Inhibition of Tumor Growth by the Antiangiogenic Placental Hormone, Proliferin-Related Protein

Nancy W. Bengtson and Daniel I. H. Linzer

Department of Biochemistry, Molecular Biology, and Cell Biology Northwestern University Evanston, Illinois 60208


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Proliferin-related protein (PRP) is a potent placental antiangiogenic hormone. To test the antiangiogenic potential of PRP to block tumor growth, we engineered tumor cells to express this hormone. Both SV40-transformed BALB/c mouse 3T3 fibroblasts and rat C6 glioma cells have markedly reduced growth rates as tumors in mice if they express high levels of PRP. In both models, the small tumors that form are largely avascular, whereas control tumors are rich in blood vessels, consistent with PRP limiting tumor growth by preventing neovascularization of the tumors. The antiangiogenic effects of PRP are also detected on human endothelial cells, suggesting that the receptor and signaling pathway of this mouse hormone are conserved between mouse and human and may represent useful targets for the development of antiangiogenic therapeutics. That signaling pathway appears to involve an inhibition of arachidonic acid release, based on the ability of arachidonic acid to overcome the antiangiogenic effects of PRP.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Angiogenesis, the growth of new blood vessels from existing vessels, is an essential process in both physiology and pathophysiology. During reproduction, remodeling of the maternal vasculature at the implantation site and the growth of fetal vessels into the placenta are required for the establishment of efficient gas, nutrient, and waste exchange. Two placental hormones in the PRL family, proliferin (PLF) and proliferin-related protein (PRP), contribute to this process in mice as positive and negative regulators, respectively, of angiogenesis (1). PLF appears to be important for attracting maternal endothelial cells toward the trophoblasts (2), and PRP is predicted to slow angiogenesis late in pregnancy, to prevent endothelial cells from resealing open vessels that create the maternal blood sinuses in which the trophoblasts bathe, and to limit the growth of maternal and fetal vessels across the placenta (3).

Both PLF and PRP circulate in the maternal bloodstream at high levels, PLF in early- to midgestation (4) and PRP in the latter part of gestation (5). Thus, the activities of these hormones may be directed both locally at endothelial cells in the uterus and placenta, and at more distal maternal targets. PLF also enters the fetal compartment (4), where it can bind to specific developing tissues (6). In contrast, PRP appears to be excluded from the fetus (6, 7), which would allow fetal vascularization to proceed in the absence of this antiangiogenic hormone.

The accumulation of PRP in the maternal circulation may be expected to have specific physiological consequences as part of normal mouse pregnancy but may also contribute to limiting any angiogenesis-dependent pathology. One such pathological condition is tumor growth (8). Avascular tumors grow slowly and depend on angiogenesis for a shift to a rapidly growing state (9, 10), whereas tumors placed into vessel-rich regions may have immediate access to a blood supply that supports rapid growth (11). Although tumors can grow independently to a small size before requiring an outside source of nutrients, more extensive growth depends upon the ability of tumors to attract endothelial cells by secreting angiogenic factors or by recruiting neighboring cells to secrete such factors. These endothelial cells will form new blood vessels and vascularize the tumor, thereby providing the tumor cells with nutrients and with access to the bloodstream, potentially leading to metastasis (12).

That a placental hormone might be responsible for the inhibition of tumor growth during pregnancy was suggested a few years ago by Gallo and colleagues (13), who pointed out that human immunodeficiency virus (HIV)-infected pregnant women were less prone to Kaposi’s sarcoma than nonpregnant women or males (13). In that report, human CG (hCG) was put forward as the placental factor responsible for the observed inhibition of tumorigenesis during pregnancy. Subsequent reports have demonstrated that an hCG-associated factor (HAF) and antineoplastic urinary protein (ANUP), but not hCG itself, are two components responsible for the observed effect on tumor cells in culture (14, 15, 16).

