ARTICLE

Role of Hypoxia-Inducible Factor 1{alpha} in Gastric Cancer Cell Growth, Angiogenesis, and Vessel Maturation

Oliver Stoeltzing, Marya F. McCarty, Jane S. Wey, Fan Fan, Wenbiao Liu, Anna Belcheva, Corazon D. Bucana, Gregg L. Semenza, Lee M. Ellis

Affiliations of authors: Department of Cancer Biology (OS, FF, WL, AB, CDB, LME) and Department of Surgical Oncology (MFM, JSW, LME), The University of Texas M. D. Anderson Cancer Center, Houston; McKusick-Nathans Institute of Genetic Medicine, Johns Hopkins University School of Medicine, Baltimore, MD (GLS)

Correspondence to: Lee M. Ellis, MD, Department of Surgical Oncology, Box 444, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030-4009 (e-mail: lellis{at}mdanderson.org)


    ABSTRACT
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Background: Hypoxia-inducible factor 1 (HIF-1), a heterodimer comprising the oxygen-regulated subunit, HIF-1{alpha}, and HIF-1{beta}, mediates transcription of the gene for vascular endothelial growth factor (VEGF). Overexpression of HIF-1{alpha} is associated with tumor angiogenesis and tumor cell proliferation and invasion. We examined the effects of inhibiting HIF-1{alpha} activity on angiogenesis and human gastric cancer growth in vivo. Methods: Human gastric cancer TMK-1 cells were stably transfected with pHIF-1{alpha}DN, an expression plasmid encoding a dominant-negative form of HIF-1{alpha} that dimerizes with endogenous HIF-1{beta} to produce HIF-1 complexes that cannot activate transcription, or with the empty expression vector (pCEP4). Two clones of pHIF-1{alpha}DN–transfected cells, DN2 and DN3, were tested in all experiments. We used an enzyme-linked immunosorbent assay to measure VEGF secretion by transfected cells cultured in hypoxic (1% O2) or nonhypoxic (20% O2) conditions. We used subcutaneous and orthotopic mouse tumor models to examine the growth of tumors derived from injected pHIF-1{alpha}DN–or pCEP4-transfected cells. Tumor cell proliferation, vessel area (a measure of functional vascular volume), and tumor endothelial cell association with pericyte-like cells (a measure of vessel maturation) were analyzed by immunohistochemical or immunofluorescent staining. All statistical tests were two-sided. Results: DN2 cells and DN3 cells secreted less VEGF than pCEP4-transfected TMK-1 cells when cultured in nonhypoxic or hypoxic conditions (e.g., DN2 versus pCEP4 in nonhypoxic conditions: 645 pg of VEGF/106 cells versus 1591 pg of VEGF/106 cells, difference = 946 pg of VEGF/106 cells [95% confidence interval {CI} = 640 to 1251 pg of VEGF/106 cells; P = .006]; DN2 versus pCEP4 in hypoxic conditions: 785 pg of VEGF/106 cells versus 2807 pg of VEGF/106 cells, difference = 2022 pg of VEGF/106 cells [95% CI = 1871 to 2152 pg of VEGF/106 cells; P<.001]). In the subcutaneous tumor model, tumors derived from DN2 or DN3 cells had lower final volumes, weights, and vessel areas, less tumor endothelial cell association with desmin-positive cells, and fewer proliferating tumor cells than tumors derived from pCEP4-transfected cells. In the orthotopic tumor model, tumors derived from DN2 cells had smaller volumes and less vessel area and maturation than tumors derived from pCEP4-transfected cells. Conclusions: Inhibition of HIF-1{alpha} activity impairs gastric tumor growth, angiogenesis, and vessel maturation.



    INTRODUCTION
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Angiogenesis is essential for tumor growth and metastasis [reviewed in (1)]. Tumor neovascularization depends on the production of specific angiogenic factors, either by host or tumor cells, that shift the angiogenic balance toward a proangiogenic phenotype (2,3). Vascular endothelial growth factor (VEGF) is one of the major factors that contribute to angiogenesis and metastasis in numerous tumor types (4,5), and VEGF overexpression has been associated with tumor progression and poor clinical outcome (612) [reviewed in (1,13)].

The transcription factor hypoxia-inducible factor 1 (HIF-1), a heterodimer consisting of an HIF-1{alpha} and a HIF-1{beta} subunit, is an important upstream mediator of VEGF expression in cancer cells (1417). Like VEGF, the HIF-1{alpha} subunit is overexpressed in a variety of human cancers (1823). HIF-1{alpha} functions as a physiologic mediator of cellular responses to hypoxia in both normal and malignant tissues (24,25). Under nonhypoxic conditions, HIF-1{alpha} is rapidly ubiquitinated and degraded by proteosomes. In in vitro assays with human HeLa cells (26), it has been shown that levels of HIF-1{alpha} protein increased exponentially as cells were subjected to decreasing O2 concentrations (i.e., from 20% O2 to 0% O2), with a half-maximal response between 1.5% and 2% O2 and a maximal response at 0.5% O2. Under such hypoxic conditions, which are common also in human tumors (27), HIF-1{alpha} heterodimerizes with the constitutively expressed HIF-1{beta} subunit, and together they bind to DNA and increase the transcription of target genes, which include VEGF, erythropoietin, transferrin, endothelin 1, inducible nitric oxide synthase, and insulin-like growth factor II (2831). Some hypoxia-independent mechanisms of HIF-1 activation in tumor cells have also been reported, such as genetic alterations in tumor suppressor genes (i.e., p53, VHL, and PTEN) and oncogenes (i.e., SRC, HER2neu, and H-RAS) (3235). Activation of certain growth factor receptors (e.g., insulin-like growth factor I receptor) has also been shown to increase expression of HIF-1{alpha} and, subsequently, VEGF in human colon carcinoma cells (36). Results of recent studies demonstrated that HIF-1{alpha} may also regulate the invasiveness of colon cancer cells by altering the expression of genes encoding intermediate filaments (i.e., vimentin, keratins), extracellular matrix components (i.e., fibronectin), and proteases (i.e., matrix metalloproteinase 2 and the urokinase plasminogen activator receptor) (37).

