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Experimental Therapeutics |
Department of Anatomy and Cell Biology, University of Bergen, 5009 Bergen, Norway [T-A. R., R. B., P. C. H.]; Department of Neurosurgery, Klinikum Mannheim, 68167 Mannheim, Germany [M. F., P. V.]; Harvard Medical School, Boston, Massachusetts 02155 [B. R. O.]; and Institute of Cancer Research and Molecular Biology, The Norwegian University of Science and Technology, N-7489 Trondheim, Norway [A. M. R.]
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ABSTRACT |
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 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Tumor growth was reduced by 35% in treated animals. It was of interest that tumor cell invasion into the surrounding tissue was also inhibited. The total vascular density was reduced by 67.6%, perfusion by 67%, and vessel diameters by 37%. This resulted in a significant reduction in tumor perfusion, although the vessel permeability was not influenced.
We have demonstrated that human endostatin not only reduces total vascular density, as shown previously, but also greatly reduces the functionality and the diameters of the vessels. Furthermore, we show that this therapeutic approach also inhibits tumor cell invasion, thus supporting the hypothesis that tumor angiogenesis and invasion represent two interrelated processes. Finally, this work further confirms the new therapeutic concept using alginate cell-encapsulation technology for the localized delivery of therapeutic compounds to central nervous system malignancies.
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Endostatin has been shown to inhibit the growth of a number of different ectopic tumors (10, 11, 12, 13, 14, 15, 16) , induce apoptosis in bovine pulmonary endothelial cells (C-PAE) in vitro (17) by activation of tyrosine kinase (18) and does not compete with fibroblast growth factor-2 or VEGF3 binding (19 , 20) .
However, the effects of endostatin on tumor microcirculation in vivo has not been thoroughly described because authors tend to report reduction in vessel count only, which does not predict vascular function and oxygenation of the tumor.
We have recently introduced a new therapeutic concept for malignant brain tumors based on the encapsulation of endostatin secreting 293 HEK cells in an immuno-protective device (bioreactor) consisting of sodium alginate (21) . Transplantation of such endostatin-secreting bioreactors together with BT4C gliosarcoma cells into the rat brain significantly prolongs animal survival (21) .
In the current study, we demonstrate using intravital fluorescence video microscopy that local release of endostatin from alginate bioreactors not only reduces vascular density but also largely reduces vascular diameters and perfusion in C6 gliomas. These findings thus suggest additional antiangiogenic mechanisms of endostatin, which may in part explain the potent antitumor action. For a direct comparison between ectopic and orthotopic tumors, dorsal skin-fold chambers as well as a cranial window model were used. Furthermore, we demonstrate that glioma cell invasion into the surrounding tissue was significantly inhibited, adding a new feature to the antitumor activity of endostatin.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Spheroid Cultures.
Spheroids were initiated as described elsewhere (22)
. Briefly, C6 monolayer cultures were trypsinized and seeded into culture flasks base coated with 0.75% agar noble in DMEM. On the day of implantation, the spheroids where stained with the vital dye DiI (Molecular Probes, Eugene, OR), which is a lipid soluble fluorescent dye that is incorporated into the cell membrane, and distributed between daughter cells upon division (23, 24, 25)
. DiI (2.5 mg) was dissolved in 1 ml of ethanol (100%) and diluted in DMEM to obtain a concentration of 0.075 mg/ml after sonication. The spheroids where stained by placing them on agar-coated, 24-well plates with an overlay of 1 ml of DiI for 4 h at 37°C.
Encapsulation of 293 Cells in Alginate.
The method of encapsulation has been described in detail elsewhere (21
, 26
, 27)
. Briefly, droplets of cells dispersed in 1.5% LVG sodium alginate (Pronova Biomedical, Torshov, Norway; 2 x 107 cells/ml alginate) were released into a 0.1 M CaCl2 solution prepared from 0.13 M NaCl stock solution, initiating gel-formation by cross-linking. The resulting alginate beads were washed three times in DPBS (Sigma Chemical Co. St. Louis, MO) and once in the growth medium described previously. The encapsulated cells were cultured in 175-cm2 culture flasks containing 50 ml of growth medium, under the same conditions as described above.
