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Tumor Biology |
Division of Surgical Oncology, Department of Surgery, Massachusetts General Hospital, and Harvard Medical School, Boston, Massachusetts 02114
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ABSTRACT |
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 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Tumor angiogenesis varies in different organ environments. For example, human renal carcinoma cells implanted under the renal capsule of nude mice produce 10–20 times more bFGF4 mRNA than when implanted into s.c. tissue (4) . Similarly, LS174T human colon carcinoma cells produce less VEGF mRNA when experimentally grown in the liver than when grown in s.c. tissue (5) . In addition, the endothelial cells that form new tumor blood vessels differ according to the host organ. New tumor blood vessels in the lung originate from capillary endothelial cells (2) , whereas new tumor blood vessels in the liver originate from sinusoidal endothelial cells (6) that, unlike lung capillary endothelial cells, are fenestrated and lack a basement membrane (7) . Because of these differences, the influence of antiangiogenesis factors on tumor growth may vary in different host organ environments. However, few studies have examined this aspect of angiogenesis.
One of the more potent antiangiogenesis agents, endostatin, is a cleavage product consisting of the COOH-terminal 184 amino acids of collagen XVIII (8) . This Mr 20,000 protein inhibits endothelial cell migration and proliferation (8, 9, 10, 11) , and induces the regression of a wide variety of tumors grown s.c. in mice (8 , 10 , 11) . The efficacy of endostatin in organ environments other than s.c. tissue is not well established. Endostatin has been shown in two studies to inhibit the formation or growth of lung metastases (11 , 12) . To our knowledge, there are no published reports on the efficacy of endostatin against liver metastases.
The aim of the present study was to determine the efficacy of endostatin in preventing tumor formation in different organ environments. We engineered stable transfectants from RenCa mouse renal carcinoma cells and SW620 human colon carcinoma cells that constitutively secrete mouse endostatin and determined the effect of endostatin on tumor formation in the flank, lung, and liver.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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chain signal peptide and upstream of a c-myc epitope and polyhistidine (His) tag. This plasmid was designated pEndoSTHB. DNA sequence analysis confirmed that the mouse endostatin cDNA sequence was inserted in the proper reading frame and without mutations.
Cell Lines.
The mouse renal carcinoma cell line RenCa (13)
was provided by Dr. K. Okumura (Jutendo University, Tokyo, Japan). The human colon carcinoma cell line SW620 was obtained from the American Type Culture Collection (Manassas, VA). RenCa and SW620 cells were maintained in DMEM supplemented with 10% fetal bovine serum, 100 units/ml penicillin, and 100 µg/ml streptomycin. HUVECs were obtained from Clonetics (San Diego, CA) and maintained in EGM-2-MV medium (Clonetics).
Generation of Stable Transfectants That Secrete Mouse Endostatin.
Endostatin transfectants were generated by transfection of pEndoSTHB into RenCa and SW620 cells using LipofectAMINE and LipofectAMINE PLUS Reagent (Life Technologies, Inc., Gaithersburg, MD) following the manufacturer’s instructions. Two days later, cells were placed under hygromycin selection at 800 µg/ml. Four weeks after transfection, hygromycin-resistant colonies were expanded and tested for endostatin-c-myc-His fusion protein production and secretion by immunofluorescence and Western blot analysis as described below. Control transfectants were generated in a similar manner, except the parent plasmid pSecTag2/HygroB (without endostatin cDNA) was substituted for pEndoSTHB.
Western Blot Analysis.