Several different approaches have been used to assess the effects of inhibiting HIF-1{alpha} function on in vivo tumor growth and angiogenesis [reviewed in (17)]. To date, all studies have used various mouse xenograft models with subcutaneously implanted tumors derived from mouse embryonic stem (ES) cells (38,39) or mouse embryonic fibroblasts (40) that were genetically engineered to lack HIF-1{alpha}; human colon cancer HCT116 cells stably transfected with a construct that expresses mutated p300 protein (wild-type p300 physiologically heterodimerizes with the HIF-1{alpha}/{beta}complex under hypoxic conditions) (41); or human pancreatic cancer PCI-43 cells stably transfected with a construct that expresses a dominant-negative mutant HIF-1{alpha} (42). However, these experimental models have led to seemingly contradictory findings with respect to the effects of HIF-1 inhibition on tumor growth, VEGF expression in vivo, and angiogenesis. For example, one study (40) reported that HIF-1 inhibition was not associated with reduced VEGF expression in vivo or vessel density in tumors despite reducing subcutaneous growth of ES cells, whereas another study (38) reported an accelerated subcutaneous growth of HIF-1{alpha}–/– ES cells. Furthermore, another study demonstrated that decreased HIF-1{alpha} activity (HIF{alpha}–/–) led to reduced tumor growth of astrocytoma cells in a subcutaneous tumor model but that the same cells exhibited a much more aggressive phenotype when implanted at an orthotopic site (43). Thus, inherent differences among tumor models (e.g., in cell types, sites of tumor growth, and approaches used to inhibit HIF-1{alpha} activity) have made the data on the effects of HIF-1{alpha} inhibition difficult to interpret.

Because HIF-1{alpha} is an important regulator of solid tumor growth, we hypothesized that HIF-1{alpha} is an important mediator of gastric tumor growth in vivo and that inhibition of its function would substantially impair tumor growth and angiogenesis. To investigate this hypothesis, we stably transfected human gastric cancer cells with a construct expressing a dominant-negative mutant version of HIF-1{alpha} (pHIF-1{alpha}DN) that competes with endogenous HIF-1{alpha} for dimerization with HIF-1{beta} but forms HIF-1 complexes that can neither bind to DNA nor activate transcription (15,44). We analyzed the effect of disrupting HIF-1 with this mutant on the biology of human gastric cancer cells in vitro and i vivo.


    MATERIALS AND METHODS
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Cell Culture

Human gastric cancer TMK-1 cells (obtained from Eiichi Tahara, University of Hiroshima, Hiroshima, Japan) were maintained in Dulbecco's modified Eagle's Medium (DMEM) supplemented with 10% fetal bovine serum (10% FBS–DMEM), 2 U/mL of penicillin–streptomycin mixture (Flow Laboratories, Rockville, MD), 1% vitamin solution (Life Technologies, Grand Island, NY), 1 mM sodium pyruvate, 2 mM L-glutamine, and nonessential amino acids (Life Technologies) and incubated in 5% CO2–95% air at 37 °C.

Stable Transfections

We used a previously described (44) expression construct encoding a dominant-negative mutant HIF-1{alpha} (pHIF-1{alpha}DN). The vector for pHIF-1{alpha}DN, pCEP4, contains a hygromycin resistance gene (Invitrogen, Carlsbad, CA). The protein expressed by pHIF-1{alpha}DN consists of residues 1–3 of HIF-1{alpha} fused in-frame to residues 28–390 of HIF-1{alpha} and lacks the DNA binding and transactivation domains of the 826-amino-acid wild-type protein. Cells transfected with the empty pCEP4 expression vector served as controls for all experiments. After transfection, cells were grown in selective medium (10% FBS–DMEM containing 200 µg of hygromycin per milliliter), and several clones of each were randomly selected and subsequently expanded in selective medium. Early passages of transfected TMK-1 cells (clones) were frozen and stored in liquid nitrogen before being used for in vitro and in vivo experiments. Screening for successful transfection was done by western blotting analysis for nuclear HIF-1{alpha} protein levels after hypoxic stimulation of cells, as described below. Cells were kept in culture for a maximum of 4–5 additional passages in selective medium. For in vivo experiments, clones of stably transfected cells were harvested from subconfluent cultures by trypsinization (0.25% trypsin and 0.02% EDTA) for 3 minutes, washed in 10% FBS–DMEM, and counted. Cell viability was assessed with the trypan blue exclusion assay.

Western Blot Analysis of HIF-1{alpha} Expression in Nuclear Extracts

Western blot analysis of HIF-1{alpha} expression in nuclear extracts was performed as described elsewhere (45). In brief, clones of pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells (at 50%–60% confluence) were incubated in 1% FBS–minimal essential medium (MEM) to reduce the effects of serum on intracellular signaling pathways for 16 hours under hypoxic (1% O2, 5% CO2, 94% nitrogen) or nonhypoxic (20% O2) conditions in a NAPCO incubator (Precision Scientific, Winchester, VA), washed twice with ice-cold phosphate-buffered saline (PBS), harvested by trypsinization, and resuspended in ice-cold PBS (1 mL). After centrifugation at 400g for 5 minutes at 4 °C, cell pellets were resuspended in 200 µL of Buffer A (10 mM HEPES, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM dithiothreitol [DTT], 0.2 mM phenylmethylsulfonyl fluoride [PMSF], and 1x protein inhibitor tablets [Complete Mini; Roche Diagnostics, Indianapolis, IN]), incubated for 30 minutes on ice, and centrifuged again (at 7150g for 3 minutes at 4 °C). The resulting cell pellets were washed twice with ice-cold Buffer A. To extract nuclear proteins, we resuspended the washed pellets in 50 µL of ice-cold Buffer C (20 mM HEPES, 25% glycerol, 450 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM DTT, 0.5 mM PMSF, and 1x protein inhibitor), incubated for 30 minutes on ice, and centrifuged (22 000g for 15 minutes at 4 °C). We determined the protein concentrations of the resulting supernatants with a commercially available kit (Bio-Rad) and spectrometry and resolved 70-µg aliquots of each supernatant by electrophoresis on a sodium dodecyl sulfate–6% polyacrylamide gel. The resolved proteins were transferred to membranes and incubated overnight at 4 °C in 3% milk, 0.5 % Tris-buffered saline, and 0.1% Tween 20 containing a mouse monoclonal anti–HIF-1{alpha} antibody (1 : 5000 dilution; clone H1{alpha}67 from BD Biosciences Pharmingen, San Diego, CA) or a mouse monoclonal anti–HIF-1{beta} antibody (1 : 1000 dilution in 5% milk; Calbiochem, San Diego, CA). The membranes were then washed in PBS for 30 minutes and incubated with an anti–mouse secondary antibody (1 : 5000 dilution; Amersham Biosciences) for enhanced chemiluminescent detection of monoclonal antibody binding (ECL; Amersham Biosciences, Piscataway, NJ). Densitometric analysis was performed with NIH Image Analysis software (version 1.62) to quantify the results of western blot analyses. The anti–HIF-1{alpha} monoclonal antibody does not recognize the HIF-1{alpha}DN protein. Two clones of pHIF-1{alpha}–transfected TMK-1 cells that showed marked reduction in levels of nuclear HIF-1{alpha} protein after hypoxic induction were selected for the experiments. Pooled pCEP4-transfected cells served as additional control in subsequent studies.