Despite the mistaken identification of hCG as the active factor in inhibiting tumor growth, it is likely that placental hormones provide a range of unusual activities as evidenced by the dramatic physiological changes that occur specifically during pregnancy. The activities of these placental hormones might then potentially be exploited to regulate pathophysiology in the nonpregnant state. PRP provides one such possibility. We therefore decided to test whether the antiangiogenic activity of PRP is able to restrict tumor growth. Furthermore, if any such activity is to prove useful in the design of novel antitumor therapies, the effect of this hormone on mouse endothelial cells must also be demonstrated to extend to human endothelial cells, a prediction that we have also now examined.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Generation of SVT2 Tumor Cell Lines Expressing PRP
To test the effect of PRP on tumor growth, an SV40-transformed BALB/c 3T3 fibroblast cell line (SVT2) was engineered to secrete PRP. The SVT2 cell line (17) was selected in part due to its rapid growth as a tumor when injected subcutaneously into BALB/c mice. In addition, this would provide a homologous system in which effects of a mouse hormone could be examined on mouse tumor cells, in an appropriate location for the tumor cell type, and in immunocompetent mice. A PRP expression construct containing the neomycin resistance gene was transfected into SVT2 cells, and G418-resistant cell lines were selected and then individually screened for the production of PRP. Four stable transfectants were chosen for further analysis based on their secretion of high levels of PRP (Fig. 1Go). PRP produced by stably transfected tumor cells is secreted, soluble, and of a higher apparent molecular mass than the 24 kDa predicted from its amino acid sequence. This higher apparent molecular mass is identical to that of PRP produced by stably transfected Chinese hamster ovary cells; this larger size has been demonstrated to result from N-linked glycosylation (5). Three of these stably transfected cell lines, designated SVT2-PRP2, SVT2-PRP3, and SVT2-PRP4, secrete quite high levels of PRP; by comparison to the purified protein control, PRP represents approximately 10% of the total secreted protein that accumulated in the conditioned medium from each of these cell lines. No PRP is detected in the conditioned medium of cultures of SVT2 cells stably transfected with the empty expression vector, indicating that SVT2 cells do not express the endogenous PRP gene.



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Figure 1. PRP Expression in SVT2-PRP Cell Lines

Conditioned media from stably transfected SVT2 cell lines were harvested and analyzed for PRP expression. Equal amounts (15 µg) of total protein from vector-transfected SVT2 cells (SVT2-V) or from four different SVT2-derived cell lines stably transfected with the PRP expression construct were loaded per lane. After transfer to a filter, the samples were incubated with an antiserum against PRP followed by an enzyme-linked secondary antibody. The leftmost lane contains 2 µg of purified PRP. The positions of the mol wt standards are indicated on the left.

 
Expression of PRP by the SVT2 tumor cells had no effect on cell growth in culture. All four cell lines were readily recloned in soft agar, indicating that SVT2 cells expressing PRP retain their fully transformed phenotype. Furthermore, the rate of cell growth for each of these cell lines was indistinguishable from the growth rate of the control, vector-transfected SVT2 cell line (Fig. 2Go). Thus, any effect that might be observed on tumor growth in vivo would not be attributable to direct effects of PRP on the growth of these cells.



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Figure 2. Cell Growth Curve of Stably Transfected SVT2 Cell Lines

SVT2-V and each of the four SVT2-PRP cell lines were plated at 7,000 cells per well on day 0 and allowed to grow under standard culture conditions. Cell number was determined in triplicate every day over a period of 7 days, and the mean ± SD is shown.

 
Inhibition of SVT2 Tumor Growth by PRP
Equivalent numbers of cells from each of the PRP-expressing SVT2 cell lines or the vector-transfected cell line were injected subcutaneously into the flanks of BALB/c mice. After 3 weeks, tumors were excised and analyzed; tumor growth was not maintained beyond this point because the size of the control tumors had reached the maximum allowed under the animal care guidelines. Each of the four SVT2 cell lines producing PRP resulted in tumors that were significantly reduced in volume and mass compared with the control (P < 0.01) (Fig. 3Go). Notably, even the SVT2-PRP1 cells, which express lower levels of PRP than the other cell lines, grew poorly as a tumor.



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Figure 3. PRP Inhibition of SVT2 Tumor Growth

Tumors formed from injected SVT2-V cells or from each of the four different SVT2-PRP cell lines were isolated after 3 weeks of growth, and their weights (A) and volumes (B) were determined. Values are presented as mean ± SEM with n = 7 to 12 mice per group; *, P < 0.01; **, P < 0.001.