One million cells were plated onto 60-mm plates and incubated for 24 h. The medium was replaced with 1 ml DMEM, and cells were incubated for 24 h. One ml of conditioned medium was concentrated in a Microcon 10 microconcentrator (Amicon, Beverly, MA) to 20 µl and subjected to electrophoresis under reducing conditions on an 18% polyacrylamide gel. Proteins were transferred onto a nitrocellulose membrane (Schleicher & Schuell, Keene, NH) and incubated overnight in 5% nonfat milk in PBS at 4°C. After briefly washing in 1% nonfat milk and 0.1% Tween 20 in PBS, the membrane was incubated with anti-c-myc mouse mAb (Sigma Chemical Co., St. Louis, MO) diluted 1:100. After three 10-min washes in 1% nonfat milk and 0.1% Tween 20 in PBS, membranes were incubated in horseradish peroxidase-conjugated antimouse immunoglobulin (Amersham, Arlington Heights, IL) diluted 1:4000. After three 10-min washes in TBS containing 0.05% Tween 20, proteins were detected using the enhanced ECL kit (Amersham).
Immunofluoresence.
Cells were grown on eight-well glass slides (Lab-Tek, Naperville, IL), fixed in 4% paraformaldehyde in PBS, and permeabilized with 0.2% Triton X-100 in PBS. After washing in PBS, cells were incubated with either 1.25 µg/ml anti-His mouse mAb (Invitrogen, Carlsbad, CA) for RenCa cells or anti-c-myc mouse mAb (Sigma) diluted 1:200 for SW620 cells for 1 h. Cells were washed in PBS and incubated in 9.3 µg/ml FITC-conjugated antimouse immunoglobulin (Biosource, Camarillo, CA) in 10% nonfat milk in PBS for 1 h. After washing in PBS, cells were mounted in Vectashield (Vector Laboratories, Burlingame, CA) and photographed with an Axiophot immunofluorescence microscope (Carl Zeiss, Thornwood, NY) at x60. Background staining using the anti-c-myc mAb on RenCa cells was too high for accurate discrimination.
In Vitro Growth Rates.
Cells were plated onto 96-well plates at a density of 2000 cells/well in quadruplicate. At designated time points, the numbers of cells were quantified using a colorimetric MTT assay as described previously (14)
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Endothelial Cell Proliferation Assay.
Stable transfectants were plated onto 150-mm plates at a density of 5–10 million cells/plate and incubated for 48–72 h. The cells were washed with PBS, and 5 ml of DMEM were added to each plate. Cells were incubated for an additional 24 h, and 5 ml of conditioned serum-free medium were collected and concentrated to 1 ml using a Centriplus 10 concentrator (Amicon). HUVEC cells were plated at a density of 12,500 cells/well in 24-well collagen-coated plates for 24 h, after which 250 µl of concentrated, conditioned medium were added to each well for 30 min. An equal volume of Medium 199 supplemented with 10% fetal bovine serum and 5 ng/ml bFGF (Sigma) was then added for 72 h. The numbers of cells were then quantified using a colorimetric MTT assay. Tests were performed in quadruplicate.
Animal Studies.
Animal studies were performed in accordance with the policies of the Massachusetts General Hospital Subcommittee on Research Animal Care. Flank tumors were generated by s.c. injection of 5 x 106 cells in 100 µl of HBSS into the right flank of immune-competent BALB/c mice for RenCa cell lines or athymic BALB/c (nu/nu) mice (Charles River Laboratories, Wilmington, MA) for SW620 cell lines (n = 5/group). Tumor width (W) and length (L) were measured every 3–4 days with calipers. The tumor volume (TV) was determined by the following formula: TV = (L x W2)/2.
Experimental lung metastases were generated by injection of 5 x 105 cells in a single-cell suspension in 100 µl of HBSS without calcium or magnesium into the tail veins of immune-competent BALB/c mice (n = 5/group). Mice were sacrificed after 4 weeks and examined for lung metastases. Experimental liver metastases were generated by injection of 5 x 106 cells in a single-cell suspension in 100 µl of HBSS without calcium or magnesium into the spleens of nude mice (n = 5/group). Mice were sacrificed after 12 weeks to examine for liver metastases. Organs were fixed in 10% buffered formalin prior to being weighed and photographed.
To generate flank tumors from lung metastases, lung metastases from recently sacrificed mice were resected from the lung and soaked in PBS. The metastases were implanted into the s.c. flanks of BALB/c mice using a trocar. A single lung metastasis of 
2–3 mm in diameter was implanted into each mouse (n = 6/group). Tumor volume was determined every 4 days.