Cell Proliferation and Cell Cycle Analysis of Transfected Cells

pHIF-1{alpha}DN–or pCEP4-transfected cells in 10% FBS–DMEM were plated at 1000 cells/well in 96-well plates and incubated for 24 or 48 hours at 37 °C. We used the methylthiazole tetrazolium (MTT) assay to quantify cell proliferation as previously described (46). Fluorescence-activated cell sorting (FACS) analysis was performed to detect potential changes in the cell division cycle of the transfected cell lines. For the FACS analysis, cells were grown to approximately 50% confluence, washed in PBS, and fixed in 70% ethanol overnight at 4 °C. Propidium iodine (10 µg/mL) supplemented with RNase A (200 µg/mL) was added to the cells for 30 minutes (at 37 °C) prior to FACS analysis.

Enzyme-Linked Immunosorbent Assays

VEGF secretion from transfected gastric cancer cells. pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells were plated at similar densities (30%–40% confluence) and incubated for 16 hours in 5% FBS–MEM under hypoxic or nonhypoxic conditions. We used an enzyme-linked immunosorbent assay (ELISA) kit specific for the 165-amino-acid form of human VEGF (Biosource International, Camarillo, CA) according to the manufacturer's protocol to measure VEGF protein concentrations in 3 mL of conditioned medium collected from the cells. The cells were rinsed with cold PBS, harvested by trypsinization, and counted. VEGF levels were expressed as picograms of VEGF/106 cells.

Assessment of genes known to regulate pericyte function. We used ELISA kits for human platelet-derived growth factor (PDGF)–BB (homodimer) or human angiopoietin 2 (Ang-2; both kits from R&D Systems, Minneapolis, MN, were used according to the manufacturer's protocols) to measure expression of these proteins in conditioned medium collected from pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells. The conditioned media were collected after the cells were incubated for 16 hours in 5% FBS–DMEM under nonhypoxic or hypoxic conditions as described above. Cells were harvested by trypsinization and counted. The conditioned media were centrifuged for 5 minutes at 350g, passed through 0.22-µm-pore-size filters (Corning, Corning, NY), and subjected to ELISA without further concentration or after fivefold concentration by ultracentrifugation at 2500g for 30 minutes at 4 °C in Centriprep YM-10 filter units (Millipore, Burlington, MA).

Gene Array Analysis for Expression of Angiogenic Molecules

To investigate the effects of inhibiting HIF-12 function on the expression of other potential mediators of angiogenesis, we performed gene array analysis with commercially available gene array kits according to the manufacturer's protocol (Angiogenesis Array, Q-Series; SuperArrays, Frederick, MD). Briefly, transfected TMK-1 cell clones were incubated for 8 hours in 5% FBS–DMEM under nonhypoxic (20% O2) or hypoxic (1% O2) conditions, and total RNA was extracted from the cells by using Trizol, as described above. RNA samples (2-µg aliquots) were labeled and used to hybridize to the angiogenesis gene array membrane. After overnight hybridization at 60 °C (with constant rotation), the membranes were washed as recommended by the manufacturer and autoradiography was performed.

Animal Models

Eight-week-old male athymic nude mice were obtained from the Animal Production Area of the National Cancer Institute and Development Center (Frederick, MD) and acclimated for 1–2 weeks while caged in groups of five. Mice were housed under pathogen-free conditions and fed a diet of animal chow and water ad libitum throughout the experiment. All experiments were approved by the Institutional Animal Care and Use Committee of The University of Texas M. D. Anderson Cancer Center.

Subcutaneous xenograft model. Mice randomly assigned to one of three groups (10 mice per group) were injected subcutaneously with HIF-1{alpha}DN–transfected TMK-1 cells (clone DN2 or clone DN3) or pCEP4-transfected TMK-1 cells (clone 2) (106 cells in 100 µL of Hanks’ balanced salt solution; cell viability >90%). Tumors were measured every other day, starting at day 4 after injection, when they became palpable and visible. Tumor volumes (in cubic millimeters) were calculated according to the equation width2 x length x 0.5. When any mouse had a tumor diameter that exceeded 1.5 cm, all mice in all groups were given bromodeoxyuridine (BrdU; 1 mg/mouse) by intraperitoneal injection and then 4 hours later were anesthetized by intraperitoneal injection of pentobarbital (50 mg/kg of body weight) and killed by cervical dislocation. Subcutaneous tumors were excised and weighed, and a portion of each was placed in 10% formalin for paraffin embedding or snap-frozen in Optimum Cutting Temperature (OCT) solution (Miles, Elkhart, IN) in preparation for subsequent immunohistochemical analyses.

Orthotopic gastric cancer model. Mice were randomly assigned to one of two groups (10–12 mice per group), anesthetized with methoxyflurane via inhalation while being observed for sufficient cardiopulmonary function (Metofane; Medical Developments, Australia Pty. Ltd.), and subjected to a mini-laparotomy during which pHIF-1{alpha}DN–transfected TMK-1 cells (clone DN2 or DN3) or pCEP4-transfected TMK-1 cells (2 x 106 cells in 50 µL of Hanks’ balanced salt solution; cell viability >90%) were injected in the gastric wall. Mice were observed daily, and all mice were killed when more than three mice in any group exhibited lethargy or reduced mobility. We dissected the mice, excised tumors that had developed, calculated tumor volumes, and harvested tumor tissue as described above. For all in vivo experiments, tumors were harvested by two independent investigators who had no knowledge of the treatment group assignments.