 
Not all of the tumors from the SVT2-PRP lines were large enough for further study. In each case, for those tumors that could be analyzed, PRP mRNA was present in the tumor extracts (Fig. 4AGo) and PRP was secreted by tumor tissue placed in culture (Fig. 4BGo). Thus, tumors generated from the PRP-expressing SVT2 cell lines maintain PRP expression. The RNA and protein expression analyses also revealed that distinct tumors arising from a PRP-expressing SVT2 cell line display very similar levels of PRP expression, demonstrating that this approach leads to consistent hormone expression in vivo. In addition, compared with the purified PRP standard, the glycoprotein hormone produced by the tumor tissue was again similar in size and produced at high levels, indicating that no significant change occurred in PRP expression upon growth of the cell lines as tumors.



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Figure 4. PRP Expression in SVT2 Tumors

A, RNA (10 µg per lane) from an SVT2-V tumor and from two different tumors derived from each of two SVT2-PRP cell lines was purified and analyzed for PRP mRNA by filter hybridization. B, Tumor tissue samples were also placed in culture, and conditioned media (15 µg protein per lane) from these cultures were examined for PRP by immunoblotting. Three independent tumors from one SVT2-PRP cell line are shown to highlight the consistent levels of PRP expression. The leftmost lane contains 2 µg of purified PRP.

 
Tumors that developed from the three cell lines expressing high levels of PRP (SVT2-PRP2, SVT2-PRP3, and SVT2-PRP4) were uniformly small, solid masses. Closer inspection of these tumors by sectioning and staining for the endothelial cell marker PECAM-1 revealed a marked reduction in vascular density in the PRP-expressing tumors, in contrast to the vessel-rich control tumors (Fig. 5Go and Table 1Go). SVT2-PRP1 cells, which secrete lower levels of this antiangiogenic hormone, gave rise to tumors that were reproducibly soft and fluid filled. Histologically, these tumors have an outer vascularized region overlaying a region of high tumor cell density but lacking vascular structures, and a core that appears to have mostly cellular debris (Fig. 5Go).



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Figure 5. PRP Inhibition of SVT2 Tumor Vascularization

Tumor tissue was isolated and sectioned for detection of endothelial cells and vessels using an antibody against the endothelial antigen, PECAM-1. Many cross-sectioned vessels are present in tumor tissue from SVT2-V cells (A and B), several of which are indicated by the arrows, but not in high expressing SVT2-PRP tumors (C and D). SVT2-PRP1 tumor tissue (E) reveals a vascularized cortex (1 ), an underlying avascular and dense cellular layer (2, with a higher magnification of this region in panel F), and a core that appears to be filled with fluid and cellular debris (3 ). Antibody binding to mark endothelial cells is evident as a dark precipitate in the higher magnification of SVT2-V tumor tissue (B). Tissue sections were counterstained with Harris’s hematoxylin. The bar in the low magnification panels (A, C, and E) corresponds to 100 µm, and in the high magnification panels (B, D, and F) to 10 µm.

 

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Table 1. Vascular Density in PRP-Expressing Tumors

 
PRP Inhibition of C6 Glioma Tumor Growth
To verify that the effects of PRP on tumor growth were not restricted to a single model system, we repeated the above experiments with a second tumor cell line, C6 glioma cells (18). These cells also grow rapidly as a tumor and secrete high levels of angiogenic activity (19, 20, 21). Similar transfections were carried out with the PRP expression construct, and the resultant stable transfectants were screened for PRP expression by immunoblot analysis (Fig. 6Go). Again, a range of PRP secretion was observed among the isolated lines, and both high-expressing and low-expressing lines were chosen for further study. As with the SVT2-derived cell lines, all of the stably transfected C6 lines displayed identical growth kinetics in cell culture (data not shown), demonstrating that neither the transfection procedure nor the production of PRP was detrimental to cell growth.



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Figure 6. PRP Expression in C6-PRP Cell Lines

Conditioned media from stably transfected C6 cell lines were harvested and analyzed for PRP expression. Equal amounts (15 µg) of total protein from parental (C6) and vector-transfected (C6-V) cells or from two different C6-derived cell lines stably transfected with the PRP expression construct (one high expressing and one low expressing line) were loaded onto each lane, and after transfer to a filter the samples were incubated with an antiserum against PRP followed by an enzyme-linked secondary antibody. A sample of serum-free medium from placental cultures generated from tissue isolated at day 12 of gestation (day 12 PCM) was included as a positive control. The positions of the mol wt standards are indicated on the left; the arrow points to PRP.

