or 25-HC significantly suppressed IL-8-induced cell migration (Fig. 6A and 6B ). Similar results were obtained by using a modified Boyden-Chamber assay [27 ] to test the involvement of SREBPs on IL-8-induced chemotaxis; SCAPi and 25-HC inhibited hMVEC migration through the filter pores up the IL-8 concentration gradient (Fig. 6C) .


Figure 6
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  		Figure 6. IL-8-induced SREBP activation in hMVEC migration and chemotaxis. (A) hMVECs were transfected with SCAPi or GL3i or were treated with 25-HC or DMSO. After 24 h, the hMVECs were used in the cloning ring or the chemotaxis assays. Data are shown as mean ± SD from four separate experiments. (B) Representative images of the extent of cell migration at 48 h after treatment. The heavy line indicates the original edge of the cells after removal of the cloning ring. Original scale bar = 150 µm. (C) hMVECs were transfected or pretreated as above. After 24 h, they were replated on the underside of collagen-coated membrane transwell inserts (8 µm pores). After 3 h of treatment with IL-8, the cells on the upper side of the membrane were counted and the numbers averaged. The experiments were performed in triplicate. *, P < 0.05; **, P < 0.001.

 
SREBP regulation of Rho activation is isoprenoid-mediated and is required for IL-8-induced angiogenesis
To gain insight into the molecular mechanisms by which SREBPs function and contribute to an increase of permeability, cell migration, and proliferation, we analyzed the importance of SREBPs in RhoA activation using a Rho-GTP pull-down assay. This small guanosinetriphosphatase (GTPase) is responsible for cytoskeletal reorganizations critical for accomplishment of these three processes important in angiogenesis. Two peaks of RhoA activation were observed following IL-8 treatment of hMVEC (Fig. 7A ). The late IL-8-induced RhoA activation was inhibited by SCAPi and 25-HC, whereas the early Rho activation, albeit decreased, was not inhibited. These data strongly suggest that SREBP is necessary for the late RhoA activation following IL-8 treatment (Fig. 7B) . To verify that IL-8 and VEGF share this segment of the signal-transduction pathway, we repeated the RhoA pull-down assay using VEGF as the stimulator (Fig. 7C) . Like IL-8, VEGF treatment led to a bimodal activation of RhoA, and the late activation of this GTPase was eliminated by inhibition of the SREBPs.


Figure 7
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  		Figure 7. SREBP regulates RhoA activity in response to IL-8 or VEGF treatment. (A) hMVECs were treated with IL-8 (50 ng/ml) as indicated. Cell lysates were mixed with 30 µg Rhotekin beads for 1 h to pull down active RhoA, and the precipitate was examined by immunoblot analysis for RhoA. Total cell lysate was examined for total RhoA. (B) Cells were transfected with SCAPi or were pretreated with 25-HC for 24 h to determine the effect of SREBP on RhoA activation after IL-8 treatment. (C) Similar RhoA pull-down assays performed after VEGF treatment. GAPDH was used to verify equal loading.

 
Increasing evidence suggests that SREBPs are the primary regulators of HMGCoAR activity [33 ], which regulates the biosynthesis of FPP and GGPP, an isoprenoid-mediated fine-tuning of RhoA functional regulation. To determine whether the signaling pathway from SREBPs to RhoA in angiogenesis involves HMGCoAR, we incubated cells with 25-HC + FPP + GGPP for 24 h, followed by IL-8 treatment (Fig. 8A ). This treatment markedly rescued the inhibition of RhoA activity by 25-HC. Futhermore, treatment of CAMs with IL-8 along with 25-HC + FPP + GGPP significantly reversed inhibition by 25-HC of IL-8-stimulated microvessels (Fig. 8B) . As the CAM assay is a chicken assay, it has limitations (see above). Therefore, to demonstrate further that regulation of RhoA activation by SREBPs occurs via isoprenylation and is required for IL-8-induced angiogenesis, we performed experiments using the angiogenesis tube formation assay in vitro. Addition of FPP and GGPP to the hMVEC cultures significantly rescued the inhibition by SCAPi or 25-HC of IL-8-induced capillary tube formation (Fig. 8C) . To further investigate Rho involvement in this signaling pathway, we transfected the hMVECs with an expression vector containing the C. botulinum C3 exoenzyme and plated them for tube formation assay. The C3 exoenzyme is a specific inhibitor for Rho GTPases, as C3 catalyzes adenosine 5'-diphosphate-ribosylation of these proteins, which inactivates them [24 , 34 ]. Treatment of cells with IL-8 in the presence of C3 abolished the isoprenoid-restored tube formation, whereas cells transfected with vector alone as a control are rescued by FPP and GGPP, allowing tube formation to occur (Fig. 8C) .