Immunohistochemical Analyses

For all immunohistochemical analyses, we stained multiple sections obtained from complete cross-sections of the tumors. Tumors that had been frozen in OCT were sectioned in 8-µm slices, mounted on positively charged slides, and air-dried for 30 min. For all immunohistochemical and immunofluorescent analyses, controls were done by omitting the primary antibody.

Tumor vessels. We used a rat anti–mouse CD31/platelet endothelial cell adhesion molecule 1 (PECAM-1) antibody (BD Biosciences Pharmingen) and horseradish peroxidase–conjugated goat anti–rat immunoglobulin G (IgG) (Jackson ImmunoResearch Laboratories, West Grove, PA) to stain tumor sections for CD31 as previously described (47). Briefly, sections of OCT-embedded tumors were fixed sequentially in cold 100% acetone, acetone : chloroform (1 volume : 1 volume), and 100% acetone and then washed with PBS. Endogenous peroxidase activity was blocked by incubating the sections in 3% H2O2 in methanol. Sections on slides were incubated in protein blocking solution (1% normal goat serum, 5% normal horse serum) for 20 minutes to block nonspecific antibody binding. The primary antibody was then applied to the sections (at 1 : 800 dilution in protein blocking solution), and the slides were incubated overnight at 4 °C. The secondary antibody (diluted 1 : 200 in protein blocking solution) was subsequently applied to the sections, and the slides were incubated for 1 hour, after which they were washed in PBS and incubated with stable diaminobenzidine substrate (Research Genetics, Huntsville, AL). Substrate development was monitored with bright-field microscopy; the reactions were stopped by washing the slides in distilled water. Sections were counterstained with Gill's No. 3 hematoxylin (Sigma) and mounted to coverslips with Universal Mount (Research Genetics). We counted the number of CD31-stained vessels (at x50 magnification) in sections obtained from four different quadrants of each tumor (all tumor sections were taken 2 mm from the tumor–normal tissue interface) and calculated the mean number of vessels. Vessel counts were performed by an investigator who did not participate in the immunohistochemical staining procedure and who had no knowledge of the treatment group assignments. The cross-sectional area of CD31-positive structures (i.e., vessel area) was also determined in the same four quadrants by capturing images, converting them to grayscale, and analyzing the CD31-stained areas with NIH Image Analysis software (version 1.62; National Institutes of Health, Bethesda, MD) by setting a consistent threshold for all slides so that only CD31-stained cells were apparent. The CD31-positive area was then expressed as pixels-squared per high-power field [HPF]. The CD31-positive area was measured for each tumor in all groups of mice.

Tumor cell apoptosis. We used a mouse anti-BrdU antibody (at 1 : 100 dilution; Beckton Dickinson, San Diego, CA) and horseradish peroxidase–conjugated anti–mouse IgG2 (1 : 200 dilution; BD Biosciences Pharmingen) to stain sections of frozen tumor tissue for nuclear BrdU uptake. Slides were fixed as described above. Binding of the primary antibody was visualized by incubating the slides with stable diaminobenzidine for 10–20 minutes. The sections were rinsed with distilled water, counterstained with Gill's No. 3 hematoxylin for 1 minute, and mounted to coverslips with Universal Mount. Some slides were counterstained with hematoxylin–eosin to allow us to study overall tissue structure. We calculated the average number of BrdU-positive tumor cells counted in four fields representing the proliferative front (i.e., at the interface between tumor tissue and nontumor tissue) at x100 magnification.

Tumor cell proliferation. For immunofluorescent terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL) staining, frozen tissue was fixed in cold acetone and chloroform as described above. The TUNEL assay was performed with a commercial kit (Promega, Madison, WI) according to the manufacturer's protocol. Tissue sections were also counterstained with Hoechst dye (1 : 2000) to allow identification of tumor cells. Immunofluorescence microscopy was done on an epifluorescence microscope equipped with narrow-bandpass excitation filters (Chroma Technology, Brattleboro, VT). Images were captured with a C5810 Hamamatsu camera (Hamamatsu Photonics K.K., Hamamatsu City, Japan) mounted on a Zeiss Axioplan microscope (Carl Zeiss) with Optimas image analysis software (Media Cybernetics, Silver Spring, MD).

Immunofluorescence Analysis of Tumor Vessel Maturation

Pericytes at different stages of differentiation express distinct markers (48). Therefore, we analyzed tumor vessel maturation by staining tumor sections for two different pericyte-associated antigens: {alpha}–smooth muscle actin ({alpha}SMA) and desmin. Because there is no universal definition for pericytes (they are most commonly viewed as the single vascular smooth muscle cells adjacent to the endothelial cells of the microvasculature), we hereafter refer to cells that are associated with endothelial cells and stain positive for pericyte or vascular smooth muscle cell markers as "pericyte-like cells." We examined vessel morphology and the association of tumor endothelial cells with pericyte-like cells in tumors derived from pCEP4- and pHIF-1{alpha}DN–transfected TMK-1 cells by costaining sections with antibodies to CD31 and to {alpha}SMA or desmin (both from DAKO) (48) as previously described (46). Briefly, sections of frozen tumors were stained overnight (4 °C) for CD31/PECAM-1 (BD Biosciences PharMingen) after acetone fixation as described above and visualized by using a Texas Red–conjugated goat anti–rat secondary antibody (1 : 200) (Jackson ImmunoResearch Laboratories). Slides were washed with PBS, and background staining was blocked by incubating the slides with nonspecific goat anti–mouse IgG Fab fragment (diluted 1 : 10 in protein blocking solution; Jackson ImmunoResearch Laboratories) as described above for 1 hour at room temperature. For evaluation of vessel maturation, pericyte-like cells were defined as cells positive for {alpha}SMA or desmin that were in direct contact with tumor endothelial cells. Tumor sections were then incubated overnight at 4 °C with mouse anti–{alpha}SMA antibody (DAKO) (1 : 2000 in protein-blocking solution). The slides were washed, and Alexa 488 (green) (Jackson ImmunoResearch Laboratories) goat anti–mouse secondary antibody (1 : 200 in protein block solution) was added for 1 hour at room temperature. Slides were rinsed in PBS, and nuclei were counterstained with Hoechst dye (1 : 2000) for 2 minutes. Slides were analyzed with an epifluorescence microscope equipped with narrow-bandpass excitation filters (Chroma Technology) to individually select for green, red, and blue fluorescence. Images were captured with a C5810 Hamamatsu camera (Hamamatsu Photonics K.K., Bridgewater, NJ) mounted on a Zeiss Axioplan microscope (Carl Zeiss, Oberkochen, Germany) and using Optimas image analysis software (Media Cybernetics). Images were further processed with Adobe Photoshop software (Adobe Systems, Mountain View, CA). Double-stained slides were analyzed at x100 magnification for pericyte/CD31 colocalization as described elsewhere (46). The percentage of tumor endothelial cells associated with pericyte-like cells was determined by counting CD31-positive vessels in direct contact with {alpha}SMA-positive or desmin-positive cells (coverage) and the number of CD31-positive vessels lacking adjacent staining for {alpha}SMA or desmin (lack of coverage) as described elsewhere (49). The protocol used for desmin staining (1 : 200 dilution of primary antibody in protein blocking solution) was similar to the {alpha}SMA staining protocol except that slides were not blocked with goat anti–mouse IgG Fab prior to the addition of primary antibody. All immunofluorescence staining was analyzed by an examiner blinded to the experimental groups who had not taken part in the immunohistochemical staining.