 
Because C6 cells are of rat origin, immunodeficient ("nude") mice were used as hosts for the in vivo tumor growth analysis. The absence of hair on these mice also made it possible to monitor tumor size throughout the 3-week period after tumor cell injection. (In contrast, SVT2 tumor size in BALB/c mice could only be measured accurately at the end of the experiment.) Consistent with the results in the SVT2 system, two C6-PRP cell lines expressing high levels of the antiangiogenic hormone grew at significantly slower rates than the control C6 tumors (P < 0.05) (results are shown for one tumor line in Fig. 7Go); C6-derived cell lines expressing lower levels of PRP, though, grew at a rate indistinguishable from that of the parental and vector-transfected C6 controls (Fig. 7Go). The smaller tumors resulting from the C6-PRP high-expressing lines were largely avascular, as revealed by the histology of tumor tissue sections (Fig. 8Go) and immunostaining with the antiserum against the endothelial cell marker PECAM-1 to count vessels (Table 1Go). High PRP expression resulted in a greatly reduced density of larger vessels, but only a modest reduction in small vessels (Table 1Go); the latter category, though, includes small groupings of PECAM-1-positive cells even if a lumen is not evident and may therefore underestimate the effectiveness of PRP.



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Figure 7. PRP Inhibition of C6 Tumor Growth

Equal numbers of C6-V (filled circles and solid line), low expressing C6-PRP (open diamonds and small dashed line) or high expressing C6-PRP (open squares and large dashed line) cells were injected subcutaneously into nude mice. The slopes of the lines are 0.26 ± 0.04 for C6-V, 0.23 ± 0.04 for low expressing C6-PRP, and 0.14 ± 0.01 for high expressing C6-PRP (P < 0.05 vs. control). Tumor size (mean ± SEM with n = 4 mice per group) was measured until 3 weeks after injection. P < 0.05 for every individual time point comparing high expressing C6-PRP tumor size to C6-V control.

 


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Figure 8. PRP Inhibition of C6 Tumor Vascularization

Tumor tissue from parental C6 (A) and vector-transfected C6 cells (B) display numerous vessels. High-PRP-expressing tumors (C and D), but much less so low-PRP-expressing tumors (E and F), showed a marked diminution in vessel density. Tissue sections were counterstained with Harris’s hematoxylin. Bar = 100 µm.

 
Effect and Mechanism of Action of PRP on Human Endothelial Cells
We have previously demonstrated that PRP is able to inhibit the migration of bovine endothelial cells in culture and angiogenesis in the rat cornea (1). If the currently unidentified PRP receptor and downstream signaling pathways are to provide potential targets for the design of novel antitumor therapeutics, then it is also essential to determine whether this conservation of PRP action on endothelial cells from other species extends to human endothelial cells. We therefore examined the effect of PRP on the migration of human dermal microvascular endothelial cells in response to basic fibroblast growth factor (bFGF) in a Boyden chamber assay. Cells on a collagen-coated filter in upper wells of the chamber were exposed to medium alone or medium supplemented with bFGF, PRP, or the combination of bFGF and PRP in the lower chambers. Six hours later, the number of endothelial cells that had migrated from the upper to the lower surface of the filter was determined. Treatment with bFGF resulted in a 7-fold increase in migration (Fig. 9Go). PRP alone had no effect on cell migration, but this hormone did reverse the effect of bFGF. A monoclonal antibody against PRP (22), but not irrelevant antibodies, completely reversed the inhibitory effect of the PRP preparation, demonstrating that the antiangiogenic activity is fully attributable to PRP (Fig. 9Go). Thus, PRP is able to act directly on human endothelial cells to block an angiogenic response.



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Figure 9. PRP Inhibition of bFGF-Induced Human Endothelial Cell Migration

Total migration of human dermal microvascular endothelial cells in a Boyden chamber assay was measured in response to 10 ng/ml bFGF, 1 µg/ml PRP, 10 ng/ml bFGF + 1 µg/ml PRP, and the combination of these two factors in the presence of a 20 µg/ml monoclonal antibody (mAb) against PRP or an equivalent amount of irrelevant immunoglobulin (IgG). Data are the mean ± SEM for the number of cells migrated in 10 high-power microscope fields per well, with n = 6 wells; *, P < 0.01 relative to the untreated control.

