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Clinical Cancer Research 13, 4051, July 15, 2007. doi: 10.1158/1078-0432.CCR-06-3032
© 2007 American Association for Cancer Research
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Human Cancer Biology

Intraperitoneal Injection of a Hairpin RNA–Expressing Plasmid Targeting Urokinase-Type Plasminogen Activator (uPA) Receptor and uPA Retards Angiogenesis and Inhibits Intracranial Tumor Growth in Nude Mice

Christopher S. Gondi1, Sajani S. Lakka1, Dzung H. Dinh2, William C. Olivero2, Meena Gujrati3 and Jasti S. Rao1,2

Authors' Affiliations: Departments of 1 Cancer Biology and Pharmacology, 2 Neurosurgery, and 3 Pathology, University of Illinois College of Medicine at Peoria, Peoria, Illinois

Requests for reprints: Jasti S. Rao, Department of Cancer Biology and Pharmacology, University of Illinois College of Medicine, One Illini Drive, Peoria, IL 61605. Phone: 309-671-3445; E-mail: jsrao{at}uic.edu.


   Abstract
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 Abstract
Materials and Methods
 Results
 Discussion
 References
 
Purpose: The purpose of this study was to evaluate the therapeutic potential of using plasmid-expressed RNA interference (RNAi) targeting urokinase-type plasminogen activator (uPA) receptor (uPAR) and uPA to treat human glioma.

Experimental Design: In the present study, we have used plasmid-based RNAi to simultaneously down-regulate the expression of uPAR and uPA in SNB19 glioma cell lines and epidermal growth factor receptor (EGFR)–overexpressing 4910 human glioma xenografts in vitro and in vivo, and evaluate the i.p. route for RNAi-expressing plasmid administered to target intracranial glioma.

Results: Plasmid-mediated RNAi targeting uPAR and uPA did not induce OAS1 expression as seen from reverse transcription-PCR analysis. In 4910 EGFR-overexpressing cells, down-regulation of uPAR and uPA induced the down-regulation of EGFR and vascular endothelial growth factor and inhibited angiogenesis in both in vitro and in vivo angiogenic assays. In addition, invasion and migration were inhibited as indicated by in vitro spheroid cell migration, Matrigel invasion, and spheroid invasion assays. We did not observe OAS1 expression in mice with preestablished intracranial tumors, which were given i.p. injections of plasmid-expressing small interfering RNA–targeting uPAR and uPA. Furthermore, the small interfering RNA plasmid targeting uPAR and uPA caused regression of preestablished intracranial tumors when compared with the control mice.

Conclusion: In conclusion, the plasmid-expressed RNAi targeting uPAR and uPA via the i.p. route has potential clinical applications for the treatment of glioma.

Nucleic acid–based methods have been used to target the transcriptional regulation of gene expression. One of these methods, RNA interference (RNAi), is now routinely used for the transient knockdown of gene expression in a wide range of organisms (1). Unlike antisense and triple-helix approaches, the RNAi-mediated method of gene regulation uses normal cellular responses to double-stranded RNA, which lead to the highly specific degradation of the target mRNA (1, 2). In addition and perhaps more importantly, this process triggers a cell-to-cell spreading of gene silencing as seen in several RNAi models (3–5). One potential obstacle for the therapeutic use of RNAi in mammalian cells is the activation of double-stranded RNA–dependent protein kinase (PKR) by long double-stranded RNA (6, 7). The ability to express intracellular small interfering RNA (siRNA) has been shown by several groups, mainly through the use of plasmids containing RNA pol III promoter (8–10).