Figure 8
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  		Figure 8. SREBP regulates RhoA activity through isoprenylation in response to IL-8 treatment. (A) Cells were preincubated with 25-HC + FPP (10 µmol/L) + GGPP (10 µmol/L) for 24 h and then treated with IL-8 (50 ng/ml) as indicated for the GST-RhoA pull-down assay. The results shown are from one of two experiments. (B) Images of IL-8-induced angiogenic response in CAMs incubated with pellets containing DMSO (control), IL-8 (3 µg), IL-8 + 25-HC (5 µg), or IL-8 + 25-HC + FPP (10 µmol/L) + GGPP (10 µmol/L). The data are from four independent experiments, each carried out in triplicate. (C) Inhibition of IL-8-induced tube formation was rescued by isoprenylation. hMVECs were transfected with SCAPi or GL3i + FPP (10 µmol/L) + GGPP (10 µmol/L) or preincubated with 25-HC for 24 h. The cells were then replated on Matrigel and treated with or without IL-8 (50 ng/ml) for another 24 h. In addition, hMVECs were transfected with an expression vector containing C. botulinum C3 exoenzyme (specific inhibitor of Rho) or with pEGFP-C1 (vector alone) for 24 h and then placed in the tube formation assay. Representative images come from three independent experiments. (D) RhoA pull-down assay similar to A shows that Mevastatin (10 µmol/L; inhibitor for HMGCoA) abrogates RhoA activation. GAPDH was used to verify equal loading.

 
To confirm that HMGCoAR is involved in IL-8-induced RhoA activation, we used Mevastatin in conjunction with IL-8 treatment (Fig. 8D) . The hMVECs were pretreated with 10 µmol/L Mevastatin for 24 h to inhibit isoprenylation. Mevastatin is a potent, competitive inhibitor of HMGCoAR, thus blocking the geranylgeranylation and farnesylation necessary for membrane-associated activity of many proteins, including RhoA. As shown, this inhibitor completely abrogated RhoA activation by IL-8, strongly suggesting that HMGCoAR is involved in IL-8-induced RhoA activation.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The mechanisms of IL-8-incuded angiogenesis are complex and remain poorly understood. In this study, we investigated in detail the role of SREBPs in IL-8-induced angiogenesis and key underlying signaling mechanisms. Specifically, we show that IL-8 stimulates SREBP1 and SREBP2 gene expression and activation in hMVECs and that these effects are IL-8 receptor-dependent; IL-8 stimulates the expression of SREBP-induced genes such as FAS, LDLR, and HMGCoAR; in vivo, SREBP expression is increased in endothelial cells during IL-8-induced neovascularization; inhibition of SREBP by SCAP siRNA and 25-HC abrogates IL-8-stimulated SREBP cleavage and suppression of IL-8-induced tube formation and angiogenesis (CAM assay) in vivo; SCAPi or 25-HC failed to inhibit tube formation in hMVECs overexpressing the active form of SREBP2; inhibition of SREBP suppresses IL-8-induced permeability, migration, and proliferation of hMVECs; targeting HMGCoAR effects on isoprenylation in IL-8-induced activation of SREBPs regulates RhoA activation; signaling through SREBP-HMGCoAR-RhoA is critical for IL-8-induced angiogenesis.