Statistical Analyses

All statistical analyses were performed using InStat Statistical Software (version 2.03; GraphPad Software, San Diego, CA); P values less than .05 were considered statistically significant. We used Grubb's test (http://www.graphpad.com/quickcalcs/Grubbs1.cfm [last accessed: May 11, 2004]) for assessing the statistical significance of outlying results from the in vivo experiments. Tumor-associated variables were tested for statistical significance by using a two-tailed Student's t test or the Mann-Whitney U test (for nonparametric data).


    RESULTS
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Nuclear HIF-1{alpha} Protein Levels and VEGF Expression in Transfected Gastric Cancer Cells

We examined the effect of hypoxia on nuclear HIF-1{alpha} protein and VEGF secretion in human gastric cancer TMK-1 cells stably transfected with pHIF-1{alpha}DN, which encodes a dominant-negative form of HIF-1{alpha} (DN HIF-1{alpha}), or empty pCEP4 (15,44). After exposure to hypoxic conditions for 16 hours, two clones of pHIF-1{alpha}DN–transfected cells (clones DN2 and DN3) exhibited reduced (i.e., by 70%–80%) nuclear levels of endogenous wild-type HIF-1{alpha} compared with levels in pCEP4-transfected cells, as determined by densitometry (Fig. 1, A). By contrast, HIF-1{beta} protein levels in did not vary substantially among the three cell lines after exposure to hypoxia (data not shown).



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Fig. 1. Effects of HIF-1{alpha} inhibition on nuclear HIF-1{alpha} protein levels and VEGF expression. A) Human gastric cancer TMK-1 cells stably transfected with pHIF-1{alpha}DN (clone DN2 or DN3) or empty vector (pCEP4) were incubated for 16 hours in 1% FBS-MEM under nonhypoxic (20% O2) or hypoxic (1% O2) conditions. The cells were harvested, nuclear proteins were extracted, and endogenous nuclear HIF-1{alpha} was examined by western blot analysis (equal amounts of protein were loaded in each lane). Differences in hypoxia-induced nuclear HIF-1{alpha} protein levels were quantified by densitometry (100% reference induction of control pCEP4 cells). B) VEGF secretion by HIF-1{alpha}DN–or pCEP4-transfected cells incubated under nonhypoxic or hypoxic conditions. Conditioned medium was collected and used for VEGF enzyme-linked immunosorbent assay; VEGF levels were normalized for cell number. *, P = .006 for DN2 (20% O2) versus pCEP4 (20% O2); +, P = .005 for DN3 (20% O2) versus pCEP4 (20% O2). +, P<.001 for DN2 (1% O2) versus pCEP4 (1% O2); #, P<.001 for DN3 (1% O2) versus pCEP4 (1% O2). Error bars indicate 95% confidence intervals.

 
We next examined the effects of hypoxia on HIF-1–regulated target gene expression by measuring the amount of VEGF secreted by pCEP4- and pHIF-1{alpha}DN–transfected TMK-1 cells incubated in nonhypoxic and hypoxic conditions. Under nonhypoxic conditions, the mean VEGF protein levels secreted by pCEP4-transfected TMK-1 cells, DN2 cells, and DN3 cells were 1591, 645, and 477 pg/106 cells, respectively. Compared with mean VEGF protein levels secreted by pCEP4-transfected TMK-1 cells, nonhypoxic mean VEGF protein levels were statistically significantly lower in DN2 cells (difference = 946 pg/106 cells, 95% confidence interval [CI] = 640 to 1251 pg/106 cells; P = .006) and DN3 cells (difference = 1114 pg/106 cells, 95% CI = 640 to 1251 pg/106 cells; P = .005) when cells were incubated at 20% O2 in 5% FBS–MEM for 16 hours. Under hypoxic conditions, mean VEGF protein levels secreted by pCEP4-transfected TMK-1 cells, DN2 cells, and DN3 cells were 2807, 785, and 586 pg/106 cells, respectively. Compared with pCEP4-transfected TMK-1 cells, mean VEGF protein levels were statistically significantly lower in DN2 cells (difference = 2022 pg/106 cells, 95% CI = 1891 to 2152 pg/106 cells; P<.001) and DN3 cells (difference = 2221 pg/106 cells, 95% CI = 2109 to 2332 pg/106 cells; P<.001) when cells were incubated at 1% O2 in 5% FBS–MEM for 16 hours (Fig. 1, B).