The viability of the encapsulated cells was determined using a two-color fluorescence viability assay (Live/Dead Viability/Cytotoxicity Assay; Molecular Probes, Eugene, OR) according to the protocol provided by the manufacturer. Green fluorescence is emitted from the intracellular esterase-converted calcein in viable cells, whereas red fluorescence indicates dead cells (26) . Fluorescence was measured in optical sections through the alginate using a confocal scanning laser microscope with a krypton-argon laser (Leica TCS-NT, Heidelberg, Germany), using rhodamine and FITC filter optics. Fluorescence was recorded in a plane between 70 and 120 µm inside the beads.
Western Blotting.
To verify that endostatin was released from the beads, conditioned medium from encapsulated endo-293 cells was collected and used for standard SDS-PAGE Western blotting (Ref. 28
; endostatin antibody concentration, 1:1000).
In Vitro Studies.
To evaluate whether endostatin had any cytotoxic or antimigratory effects on the tumor cells, monolayers were exposed to endostatin-secreting alginate beads. C6 cells were seeded into 24-well culture plates (8000 cells/well, 5 parallels at each time point, and 5 time points) and were allowed to adhere before being exposed to either clear alginate beads or beads containing endo-293 or 293-EBNA cells (mock transfected). The cells were counted every second day for 10 days, using a Coulter counter (Coulter Electronics, Ltd., Herpenden Herts, United Kingdom), and growth curves were established.
A similar assay was chosen for studying migration, where C6 spheroids were generated as mentioned above and thereafter allowed to adhere to the plastic surface of 24-well plates (Nunc). The area of migration was studied as described previously (27) . At each time point, migrating spheroids were fixed in 4% paraformaldehyde, stained with hematoxylin, and photographed for documentation.
Animals and Dorsal Skin-Fold Chamber Model.
The animal experiments were approved by the ethics committee for animal research. Athymic nude mice (nu/nu; 18 male/female, 28–32 g) were bred and maintained within a specific-pathogen, germ-free environment. The technique for implantation of the dorsal skin-fold chamber has been described previously (29, 30, 31, 32, 33)
. Briefly, animals were anesthetized by s.c. injections of 7.5 mg of ketamine hydrochloride and 2.5 mg of xylazine per 100 mg of body weight. Two symmetrical titanium frames flanked the dorsal skin-fold of animals to sandwich the extended double layer of skin and create the dorsal skin-fold chamber, which consisted of one layer of striated muscle, s.c. tissue, and epidermis. An observation window, covered with a glass coverslip, allowed for repeated intravital microscopic observations. Two days after chamber preparation, the coverslip was temporarily removed, and a single tumor spheroid (200 µm in diameter) and four to six alginate beads (depending on the size) were placed at the surface of the striated skin muscle. Seven mice received alginate beads containing 293-endo cells, and 6 received clear alginate beads (controls). The animals tolerated the skin-fold chambers well and showed no signs of discomfort or changes in sleeping and feeding behavior.
Cranial Window Preparation.
The procedure for the preparation of the cranial window was similar to that described in previous reports (34)
. The heads of the animals were fixed in a rodent stereotactic frame (David Kopf Instruments, Tujunga, CA). The skin on top of the frontal and parietal skull was cut, and the underlying periosteum was scraped off to the temporal crests. Using an electrical high-speed drill with a burr tip size of 0.5 mm, a circular bone flap (0.7–1 cm in diameter) was created and freed from the underlying dura and sagittal sinus with the aid of a modified microdissector. The dura was removed with Iris microscissors, avoiding any damage to the sinus and bridging veins. Finally, the glioma spheroid (200 µm in diameter) and alginate bioreactors were directly placed onto either hemisphere, and the window was sealed with a glass coverslip adhered to the bone using a histocompatible glue.
Experimental Protocol.
Intravital multifluorescence microscopic studies of glioma growth, angiogenesis, and microcirculation were performed on days 3, 6, 10, 14, and 18 after implantation in dorsal skin-fold chambers and on days 3, 6, 10, and 14 after implantation in cranial windows. The newly formed microvasculature within the fluorescent glioma mass (intratumorally) and at the glioma periphery (peritumorally), i.e., outside the tumor and next to the tumor edge, were assessed separately. Vascular measurements included newly formed tumor microvessels only, which can be clearly distinguished from the autochthonous host striated muscle and cerebral microvessels by their chaotic arrangement and heterogeneous diameters (35
, 36) . In addition, it is possible to evaluate whether the vessels are in the tumor or in the normal tissue simply by changing the filters. Vascular densities were measured in four to six regions of interest per animal and per observation time point. Microvascular diameters as well as hemodynamic parameters were determined by analyzing 5–10 microvessels per region of interest. At the end of the in vivo experiments (14 and 18 days after implantation), the animals were sacrificed with an overdose of ketamine/xylazine, and the skinfold chamber preparations and brains were processed for histological and immunohistochemical analysis.