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RESULTS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. pEndoSTHB or pSecTag2/HygroB was transfected into RenCa and SW620 cells. Two endostatin-transfected RenCa clones (R
E15 and RE17) that secreted the expected Mr 29,000 protein and one control clone (R0) were selected (Fig. 1B)
. Similarly for the SW620 cells, two endostatin-transfected clones (SED2 and SED9) and one control clone (S0C1) were selected.
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and SW620 clones (Fig. 2d)
demonstrated cytoplasmic staining consistent with the presence of the endostatin-c-myc-His fusion protein prior to secretion. The control RenCa clone (Fig. 2a)
and control SW620 clone (Fig. 2c)
showed only minimal background staining.
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. In addition, SW620 stable transfectants and parental SW620 cells displayed similar in vitro growth rates (Fig. 3B)
. Thus, endostatin secreted into the culture medium by endostatin-expressing clones had no inhibitory effect on the proliferation of these nonendothelial cells.
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0 control cells did not inhibit HUVEC cell growth as compared with DMEM alone (Fig. 4)
. Conditioned medium from RE15 and RE17 endostatin-transfected cells inhibited HUVEC proliferation by 51 and 40%, respectively, compared with conditioned medium from R0 control cells. Similar results were obtained for conditioned medium from SW620 stable transfectants, with conditioned medium from SED2 and SED9 endostatin-transfected cells inhibiting HUVEC proliferation by 39 and 36%, respectively, compared with conditioned medium from S0C1 control cells.
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0 control cells formed tumors rapidly, but RE15 and RE17 endostatin-transfected cells formed s.c. tumors that were 73 and 84% smaller, respectively, compared with R0 tumors after 21 days (Fig. 5A)
. S0C1 control cells also formed tumors rapidly, whereas SED2 and SED9 endostatin-transfected cells formed tumors that were 91 and 82% smaller, respectively, after 21 days (Fig. 5B)
. Mice bearing R0 and S0C1 tumors were euthanized on day 21 because of tumor burden. Mice bearing RE15, RE17, SED2, and SED9 tumors were followed after 21 days, and their tumors eventually grew at an appreciable rate (data not shown). Seven to 8 weeks after initial tumor inoculation, mice with tumors from endostatin-transfected cells were euthanized because of tumor burden.
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0C1 control cells and SED9 endostatin-transfected cells were mixed in various ratios before s.c. implantation into nude mice. Tumor growth was equally inhibited, independent of whether 100, 75, or 25% of the cells expressed endostatin (Fig. 5C)
. In a separate experiment, each clone was injected s.c. into mice to generate tumors for immunohistochemical analysis. During the first 3 weeks after inoculation, flank tumors from endostatin-transfected cells were too small to harvest and section; therefore, tumors from endostatin-transfected cells were harvested between 3 and 5 weeks after inoculation, when they had grown beyond 2–3 mm in diameter. No immunohistochemical staining for endostatin-c-myc-His fusion protein was observed in any of the sections, suggesting down-regulation or loss of the endostatin transgene by the time these tumors had grown several mm in diameter (data not shown). Sections were also stained for tumor blood vessels using an anti-CD31 mAb. Blood vessel densities were determined and were not significantly different between tumors from endostatin-transfected clones and control clones (data not shown).
Mouse Endostatin Inhibits Formation of Lung Metastases.
To investigate the effect of endostatin on tumor formation in the lung, we injected 5 x 105 cells from each RenCa clone into the tail veins of BALB/c mice. The parental RenCa cells in this well-established model produce abundant lung metastases after 4 weeks (15)
. After sacrificing mice at 4 weeks, lungs from mice injected with RE15 and RE17 endostatin-transfected cells had few or no metastases, whereas lungs in mice injected with control cells had metastases that were too numerous to count (Fig. 6A
; Table 1
). Lungs from mice injected with R0 cells also weighed significantly more than lungs from mice injected with RE15 and RE17 cells. This experiment was repeated in 15 more mice with similar results (data not shown).