Effect of Dominant-Negative Mutant HIF-1{alpha} on Xenograft Tumor Growth

We first investigated the effects of impaired HIF-1{alpha} function on tumor growth and angiogenesis by gastric cancer cells in vivo in a xenograft tumor model in which mice were injected subcutaneously with TMK-1 cells stably transfected with pCEP4 or with pHIF-1{alpha}DN (clone DN2 or DN3). Palpable tumors were first detected in all mice by 8 days after cell injections. At 22 days after cell injections, mean tumor volumes among mice injected with DN2 cells (120 mm3) or DN3 cells (112 mm3) were statistically significantly smaller than those among mice injected with pCEP4-transfected cells (885 mm3) (difference between DN2 and pCEP4 = 765 mm3, 95% CI = 304 to 1226 mm3; P = .003; difference DN3 and pCEP4 = 773 mm3, 95% CI = 289 to 1256 mm3; P = .004) (Fig. 2, A). In addition, 22 days after injection (when the experiment was terminated because of the formation of large tumors in the control group), mice injected with pHIF-1{alpha}DN–transfected cells had statistically significantly lower mean tumor weights than mice injected with pCEP4-transfected cells (DN2 versus pCEP4: 0.08 g versus 0.70 g, difference = 0.62 g, 95% CI = 0.23 to 0.93 g; P = .002; DN3 versus pCEP4: 0.05 g versus 0.70 g, difference = 0.65 g, 95% CI = 0.22 to 0.95 g; P = .004) (Fig. 2, B). Results of MTT and FACS analyses showed that, in vitro, pCEP4- and pHIF-1{alpha}DN–transfected cells had similar growth rates and cell cycle parameters under nonhypoxic conditions (data not shown), which demonstrates that the differences in in vivo growth rates were not due to inherent differences in cell proliferation rates.



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Fig. 2. Effects of HIF-1{alpha} inhibition on tumor growth in a subcutaneous xenograft tumor model. Human gastric cancer TMK-1 cells stably transfected with pHIF-1{alpha}DN (clone DN2 or DN3) or empty vector (pCEP4) were injected into the subcutis of mice (10 mice per group). Tumor volumes (A) were measured every other day when palpable tumors were first detected on day 8 after injection, and tumor weights (B) were determined on day 22 after injection, when the experiment was terminated (*P = .003 for DN2 and P = .004 for DN3 versus pCEP4). Inset shows representative excised tumors from the pCEP4 group and HIF-1{alpha}DN2 group. Scale bar indicates 10 mm. Error bars indicate 95% confidence intervals.

 
Effect of Dominant-Negative Mutant HIF-1{alpha} on Xenograft Tumor Angiogenesis, Vessel Morphology, and Cell Proliferation In Vivo

We next used immunohistochemical staining to investigate the effect of inhibiting tumor cell HIF-1{alpha} function on in vivo angiogenesis and vessel morphology in the subcutaneous tumor xenografts. Mice injected with pCEP4-transfected cells had tumor vessels that appeared mature (i.e., they had thick walls), displayed strong expression of CD31 along their lengths, and had open lumina. By contrast, mice injected with pHIF-1{alpha}DN–transfected cells had tumor vessels that appeared smaller and thinner and had less branching. However, there was no statistically significant difference in the mean number of tumor vessels per HPF (classical microvessel density) between mice injected with DN2 or DN3 cells and mice injected with pCEP4-transfected cells (Fig. 3, A and 4).



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Fig. 3. Effect of HIF-1{alpha} inhibition on tumor angiogenesis. Frozen sections of subcutaneous tumors were stained for CD31. A) Vessel density was determined by counting the number of CD31-stained vessels per high-power field (HPF) in four quadrants in each tumor (at 2 mm from inside the tumor–normal tissue interface). B) CD31-positive vessel area per HPF (i.e., 103 pixels2) was measured in tumor sections using optical image analysis software. *, P<.001 for DN2 versus pCEP4 and for DN3 versus pCEP4. Error bars indicate 95% confidence intervals.

 


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Fig. 4. Immunohistochemical analysis of angiogenesis, tumor cell proliferation, and immunofluorescence analysis of pericyte-like cell coverage of tumor endothelial cells in subcutaneous xenograft tumors derived from pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells. Tumor sections were stained with hematoxylin–eosin (H & E), with an anti-CD31 antibody (CD31) to detect vessel-associated endothelial cells (brown), with an anti-BrdU antibody (BrdU) to detect proliferating tumor cells (brown), or with an anti-CD31 antibody (red) plus either anti-{alpha}SMA or anti-desmin antibodies (green) to evaluate pericyte-like cell coverage of tumor endothelial cells. Scale bars indicate 100 µm for H & E and CD31 and 50 µm for BrdU, CD31/{alpha}SMA, and CD31/desmin.

 
Vessel area is a critical determinant of blood flow (Poiseuille's law). Therefore, we investigated the observed differences in vessel morphology by determining the area of CD31-positive–staining structures in each tumor section by optical image analysis. In addition, we performed vessel counts as described above. Tumors derived from pHIF-1{alpha}DN–transfected cells had a smaller mean CD31-positive vessel area than those derived from pCEP4-transfected cells (DN2 versus pCEP4: 1.0 x 103 pixels2 versus 3.7 x 103 pixels2, difference = 2.7 x 103 pixels2, 95% CI = 1.97 to 3.42 x 103 pixels2; P<.001; DN3 versus pCEP4: 1.2 x 103 pixels2 versus 3.7 x 103 pixels2; DN3: difference = 2.5 x 103 pixels2, 95% CI = 1.84 to 3.15 x 103 pixels2; P<.001) (Fig. 3, B and 4). In addition, double immunofluorescence staining for CD31 (an endothelial cell marker) and {alpha}SMA (a pericyte marker) revealed that most of the vessels in the pCEP4-derived tumors (especially those with open lumina) were closely associated with pericyte-like cells, whereas vessels in the pHIF-1{alpha}DN–derived tumors exhibited limited association with pericyte-like cells (Fig. 4, fourth row). These results were subsequently confirmed by costaining for CD31 and desmin (another pericyte marker): Mean tumor endothelial cell coverage by desmin-positive cells (green staining in bottom row of Fig. 4) was statistically significantly lower in tumors derived from DN2 cells (32%) or DN3 cells (37%) than in tumors derived from pCEP4-transfected cells (80%) (difference for DN2 versus pCEP4 = 48%, 95% CI = 33 to 63%; P<.001; difference for DN3 versus pCEP4 = 43%, 95% CI = 22 to 64%; P = .001) (Fig. 4, fifth row).