Intravital Multifluorescence Video Microscopy.
Intravital multifluorescence video microscopy (epi-illumination) was performed using a modified Axiotech Vario microscope with a blue (450–490 nm) and green (520–570 nm) filter block (22)
. Observations were made using x3.2 short distance, x10 long distance, and x20 water immersion working objectives (all from Zeiss, Oberkochen, Germany), resulting in magnifications of x50, x200, and x400, respectively. The glioma cells and microvasculature were visualized by means of a low-light level charge-coupled device video camera (CF8/1 FMC; Kappa, Gleichen, Germany; Ref. 22
). Microscopic images were recorded using an S-VHS video system (Panasonic, Munich, Germany) for off-line analysis. DiI labeling of glioma cells allowed for a precise delineation of the spheroid from the surrounding, unaffected host tissue as well as identification of individual tumor cells applying green light epi-illumination (22)
. By contrast enhancement with 2% FITC-conjugated dextran (0.1 ml of FITC-dextran150 i.v. via tail vein; Mr 150,000; Sigma Chemical Co., St. Louis, MO) and use of the blue light epi-illumination, angiogenic sprouts, individual microvessels, and, finally, the glioma microvasculature was visualized.
Analyzed Parameters.
Quantitative analysis of intravital microscopic observations was performed by a computer-assisted image analysis system (33)
. Tumor growth and tumor cell migration were assessed by measurement of the tissue area (mm2) covered by the fluorescent solid tumor mass and the migrating tumor cells, respectively. Analysis of microcirculatory parameters include the total tumor vascular density (cm-1), which was defined as the length of all newly formed microvessels per area of interest and observation time point, the functional vascular density (cm-1), which was defined as length of RBCs perfused, microvessels per area of interest, and observation time point and microvessel diameters (µm). To assess the permeability of the vessels, we compared intravascular with extravascular fluorescence intensity of multiple individual tumor microvessels in both peritumoral and intratumoral areas and calculated a permeability index as follows: permeability index = intravascular fluorescence/extravascular fluorescence. Finally, the vascular surface is defined as the calculated mean area covered by the tumor vessels per total area of observation. Thus, vascular surface is the product of vessel density (cm/cm2) and vessel diameter (µm; Ref. 36
).
Statistical Analysis.
Quantitative data are given as mean values ± SD. Mean values of microcirculatory data were calculated from the average values in each animal. For analysis of differences between the groups, post-hoc unpaired Bonferroni t test was used following one-way ANOVA. Results with P < 0.05 were considered significant (*, mean ± SD, P < 0.05 versus control).
Histology and Immunohistochemistry.
At experiment completion, the dorsal skin-fold chamber preparations and brains were dissected free and frozen in liquid nitrogen for histological and immunohistochemical analyses. The sections were mounted on stubs, embedded in Tissue-Tek (Miles Laboratories, Inc., Naperville, IL), frozen in 2-methylbutane (E. Merck, Darmstadt, Germany), and cooled with liquid N2.
Serial axial sections (10 and 60 µm) were cut and mounted on slides precoated with gelatin (Sigma Chemical Co.). The sections were stained with Harris H&E G (Merck) according to standard procedures, mounted in Entellan (Merck), and examined with a Nikon Diaphot light microscope.
The sections were analyzed with regard to endostatin distribution and blood vessel density (von Willebrand factor/CD31). The immunostaining was performed according to standard procedures (37) . The following antibodies were used (1:100 dilution in DPBS):anti-von Willebrand factor (DAKO AS, Copenhagen, Denmark), anti-CD31 (Becton Dickinson, San Jose, CA), and antihuman endostatin, polyclonal (Chemicon, Temecula, CA). Species-appropriate, FITC-conjugated secondary antibodies were used for primary antibody detection (1:30 dilution in DPBS). Cell nuclei were stained by treating the sections with RNase (Sigma Chemical Co.; 1 mg/in DPBS), followed by a short exposure to propidium iodide (Sigma Chemical Co.; 50 µg/ml in DPBS). Finally, the sections were washed in DPBS and mounted with Vectashield (Vector Laboratories, Inc., Burlingame, CA).