 
PRP blocks endothelial cell migration in culture and neovascularization in vivo in response to both bFGF and PLF. Stimulation of endothelial cell migration by these angiogenic factors is inhibited by pertussis toxin, implicating G protein signaling (23, 24). At least for bFGF, the G-protein signaling pathway leads to arachidonic acid production as an essential step (23), suggesting that the arachidonic acid pathway may be an important target for PRP. Indeed, addition of 10 µM arachidonic acid reversed PRP inhibition of bFGF-stimulated human endothelial cell migration (Fig. 10Go).



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Figure 10. Arachidonic Acid Reversal of PRP Inhibition of bFGF-Induced Endothelial Cell Migration

Total migration of human dermal microvascular endothelial cells in a Boyden chamber assay was measured in response to 10 ng/ml bFGF, 1 µg/ml PRP, 1 or 10 µM arachidonic acid, or to combinations of these factors. Data are the mean ± SEM for the number of cells migrated in 10 high-power microscope fields per well, with n = 6 wells; *, P < 0.001 relative to the untreated control. Note that PRP blocks the response to bFGF (compare columns 2 and 6), and that 10 µM arachidonic acid reverses this effect of PRP (column 8).

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The experiments presented here demonstrate that PRP is able to restrict both tumor angiogenesis and tumor growth in mice. These results are consistent with the antiangiogenic effects of PRP being the cause of the reduced tumor growth, but we cannot exclude at this time other possible biological effects of PRP that may be relevant to tumor growth. PRP thus joins several other antiangiogenic factors, including angiostatin (25) and endostatin (26), as agents which can be used to inhibit tumor growth, at least in model systems. While it may be tempting to assume that any antiangiogenic factor will inhibit tumor growth, experimental evidence is required to establish the effectiveness of a factor for several reasons. An antiangiogenic factor may act on only some classes of endothelial cells, or only prevent angiogenesis induced by a limited set of angiogenic factors, or not be sufficiently robust in vivo because of posttranslational modification (such as proteolytic cleavage) or rapid clearance. The establishment of PRP as an effective agent in interfering with tumor angiogenesis in mice suggests that its receptor and downstream signaling pathways may be useful targets for antitumor drug design. The finding that the antiangiogenic effects of PRP can be overcome by addition of exogenous arachidonic acid represents the first clue to the PRP signaling mechanism. Predictions to be tested based on this result are that PRP inhibits the bFGF- or PLF-induced activation of phospholipase A2, and that a specific metabolic product of arachidonic acid (for example, a particular prostaglandin) may be an essential signal for endothelial cell migration in response to both bFGF and PLF.

The ability of PRP to act on tumors at sites outside of the uterus suggests that the high levels of PRP that circulate during pregnancy may have systemic effects. One possible contribution of PRP outside of the implantation site might be to restrict angiogenesis-dependent cell proliferation during pregnancy. The brief period of PRP expression in gestation (~1 week) makes it difficult to examine the effect of endogenous PRP on tumor growth. It should become possible, though, to identify both implantation site and distal effects of PRP by disruption of this gene in mice.

The tumor cells and hosts that we chose to examine provide a combination of homologous mouse components in the case of the SVT2 system, and a potent and widely used angiogenic tumor model in the C6 system. In both cases, by engineering tumor cells to secrete PRP, rather than injecting purified hormone, the tumor environment was exposed continuously to the antiangiogenic hormone and presumably to a reasonably constant concentration of hormone relative to tumor size. Exogenous administration of the hormone, or of an agonist ligand, to block tumor growth may pose greater challenges in terms of efficiently targeting the protein to the tumor site and preventing rapid clearance of the hormone from the circulation. Nevertheless, the inhibition of growth of the SVT2-PRP1 line as a tumor, a line that secretes only modest amounts of PRP compared with the other three SVT2-PRP cell lines that were tested, gives some reason to expect that it would be possible to deliver sufficient doses of this hormone or a hormone agonist to limit tumor growth.