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E15 and RE17) and control cells (R0) were harvested and implanted s.c. into BALB/c mice. Although we had demonstrated previously that RE15 and RE17 endostatin-transfected cells grew poorly in the flank compared with R0 control cells after s.c. inoculation of a cell suspension (Fig. 5A)
, we observed different results after s.c. implantation of harvested pulmonary metastases. There was no significant difference in the size of flank tumors grown from R0 pulmonary metastases compared with those grown from either RE15 or RE17 pulmonary metastases (Fig. 6B)
. These data are consistent with down-regulation or loss of expression as an explanation for eventual tumor formation by endostatin-transfected cells.
Mouse Endostatin Prevents Formation of Liver Metastases.
We next examined the effect of endostatin expression on formation of experimental liver metastases. Others have demonstrated previously that SW620 cells injected into the spleens of nude mice generate experimental liver metastases in 60% of mice after 3 months (16)
. We injected 5 x 106 cells from each SW620 stable transfectant into the spleens of nude mice. After 12 weeks, the mice were sacrificed, and their livers were harvested. Three of the five livers from mice injected with S0C1 control cells developed liver metastases, whereas none of the 10 livers from mice injected with SED2 or SED9 endostatin-transfected cells developed liver metastases (Fig. 7A
; Table 2
). This experiment was repeated in 15 more mice with similar results (data not shown).
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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These results demonstrate the ability of endostatin to inhibit tumor or metastasis formation in different organ environments and do not examine the efficacy of endostatin against established tumors. A recent study by Bergers et al. (17)
found that in a transgenic mouse model of spontaneous 
-islet cell tumors, endostatin was highly effective in preventing progression of hyperplastic nodules to small tumors but was ineffective in inhibiting the growth of established invasive carcinomas. Achieving regression of established tumors is a more difficult undertaking than inhibiting tumor formation. Endostatin has been demonstrated to induce regression of tumors grown s.c. on the flanks of mice (8
, 10
, 11)
, but actual regression, as opposed to growth inhibition, has not been demonstrated in the lung or liver environments.
Gene transfer of endostatin and other angiogenesis inhibitors has been performed by other groups. Nguyen et al. (9) reported the use of an adeno-associated virus to transduce tumor cells with the endostatin gene in vitro and demonstrated that the secreted endostatin inhibits endothelial cell proliferation. In another study, plasmid DNA containing the endostatin cDNA was injected i.m. into mice and resulted in the inhibition of s.c. tumors and spontaneous pulmonary metastases (11) . In addition, s.c. inoculation of fibrosarcoma cells stably transfected with the gene for another angiogenesis inhibitor, angiostatin, resulted in inhibition of flank tumor growth and spontaneous pulmonary metastases (18) .
For the purposes of this study, stable transfection of the endostatin gene into cancer cells, followed by inoculation of these cancer cells into mice, avoided some potential problems associated with systemic administration of endostatin. Production of large amounts of biologically active recombinant endostatin to treat cohorts of mice can be difficult. In addition, the pharmacokinetics of endostatin absorption, distribution, and elimination after systemic administration is unknown and may result in different tissue levels in various organs, particularly in the liver, which is the primary site of metabolism of many drugs (19) . Although host organ environment may have influenced the expression of the endostatin gene from our endostatin-transfected clones, expression was enough at all sites tested to inhibit tumor formation.
Results of immunohistochemical staining studies of flank tumors indicated that endostatin expression was no longer detectable once tumors had grown beyond a small size. Endostatin expression may have been down-regulated or lost after tumor cell implantation into mice in the absence of hygromycin selection. Alternatively, proangiogenesis factors such as bFGF and VEGF may have eventually been up-regulated in these tumors, overcoming the effects of endostatin.