To examine whether IL-8-induced SREBP activation is a direct effect, we used several different approaches to block CXCR1 and CXCR2, the receptors for this chemokine. As shown in this study, IL-8 stimulated the activation of SREBPs, and specific antibodies or inhibitors for these receptors as well as antibodies to IL-8 resulted in abrogation of SREBP activation. Furthermore, SCAPi and 25-HC significantly decreased EC endothelium permeability, migration, proliferation, and tube formation induced by this chemokine. These data suggest that SREBPs are critical in the processes necessary for IL-8-induced angiogenesis. Our experiments in vivo using rabbit skin wound-healing and the CAM assay allowed us to further test our hypothesis that SREBP activation plays an important role in IL-8-induced angiogenesis. In addition, SREBPs are activated by other angiogenic factors. bFGF is a member of the heparin-binding growth factor family, whose activities are mediated by receptor tyrosine kinases. Thrombin activates members of the G-protein-coupled receptor family of proteolytically activated receptor-1, -3, and -4, through which it stimulates the proliferation and permeability responses in hMVECs associated with angiogenesis [20 , 35 ]. Although the aforementioned factors exert biological effects through different receptors, they share a common signal transduction pathway segment involving SREBP activation, strongly suggesting that SREBPs are key, common molecules in mediating or priming EC functions for angiogenesis.

Given the importance of SREBPs in the regulation of membrane biosynthesis in fine-tuning membrane biophysical properties, it is not surprising that these transcription factors can be responsible for the regulation of EC function. It has been shown that plasma membrane microviscosity, which is a function of membrane lipid composition and redistribution, increases at the leading edge of ECs treated with VEGF, resulting in polarization of the cell and pseudopodia formation. In addition, the generation of a microviscosity gradient can affect the actin dynamics needed for EC migration [36 , 37 ]. The small GTPase RhoA is essential in modulating the formation of stress fibers and focal adhesions and in promoting the reorganization of F-actin and tyrosine phosphorylation of cell adhesion proteins in cell-cell junctions, resulting in de-adhesion, which leads to permeability of the endothelium [38 39 40 ].

Our results also show that IL-8 stimulates a bimodal activation of RhoA and that the late activation is inhibited by SCAPi or 25-HC, two means of inhibiting SREBPs. This indicates that IL-8-induced late activation of RhoA occurs downstream of SREBP activation. RhoA activity is known to be controlled by its isoprenylation state; given the role of SREBP in cellular lipid homeostasis, the lipid-regulating effects of SREBP could be a link between SREBP and RhoA activation. Here, IL-8 induced the expression of genes involved in lipid biosynthesis, such as FAS, LDLR, and HMGCoAR. The latter is the rate-limiting enzyme in cholesterol biosynthesis, during which isoprenoids, including FPP and GGPP, are produced. This HMGCoAR-mediated FPP and GGPP biosynthesis is required for the post-translational lipidation, membrane localization, and function of RhoA [41 42 43 44 45 ]. Our results suggest that the lipid-regulating effects of SREBP are a link between SREBP and RhoA activation. We show that addition of FPP and GGPP to cultured hMVECs or to CAMs treated with both 25-HC in the presence of IL-8 rescues RhoA activation and angiogenesis.

In conclusion, this study furthers our understanding of the mechanisms of IL-8-induced angiogenesis by showing that activation of SREBPs is essential for the angiogenic response stimulated by this chemokine. Furthermore, we also show that this is the case for several other angiogenic factors that belong to different families of proteins and activate different receptors. These studies suggest that activation of SREBPs followed by RhoA activation is important for the angiogenic response stimulated by several different classes of angiogenic factors. Therefore, targeted inhibition of SREBPs and/or RhoA may eliminate the need to inhibit different specific receptors activated by multiple types of angiogenic factors and thus, provide a novel approach to prevention and treatment of diseases involving abnormal angiogenesis.


   ACKNOWLEDGEMENTS
 
This study was supported in part by AHA Grant #0050732Y (M. M-G.) and National Institutes of Health HL 77448 (J. S). We thank Dr. K. DeFea for the GST-TRBD fusion protein, Genentech for the CXCR1 and CXCR2 antibodies, and Dr. Miguel Angle Del Pozo for the pEGFP-C1-C3 vector.

Received March 5, 2006; revised April 24, 2006; accepted April 25, 2006.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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