All sections were viewed and evaluated using a Leica TCS NT confocal laser scanning microscope with an argon-krypton laser (Leica, Heidelberg, Germany) applying TRITC and FITC filter optics.
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RESULTS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Western blots of conditioned medium from encapsulated 293-endo cells verified endostatin secretion from the beads in vitro (Fig. 1B)
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The tumor take rate was 100%, as reported previously (22)
. On the day of implantation into both settings, the DiI-labeled glioma spheroids showed an even fluorescence, a smooth edge, and no apparent signs of passive displacement of individual cells (Fig. 2, A and D)
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. Some tumor cells even migrated beyond this peritumoral area, and the adjacent alginate beads (Fig. 2, G and H)
. In contrast, the cranial window-established microtumors showed markedly reduced cell density within the core, and more cells detached and migrated in a radial pattern from the spheroid, suggesting a higher migratory activity of tumor cells on day 3 within the orthotopic setting (Fig. 2, F and I)
. The view of this area using FITC optics revealed initial tumor-induced angiogenesis in the cranial window setting, characterized by vascular sprouts and a chaotic vascular network (Fig. 2, E and J)
, as also observed in the skin-fold chambers (Fig. 2C)
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Endostatin-secreting Alginate Bioreactors Inhibit Glioma Growth and Invasion.
Throughout the observation period, tumor growth in dorsal skin-fold chambers was significantly inhibited by endostatin. After 18 days, the tumors were 35% smaller than the control tumors (Fig. 3A)
. Surprisingly, tumor cell migration into peritumoral areas also appeared to be inhibited (Fig. 3B)
, although because of large SDs, this trend proved not to be statistically significant. Histological analysis at the end of the observation period, however, confirmed the trend in migration inhibition observed by intravital microscopy. As illustrated in Fig. 4
, the treated tumors show a clear-cut border between the tumor and host tissue (Fig. 4A)
, whereas the control tumors showed extensive infiltration of glioma cells into the adjacent muscle and s.c. tissue (Fig. 4B)
. In parallel, immuno-staining with antihuman endostatin antibody demonstrated that the locally released endostatin was not only located in the near vicinity of the beads but also in the more distant muscle and s.c. layers representing the invasion zone of control tumors (Fig. 4, C and D)
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. The endostatin-treated animals, however, displayed fewer, less tortuous, and branched vessels, resembling a granulation tissue-like microvasculature rather than tumor-induced angiogenesis (Fig. 5B)
. Tumor-induced angiogenesis was observed within both the intratumoral and peritumoral zones, although it was predominant in the latter. Consequently, this was also the area in which endostatin had the greatest inhibitory effect on angiogenesis (Fig. 5, C and D)
. At day 6, the total vascular density was already considerably reduced in the treated animals (Fig. 5C)
, showing >80% reduction in vessel density in the peritumoral areas. Furthermore, the functionality of the vessels in terms of perfusion was also greatly reduced (Fig. 5D)
, where again the peritumoral areas showed the most severe differences at day 6 with an almost 80% reduction in functional vessel density.
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. On average, the endostatin-treated vessels were 37% smaller in diameter than the controls, in both intra and peritumoural areas (Fig. 6A)
. In line with the observations regarding reduction in tumor size and total vascular density, the vascular exchange surface of the endostatin-treated tumors was 
50% smaller than the control tumors (Fig. 6B)
. Of interest, despite the inhibitory effects of endostatin on tumor angiogenesis and tumor microcirculation, microvascular permeability was not affected and remained high for the macromolecular fluorescent marker FITC-dextran (Fig. 6B)
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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It has, for example, been suggested that human endostatin does not inhibit murine or bovine endothelial cell migration/proliferation in vitro (39 , 40) and that circulating human endostatin has no antiangiogenic activity at all (41) . In contrast to this, it has also been reported that human endostatin inhibits VEGF-induced human umbilical vein endothelial cell migration in vitro (20) and inhibits the growth of several different rodent tumor models in vivo (21 , 42 , 43) . In the current study, we show that localized delivery of recombinant human endostatin from encapsulated 293-endo cells not only inhibits solid tumor growth but also affects invasion of tumor cell into the surrounding tissue. Furthermore, we show for the first time that endostatin not only affects the tumor vascular density but also greatly reduces vessel functionality, diameters, and hence microvascular perfusion in both an ectopic and orthotopic setting.