The recruitment of pericytes and their contribution to vessel maturation, morphology, and function is regulated by specific cytokines, such as PDGF-BB and Ang-2 (48,5053). Conditioned media from pCEP4-transfected and pHIF-1{alpha}DN–transfected cells had similar (i.e., low) concentrations of PDGF-BB and of Ang-2, regardless of whether the cells were incubated in nonhypoxic or hypoxic conditions (data not shown). Furthermore, in a gene array analysis, we detected no statistically significant changes in expression levels of genes involved in vessel maturation between cells transfected with pCEP4 and cells transfected with pHIF-1{alpha}DN under both hypoxic and nonhypoxic conditions (data not shown). Finally, we analyzed xenograft tumor sections for apoptosis (using TUNEL) and found no statistically significant differences in the mean number of apoptotic cells between tumors derived from pCEP4-transfected cells and tumors derived from pHIF-1{alpha}DN–transfected cells (data not shown). By contrast, tumor cell proliferation (as defined by the mean number of BrdU-positive cells per HPF) was statistically significantly lower in tumors derived from DN2 cells than in tumors derived from pCEP4-transfected cells (18.7 cells/HPF versus 50.9 cells/HPF, difference = 32.2 cells/HPF, 95% CI = 27.6 to 36.7 cells/HPF; P<.001) (Fig. 4, third row).

Effect of Dominant-Negative Mutant HIF-1{alpha} on Orthotopic Gastric Tumor Growth and Angiogenesis

To validate the results obtained using the subcutaneous xenograft tumor model, we next examined the effects of HIF-1{alpha} inhibition on tumor growth and angiogenesis in an orthotopic model of gastric cancer. Mice were injected in the greater curvature of the gastric wall with either pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells (clone DN2 or DN3) to initiate the formation of gastric wall tumors that did not obstruct the lumen. By 45 days after the injections, mice in the pCEP4 group had become lethargic, and we terminated the experiment. Tumor incidence was similar among the three groups of mice (75% in the pCEP4 group versus 60% in the DN2 group [P = .662] versus 70% in the DN3 group [P = 1.00]; Fisher's exact test)]. Among tumor-bearing mice, those injected with DN2 cells or DN3 cells had statistically significantly smaller mean tumor volumes than those injected with pCEP4-transfected cells (DN2 versus pCEP4: 75 mm3 versus 756 mm3, difference = 681 mm3, 95% CI = 81 to 1280 mm3; P = .029; DN3 versus pCEP4: 88 mm3 versus 756 mm3, difference = 668 mm3, 95% CI = 127 to 1209 mm3; P = .019) (Fig. 5, A). Figure 5, B shows representative images of orthotopic gastric tumors that developed in mice injected with pCEP4-transfected TMK-1 cells or DN2 cells.



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Fig. 5. Effect of HIF-1{alpha} inhibition on gastric tumor growth in an orthotopic model. A) pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells were injected into the gastric wall of nude mice; 45 days later, mice in the pCEP4 group were moribund and the experiment was terminated. We measured gastric tumor diameters and calculated the tumor volumes among tumor-bearing mice. *, P = .029 for DN2 versus pCEP4; #, P = .019 for DN3 versus pCEP4. Error bars indicate 95% confidence intervals. B) Representative images of gastric tumors (circled area) derived from pCEP4- and pHIF-1{alpha}DN–transfected TMK-1 cells Scale bar = 10 mm.

 
Tumor sections were also stained for CD31 to investigate the effect of HIF-1{alpha} inhibition on angiogenesis in orthotopically implanted gastric tumor cells. As we observed for the subcutaneous xenograft tumors, there was a substantial difference in vessel morphology between orthotopic gastric tumors derived from pCEP4-transfected cells and those derived from pHIF-1{alpha}DN–transfected cells. For example, the mean CD31-positive area was smaller in tumors derived from DN3 cells than in tumors derived from pCEP4-transfected cells (2.30 x 103 pixels2 versus 8.63 x 103 pixels2, difference = 6.33 x 103 pixels2, 95% CI = 5.10 to 7.43 x 103 pixels2; P<.001) (Fig. 6, A and B). In addition, double immunofluorescence staining of orthotopic tumor sections for CD31 and {alpha}SMA revealed that tumors derived from DN2 or DN3 cells had fewer pericyte-like cells associated with tumor endothelial cells (22% and 32%, respectively) than vessels in tumors derived from pCEP4-transfected tumors (58%) (difference between pCEP4 and DN2 = 36%, 95% CI = 13% to 58%; P = .004; difference between pCEP4 and DN3 = 26%, 95% CI = 1.6% to 50%; P = .038).



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Fig. 6. Effect of HIF-1{alpha} inhibition on angiogenesis in gastric tumors. Frozen sections of orthotopic gastric tumors derived from injection of pCEP4- or pHIF-1{alpha}DN–transfected TMK-1 cells were stained with anti-CD31 antibody. A) CD31-positive vessel area was measured per high-power field (i.e., 103 pixels2). *, P<.001 for DN2 and #, P<.001 for DN3 versus pCEP4; DN3 is shown. Error bars indicate 95% confidence intervals. B) Representative images of CD31-stained (brown) sections of tumors derived from pCEP4- and pHIF-1{alpha}DN–transfected TMK-1 cells. Scale bar = 100 µm.

 

    DISCUSSION
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We found that inhibition of HIF-1{alpha} function in human gastric cancer cells by stable overexpression of a dominant-negative mutant HIF-1{alpha} resulted in the reduction of VEGF secretion in vitro, the inhibition of tumor growth and angiogenesis in vivo, and alterations in tumor vessel morphology and maturation in vivo. Whereas the effect of HIF-1{alpha} inhibition on VEGF expression in in vitro assays has demonstrated mostly concordant results in the literature (i.e. reduction of VEGF), its impact on the regulation of tumor growth and angiogenesis in vivo has been controversial [reviewed in (17)]. However, Blouw et al. (43) recently showed that the growth-inhibitory effects of dysfunctional HIF-1 complexes depend on the tumor microenvironment. Their results suggested that the suitability of HIF-1 as a molecular target in cancer therapy should be tested in orthotopic tumor models.