After flank inoculation of a cell mixture consisting of only 25% of endostatin-transfected cells, flank tumor growth was inhibited as much as when the inoculum consisted of 100% endostatin-transfected cells. However, we also observed that lung metastases from endostatin-transfected cells, after implantation into the flank, formed flank tumors as rapidly as lung metastases from control cells. These results may be consistent because an endostatin-expressing cell line likely represents a heterogeneous population in which some cells express little or no endostatin. When this heterogeneous population is implanted s.c., endostatin secreted by the majority of cells appears to prevent tumor formation by any immediately adjacent cells that do not express endostatin. In contrast, when inoculated i.v. as a single-cell suspension, individual cells that express little or no endostatin may successfully establish pulmonary metastases. This scenario presumes that the endostatin-producing cells that lay dormant in the lung do not produce levels of endostatin high enough to prevent metastasis formation elsewhere in the lung.
The observed reduction in tumor growth associated with endostatin secretion was most likely attributable to the biological activity of the fusion protein and not secondary to other causes. Clonal selection for a less tumorigenic phenotype is hypothetically possible but improbable for several reasons: (a) we used two different carcinoma cell lines and selected two endostatin-expressing clones from each cell line. We also selected hygromycin-resistant control clones that had been transfected with the same expression vector without the endostatin cDNA insert. The statistical likelihood that only the four clones of the six that we selected are incapable of in vivo growth for reasons unrelated to endostatin expression is exceedingly low; (b) we demonstrated that the expressed endostatin was biologically active against endothelial cells in vitro; (c) the control clones were clearly tumorigenic, yet implantation of a mixture of cells in which only 25% of cells expressed endostatin inhibited growth of the control cells. Host-immune response to the endostatin-c-myc-His fusion protein is also an unlikely cause of the observed results because significant antitumor effects were observed in both immune-competent hosts and congenitally athymic hosts. Ideally, we would have liked to perform experiments using endostatin-neutralizing antibodies. However, the absence of an endostatin antibody capable of neutralizing all of its biological activity precluded such an experiment. The anti-c-myc and anti-His mAbs reacted with the recombinant fusion protein but did not attenuate the ability of endostatin to inhibit HUVEC proliferation in vitro (data not shown). Accordingly, these antibodies could not be used for neutralization experiments.
Many issues regarding endostatin and other angiogenesis inhibitors remain unanswered. The molecular mechanisms by which endostatin exerts its effects are not well understood. Recently, Knebelmann et al. (20) reported that endostatin inhibits activation of mitogen-activated protein kinase, which is a downstream target in the bFGF and VEGF signaling pathways. Examination of the effect of endostatin on these pathways in different environments may shed more light on organ-specific differences in endostatin function. One drawback to gene transfer of endostatin into cancer cells prior to inoculation into mice is that this strategy did not allow us to study the efficacy of endostatin against established tumors in different organ environments. We are currently examining the efficacy of endostatin against established tumors in various organ environments using baculoviral and herpes simplex viral vectors to deliver the endostatin gene.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 This work is supported by NIH Grants CA64454 (to K. K. T.), GM21700 (to C-M. L.), DK43352 (core facilities), CA76656 (to S. U. S.), and CA71345 (to S. S. Y.) and the Claude E. Welch Research Fellowship (to S. S. Y.). 
2 Present address: Department of Urology, Kobe University School of Medicine, 7-5-1, Kusunoki-cho, Chuo-ku, Kobe 650-0017, Japan.
3 To whom requests for reprints should be addressed, at Division of Surgical Oncology, Department of Surgery, Massachusetts General Hospital, Cox 626, 100 Blossom Street, Boston, MA 02114. Phone: (617) 724-3868; Fax: (617) 724-3895; E-mail: ktanabe{at}partners.org
4 The abbreviations used are: bFGF, basic fibroblast growth factor; VEGF, vascular endothelial growth factor; CMV, cytomegalovirus: mAb, monoclonal antibody; MTT, thiazolyl blue; HUVEC, human umbilical vein endothelial cell.
Received 6/15/99. Accepted 10/19/99.
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REFERENCES |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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