The concept of cell encapsulation and consequent implantation in the central nervous system for the treatment of brain tumors is novel, and we have shown recently that an experimental rat intra-cerebral gliosarcoma model (BT4C) can be treated with endostatin-secreting bioreactors, resulting in prolonged survival of the animals (21) .
The results presented herein show that this method of sustained localized therapy is not only applicable within the brain parenchyma but also on the cerebral cortex and in s.c. and muscle tissue. Both skin-fold and cranial window preparations seemed to tolerate the alginate implants well, showing no apparent signs of immune reaction in forms of increased leukocyte count/endothelial interactions or edema (data not shown). Furthermore, the physiological angiogenic response to the grafts was not inhibited by recombinant human endostatin; in fact, these vessels grew around and over the bioreactors (data not shown), suggesting that the antiangiogenic response is tumor specific. This is essential in predicting long-term viability of the encapsulated 293-endo cells because inhibition of physiological angiogenesis would also affect nutrient supply to the bioreactors.
One of the most interesting findings in this study was that the tumor cell migration/invasion pattern was different in the dorsal skin-fold chambers of treated animals (Fig. 3)
. This is in contrast to the in vitro proliferation and migration assays, which did not reveal inhibitory effects of endostatin (Fig. 7)
. When the skin-fold chambers were immunostained for human endostatin, the protein was found to be located as expected, in the immediate vicinity of the alginate bioreactors (Fig. 4C)
. However, distant deposits were also observed in the muscle and s.c. layers of the skin, into which the tumor cells did not invade (Fig. 4D)
. Because the exact mechanism behind the antiangiogenic activity of endostatins remains unknown, any explanation offered for this anti-invasive effect would be purely speculative. However, a recent study by Rehn et al. (44)
suggests that integrins may be involved in the binding of endostatin. Furthermore, it is well known that many common features are shared by tumor and endothelial cells, such as cell adhesion pathways, proteolysis, and cell migration on extracellular matrix components (45)
. Accordingly, a recent article by Kim et al. (46)
indicates that endostatin may block both endothelial and tumor cell invasion in a 3-dimensional in vitro model by preventing activation of matrix metalloproteinase-2.
Alternatively, the inhibition of tumor cell invasion in vivo may be a secondary effect to inhibition of angiogenesis. A substantial number of reports associate tumor cell invasion with tumor-associated angiogenesis (22 , 47, 48, 49, 50) , although the mechanism for such a relationship remains unclear. It is not unlikely, however, that the new vessels that originate from the adjacent peritumoral tissue grow toward the main tumor mass, providing a trail on which tumor cells can migrate in the opposite direction. Therefore, if such inward growth of vessels is reduced, it may result in fewer cells migrating away from the tumor mass, hence a smaller area of migration.
However, regardless of whether inhibition of tumor cell invasion by endostatins was direct (perhaps via matrix metalloproteinase-2) or indirect (via angiogenesis inhibition), it may represent a new effective means to control local, diffuse infiltrative glioma growth.
At day 18, the total vascular density was reduced in the treated tumors by 31% in the intratumoral areas and >67% in peritumoural areas (Fig. 5A)
, which is not surprising because the peritumoural area commonly has the highest angiogenic activity (22
, 34, 35, 36)
. Furthermore, this confirms preexisting data on vascular effects of endostatin (10, 11, 12, 13, 14, 15, 16)
.