In contrast to the intermediate effect seen with the SVT2-PRP1 cell line, only C6 cell lines expressing high levels of PRP were able to inhibit tumor growth. We were unable to extend the tumor growth studies to a point at which the effects of a lower dose of PRP might become evident, because the tumors had already reached the maximum size allowed by the animal care guidelines. The ability of a high dose, but not a lower dose, of PRP to inhibit C6 tumor growth is consistent with the idea that the net angiogenic activity, rather than a dominant effect of any single factor, drives tumor vascularization and growth. In the C6 model, very high levels of PRP are likely to be required to overcome the amount of angiogenic factor [vascular endothelial growth factor (VEGF)] produced by these tumor cells.

PRP can act directly on human microvascular endothelial cells to inhibit their response to bFGF, suggesting that the effects of PRP on tumor growth in mice may translate into similar effects of binding to the PRP receptor in humans. Although no evidence for a human PRP homolog has been obtained, the ability of PRP to act on human (and rat and bovine) endothelial cells indicates that the receptor and cell response are conserved among mammals. These results further argue that identifying placental hormones in model systems such as mice and rats can represent a useful approach to defining activities relevant to human physiology. Because the physiological changes during pregnancy are so dramatic, the characterization of these placental hormones is likely to reveal new mechanisms of cell regulation that are not evident in the nonpregnant animal. Furthermore, if PRP and other antiangiogenic factors are found to act on endothelial cells through distinct signaling pathways or to affect distinct stages of angiogenesis that contribute to tumor growth (27), then it is tempting to speculate that combinations of factors may provide an even more effective blockade of tumor angiogenesis than would be achieved by a single factor alone.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Cell Culture and Migration Assay
SV40 transformed BALB/c 3T3 cells (SVT2) (17), provided by Arnold Levine, and rat glioma C6 cells (18) were cultured in DMEM (Life Technologies, Inc., Gaithersburg, MD) supplemented with 10% calf serum and 2 mM L-glutamine in a humidified atmosphere of 5% CO2. SVT2 and C6 cells were transfected using calcium phosphate precipitation of either pMSXND (vector) or the pMSXND-PRP expression construct (5). Individual colonies were selected based on resistance to treatment with G418 (Sigma, St. Louis, MO) and then screened for PRP expression by immunoblotting. Clones expressing PRP were recloned by growth in soft agar, after which individual colonies were selected, passaged, and analyzed again for protein expression. For cell growth assays, SVT2 or C6 cells transfected with vector alone (SVT2-V and C6-V) or vector containing the PRP coding region (SVT2-PRP or C6-PRP) were plated at 7,000 cells per well in 12-well plates and grown under standard culture conditions; cell counts were performed with a hemocytometer on triplicate samples on a daily basis for 7 days.

Primary human dermal microvascular endothelial cells were obtained from Clonetics/BioWhittaker, Inc. (Walkersville, MD), and grown in EGM-MV medium (Clonetics/BioWhittaker, Inc.). To monitor migration, 50,000 endothelial cells were transferred per well of a Boyden chamber onto Nucleopore membranes with a pore size of 8 µm (Fisher Scientific, Pittsburgh, PA) and that had been coated with collagen type IV (Sigma). Medium lacking growth factors was added to the upper wells, and the identical medium with 10 ng/ml bFGF (R&D Systems, Minneapolis, MN), 1 µg/ml PRP, 1 or 10 µM arachidonic acid (Sigma), 20 µg/ml anti-PRP monoclonal antibody (22), or control mouse IgG (PharMingen, San Diego, CA), or various combinations of these factors was added to the lower wells. PRP was purified from the conditioned medium of Chinese hamster ovary cells stably transfected with a PRP expression construct, as described previously (5); monoclonal antibody was purified from hybridoma conditioned medium by Protein G affinity chromatography (Pharmacia Biotech, Piscataway, NJ). Six hours after treatment, filters were removed and stained with Dade Diff-Quick (Fisher Scientific) and then rinsed in PBS, and the number of cells that had migrated to the lower side of the filter were counted. Only cells in which the nucleus had moved through the membrane were counted.