We found that in two different experimental tumor models (i.e., a subcutaneous xenograft model and an orthotopic model), inhibition of HIF-1{alpha} function was associated with the inhibition of gastric tumor growth and angiogenesis. In addition, HIF-1{alpha} inhibition resulted in dramatic changes in vessel morphology and maturation, suggesting that HIF-1 is an important (if indirect) regulator of vascular biology. Few published studies have examined the role of HIF-1{alpha} in mediating this aspect of angiogenesis. In three different xenograft studies, blockade of HIF-1 was found to have tumor growth–inhibitory effects in vivo (41,42,54). Kung et al. (41) demonstrated that interference with HIF-1 binding to p300 reduced the subcutaneous growth of human colon cancer HCT116 cells; however, vessel counts or changes in vessel morphology were not reported. Maxwell et al. (50) showed that human hepatoma Hepa-1 cells genetically modified to express reduced levels of functional HIF-1{beta} were associated with reduced tumor growth and angiogenesis (as measured by microvessel density) compared with wild-type Hepa-1 cells. Chen et al. (42) reported that HIF-1{alpha} inhibition in human pancreatic cancer cells resulted in reduced subcutaneous growth of the pancreatic cancer cells; however, the authors did not observe substantial changes in VEGF expression or vessel density in these tumors.

The authors of these three studies did not report on vessel morphology or vessel area in the tumor specimens. However, results of two studies have suggested that HIF-1 inhibition may have an effect on vessel morphology (38,39). Carmeliet et al. (38) observed large dilated vessels in subcutaneous tumors formed by HIF-1{alpha}–expressing mouse ES cells (i.e., HIF-1{alpha}+/+ cells), resulting in a hemorrhagic appearance of excised tumors, whereas tumors formed by stem cells that did not express HIF-1{alpha} (i.e., HIF-1{alpha}–/– cells) appeared pale and lacked vessels. Tsuzuki et al. (38) reported that vessels in tumors derived from HIF-1{alpha}–/– embryonic stem cell were smaller, less functional (i.e., had reduced blood flow), and more permeable to fluorescein isothiocyanate–labeled dextran than tumors derived from HIF-1{alpha}+/+ cells, despite having only modest decreases in VEGF levels. These observations suggest that HIF-1{alpha} may mediate several functions of the tumor neovasculature, including vessel morphology and pericyte coverage of endothelial cells.

HIF-1{alpha} regulation of vessel maturation and function in tumors has not been well investigated. We therefore evaluated tumor neovasculature by measuring CD31-positive area in tumor sections in an attempt to measure a surrogate marker of the functional vascular network, because blood flow is associated with vessel area. Because vessel area is a critical determinant of blood flow (Poiseuille's law) and because functional vessels (mature vessel, open lumen, periendothelial support) are essential for blood flow in microvasculature, we addressed the issue of vessel maturation by analyzing periendothelial cell (pericyte) coverage of tumor endothelial cells. To date, pericyte coverage of tumor endothelial cells as an indicator of vessel maturation (50,52) and functionality has not been investigated in tumors having impaired HIF-1{alpha} function. We found that inhibition of HIF-1{alpha} activity in tumor cells reduced coverage of tumor endothelial cells by pericyte-like cells. In our study, the angiogenic factors Ang-2 and PDGF-BB, which have been reported to affect endothelial cell stability or to increase pericyte migration (PDGF-BB) (53,5558), were expressed at similar levels in vitro in the transfected cell lines. In addition, we detected no differences in the expression of genes for angiogenic factors that have been associated with pericyte migration or recruitment between cells that stably expressed a dominant-negative mutant HIF-1{alpha} and those that did not. However, the complexity of the tumor–host interaction in angiogenesis (and expression of angiogenic molecules) may not be reflected by in vitro assays. For example, it is possible that HIF-1 may mediate vessel maturation through in vivo intercellular cross-talk, as previously described (59).

We found that inhibition of HIF-1{alpha} function reduced nonhypoxic VEGF expression in vitro. As reported by others (3236), HIF-1{alpha} may be constitutively overexpressed in tumor cells by hypoxia-independent mechanisms. We speculate that HIF-1 is constitutively active in TMK-1 cells but at levels below the limits of detection for western blot analysis of nuclear extracts.

On the basis of our results and the reported implications of HIF-1 in tumor progression and invasion, HIF-1 may be a promising target for cancer treatment (17,21,22,60,61). Several agents that affect HIF-1 or HIF-1{alpha} function have been described [reviewed in (17)]. For example, Mabjeesh et al. (62) showed that the natural estrogen metabolite 2-methoxyestradiol disrupts microtubules in tumors, inhibits tumor growth and angiogenesis in vivo, decreases HIF-1{alpha} expression, and inhibits HIF-1{alpha} function. A substance used in the treatment of circulatory disorders, YC-1 [3-(5'-hydroxymethyl-2'-furyl)-1-benzyl indazole], has been shown to inhibit HIF-1 activity, platelet aggregation, and vascular contraction in vitro (63). Moreover, treatment of various types of human cancer cells with YC-1 led to inhibition of subcutaneous tumor growth, reduced HIF-1{alpha} expression in vivo, and reduced tumor vascularity (63). However, because neither agent is specific for HIF-1, their clinical applicability in cancer patients is uncertain at this time.

In summary, our results suggest that HIF-1{alpha} not only regulates VEGF expression in cancer cells but also contributes to the formation of a complex proangiogenic microenvironment in tumors, thereby affecting vessel morphology and, ultimately, vessel function. Inhibition of HIF-1{alpha} function decreased the growth of orthotopically injected human gastric cancer cells in vivo, suggesting that HIF-1{alpha} is a valid target for the treatment of gastric cancer. Finally, the method of transfecting tumor cells with a dominant-negative construct reported in this study may be a powerful tool for obtaining further insight into HIF-1–regulated gene expression and the functional impact of HIF-1 blockade on cancer cells.


    NOTES
 Top
 Notes
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Supported in part by NIH grants T-32 09599 (JSW, LME), and CA74821 (LME) and NIH CCSG CA 16672 and NIH P50-CA103175 (GLS).

We thank Christine Wogan from the Department of Scientific Publications and Rita Hernandez from the Department of Surgical Oncology at M. D. Anderson for editorial assistance. We also thank Donna Reynolds and Carol Oborn, both from the Department of Cancer Biology at M. D. Anderson, for technical assistance.


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 Materials and Methods
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 Discussion
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Manuscript received October 14, 2003; revised April 9, 2004; accepted April 21, 2004.


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