In addition, we report that after 18 days of treatment, >67% of the newly formed vessels were not functional (i.e., no RBC flow) and that the vessel diameters were also reduced by 37%, the latter of which is a hallmark for tumor vessel regression after VEGF neutralization (51) . These vascular effects of endostatin are previously unreported and contribute largely to the vast reduction in the vascular surface available for nutrient exchange, which was reduced from 30 to 12% at day 18. As mentioned previously, the vascular effects of endostatin were most severe in the peritumoral areas, and immunostaining with antihuman endostatin antibodies showed that most of the endostatin released from the bioreactors was indeed located beyond the main tumor mass. The reason for this regional distribution of endostatin is not clear. One explanation is that endostatin was not able to penetrate into the solid tumor mass. It is well known that the interstitial fluid pressure is high in the center of tumor and low at the periphery (52 , 53) . This pressure gradient facilitates interstitial fluid movement from the center of the tumor to the tumor periphery and surrounding tissues (52) . Therefore, any therapeutic molecule delivered at the tumor periphery would have to overcome this outward convection to diffuse into the tumor. Hence, it is possible that the endostatin delivered from the bioreactors was unable to reach the tumor center because of this unfavorable convection of interstitial fluid. Alternatively, it may well be that endostatin simply has an affinity for the basement membrane, as indicated in previous studies (19) , and therefore accumulates at the dermal-epidermal junction. Whatever the reason for the peripheral deposits of endostatin, it may not be imperative for antiangiogenic molecules to reach the central tumor mass, because it is commonly the peripheral tumor regions that show the highest angiogenic activity (35 , 36) . Hence, inhibition of tumor angiogenesis in the periphery may be sufficient to inhibit tumor growth (35 , 36) .
As mentioned earlier, the vascular effects of human endostatin were solely tumor associated, indicating that there may exist mechanistic/cellular differences between normal and tumor-induced angiogenesis. In fact, recent reports on differential gene expression in endothelial cells from normal and tumor tissue as well as cell lines show that endothelial cells from tumor tissue are quite different from that of normal tissue and cell lines (54) . It is suggested that tumor endothelium expresses genes commonly expressed only during physiological angiogenesis, such as in corpus luteum formation and wound healing. In addition, a few differentially expressed genes, such as tumor endothelial marker 8, have been identified in tumor endothelium (54) . This indicates that tumor endothelium has a genetic profile that is quite different from endothelial cells in normal tissue or endothelial cell lines. These data must be taken into consideration when interpreting previous reports on the failure of human endostatins to inhibit endothelial cell lines (both human, mouse, and bovine) in vitro, yet show inhibitory effects in murine models in vivo. As long as the receptor for endostatin remains unknown, it is clear that one should take great care in choosing the most representative model system in which to study the antiangiogenic effects of this molecule. Hence, in this study, we treated gliomas implanted in both dorsal skin-fold chambers and cranial windows. Our results show that the antivascular effects were comparable between the ectopic and orthotopic model. However, we failed to show statistical significance in tumor size between endostatin-treated and control cranial window tumors. The reason for this is not clear, because we have shown previously that intracerebrally implanted, endostatin-secreting bioreactors induce prolonged survival of rats bearing BT4C brain tumors (21) . The glioma spheroids implanted into the cranial window grow substantially faster than the dorsal skin-fold implants, and because the window is smaller, they quickly grow to a critical size with respect to chamber size (2 weeks). Given the aggressiveness of this tumor, this short growth period may not be enough for endostatin to induce significant antitumor effects. In fact, if one compares the dorsal skin-fold tumors, it becomes clear that endostatin has the greatest antitumor effects from day 14 and onward, at which point the cranial window tumors were already at a critical size where the animals had to be killed.
In conclusion, this study shows that not only does endostatin affect total vascular density, as demonstrated previously, but it also affects functional vascular density, diameter, and morphology, resulting in a large reduction of vascular surface and tumor perfusion.
In addition to the vascular effects, a significant inhibition of tumor cell invasion was observed, suggesting that this is either an indirect effect after the inhibition of angiogenesis or a direct effect on tumor cell migration. These findings propose novel/additional mechanisms for the antitumor efficacy of localized release of endostatin from cells encapsulated in alginate.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by Grant QLG1-CT-2000-00815 from the European Union commission, grants from The Norwegian Research Council, The Norwegian National Program for Gene Therapy, Innovest, and Haukeland Hospital, Grant DFG VA-151/4 from The German Research Foundation, and a grant from the Anne Magrethe and Per Jeager Research Foundation. 
2 To whom requests for reprints should addressed, at Department of Anatomy and Cell Biology, University of Bergen, Aarstadveien 19, 5009 Bergen, Norway. Phone: 47-55-58-63-73; Fax: 47-55-58-6360; E-mail: Tracy-Ann.Read{at}pki.uib.no
3 The abbreviations used are: VEGF, vascular endothelial growth factor; EBNA, EBV nuclear antigen; DiI, 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate; DPBS, Dulbecco’s PBS.
Received 4/11/01. Accepted 7/18/01.
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