Immunoblot Analysis
Conditioned media from stably transfected cell lines or from dispersed tumor cells that were placed into culture were collected and concentrated by Centricon-30 filtration (Millipore Corp., Bedford, MA). Fifteen micrograms of total protein were loaded per lane onto SDS-polyacrylamide gels. After electrophoresis, proteins were transferred to nitrocellulose membranes, which were then washed in 20 mM Tris-HCl, pH 7.6, 150 mM NaCl, 0.05% Triton X-100 (buffer A) containing 5% dried milk fat powder and then incubated for 2 h at room temperature with a 1:1,000 dilution of anti-PRP antiserum (5). After a 30-min wash in buffer A, membranes were exposed for 2 h at room temperature to a 1:1,000 dilution of horseradish peroxidase-conjugated, donkey-antirabbit immunoglobulin (Amersham Pharmacia Biotech, Arlington Heights, IL). After washing, bound antibody complexes were detected using the Super Signal chemiluminescence reagent (Pierce Chemical Co., Rockford, IL) and exposure to XAR film (Kodak, Rochester, NY).

In Vivo Tumor Analysis
Five-week-old female BALB/c mice were injected subcutaneously with 105 SVT2, SVT2-V, or SVT2-PRP cells resuspended in a volume of 100 µl of serum-free cell culture medium containing 0.5% methylcellulose (Fisher Scientific). Five-week-old nude mice were similarly injected with C6, C6-V, or C6-PRP cells. Three weeks after injection of SVT2 cells, tumors were excised and tumor weights and volumes were determined. For C6 cells, tumor growth was monitored throughout the 3-week period, with tumor volume calculated as length x width2 x 0.5. All procedures were approved by the Northwestern University Animal Care and Use Committee.

RNA Filter Hybridization
Ten micrograms of tumor RNA, isolated using Tri Reagent (Sigma), were loaded per lane of 2% agarose gels containing 6% formaldehyde. After electrophoresis, the RNA was visualized by ethidium bromide staining under UV light and then transferred to a nylon membrane. Membranes were hybridized overnight at 42 C with a 32P-labeled PRP cDNA probe and then washed and exposed to XAR film.

Immunohistochemistry
Excised tumors were fixed in 10% formalin and embedded in paraffin. Sections (15 µm) were permeabilized with 0.01% trypsin for 30 min at 37 C and washed in PBS. Endogenous peroxidase activity was quenched by incubation with 0.3% H2O2 in PBS for 15 min at 4 C followed by three PBS washes. Sections were then incubated with blocking buffer (2% rabbit serum and 5% BSA in PBS) for 30 min at room temperature, followed by 1 h at room temperature with rat-antimouse PECAM-1 monoclonal antibody (PharMingen) diluted 1:6 in blocking buffer. After washing in PBS, sections were exposed to biotinylated rabbit-antirat immunoglobulin secondary antibody (Vector Laboratories, Inc. Burlingame, CA) diluted 1:1,000 in blocking buffer for 1 h at room temperature, washed again, and treated for 1 h at room temperature with the ABC kit (Vector Laboratories, Inc.) followed by three PBS washes. Substrate solution of 0.06% (wt/vol) diaminobenzidine-0.03% (vol/vol) H2O2 in PBS was then added to the sections for 15 min at room temperature, and the sections were counterstained with Harris’s hematoxylin.

To count vessels, PECAM-1-positive cells and structures were totaled from complete cross-sections of several independent tumors for different tumor lines. Vessels were divided into either larger structures with obvious lumenal spaces, or a range of smaller PECAM-1-positive multicellular clusters, not all of which had evident lumenal spaces. With the inclusion of all PECAM-1-positive structures, numbers for "small vessels" may therefore underestimate the effects of PRP on vascular density.


    ACKNOWLEDGMENTS
 
We thank Diane Mayer and Janelle Roby for expert technical assistance and Noël Bouck for helpful comments.


    FOOTNOTES
 
Address requests for reprints to: Daniel I. H. Linzer, Department of Biochemistry, Molecular Biology, and Cell Biology, Northwestern University, Evanston, Illinois 60208. E-mail: dlinzer{at}northwestern.edu

This work was supported by NIH Grant R01 HD-24518, by the Robert H. Lurie Comprehensive Cancer Center (P30 CA-60553), and by the NIH Research Center on Fertility and Infertility at Northwestern University (P30 HD-28048).

Received for publication July 18, 2000. Revision received August 24, 2000. Accepted for publication September 7, 2000.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

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