Tumor cell invasion is a multistep process involving tumor cell attachment to the extracellular matrix followed by the degradation of host barriers by proteolysis and tumor colony formation at distinct sites (11). Urokinase plasminogen activator (uPA) plays a key role in tumor progression and invasion by virtue of its ability to initiate a cascade of proteases that can degrade most matrix and basement membrane components and interfere with cell-cell and cell-matrix interactions (12, 13). uPA and its high-affinity receptor, uPAR, which is a glycosyl phosphatidylinositol-anchored membrane protein (CD87), are believed to be critical elements in tumor biology because they control cell motility, tissue remodeling, and are involved in the bioavailability of angiogenic factors (14). Formation of the uPA-uPAR complex at the cell surface is required for efficient activation of plasmin, a protease that can degrade extracellular matrix components and release various growth factors (15, 16). Although many studies have documented that uPAR has a central role in uPA-mediated cell surface plasminogen activation, further studies with uPAR-deficient mice have shown the existence of additional pathways of uPA-mediated plasminogen activation that are independent of uPAR (17).

Glioblastomas are characterized by their invasive infiltration and destruction of surrounding normal brain tissue, making complete surgical resection of these tumors virtually impossible. Their invasive behavior seems to depend in part on a variety of proteolytic enzymes, including serine, metalloproteases, and cysteine proteases. Our previous work and that of others have suggested a direct correlation between the expression of uPA and uPAR and the invasiveness of human gliomas (18–23). Our studies have also shown that antisense clones for uPAR and uPA are unable to form tumors in nude mice intracerebrally (24–26). Further, adenoviral vectors carrying antisense uPAR inhibited the invasion of glioma cells in vitro and inhibition of ex vivo tumor formation (27). Taken together, these studies indicate the biological significance of uPA and uPAR in glioma invasion and tumor growth.

The delivery approach of adenovirus vectors and antisense technology to intracellularly target RNA seems to be a crucial limiting factor in exerting its inhibitory effect on the targeted molecule. The siRNA duplex is significantly more stable in cells than the cognate single-stranded sense or antisense RNA, with transcription, under the control of the identical promoter in each case (28). Here, we have produced a single construct driven by a cytomegalovirus (CMV) promoter to deliver hairpin RNA (hpRNA) molecules for both uPAR and uPA. We show the simplicity and ease of using vectors expressing hpRNA molecules for more than one target molecule using a single promoter and the subsequent, effective inhibition of glioma cell invasion, angiogenesis, and tumor growth both in vitro and in vivo.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
Construction of hpRNA-expressing plasmid. A pcDNA3 plasmid with a CMV promoter was used in the construction of the hpRNA-expressing vector. The uPA sequence agcttGagagccctgctggcgcgccatatataatggcgcgccagcagggctctca and uPAR sequence gatccTacagcagtggagagcgattatatataataatcgctctccactgctgtag were used for the siRNA sequence. Inverted repeat sequences were synthesized for both uPA and uPAR. The inverted repeats were laterally symmetrical making them self-complimentary with a 5-bp mismatch in the loop region. This 5-bp mismatch would aid in the loop formation of the hpRNA. Oligonucleotides were heated in a boiling water bath in 6x SSC for 5 min and self-annealed by slow cooling to room temperature. The resulting annealed oligonucleotides were ligated to pcDNA 3 at the HindIII site for uPA and BamHI site for uPAR sequentially and the plasmid was named pU2. Single constructs were also made: puPAR-targeting uPAR and puPA-targeting uPA. An inverted-repeat sequence targeting green fluorescent protein (GFP) mRNA was synthesized and cloned into the pcDNA 3 HindIII site as described above (Fig. 1 ).


Figure 1
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  		Fig. 1. Schematic representation showing the possible mechanism involved in the formation of siRNA molecules from hpRNA to induce RNAi targeting of uPAR and uPA.

 
Cell culture and transfection conditions. SNB19 (or SNB19 GFP) cell lines, established from a human high-grade glioma, and epidermal growth factor receptor (EGFR)–overexpressing human glioma xenograft tumors, designated as 4910 (xeno; kindly provided by Dr. David James at University of California at San Francisco) were used for this study. Cells were grown in DMEM/F12 medium (1:1, v/v) supplemented with 10% FCS in a humidified atmosphere containing 5% CO2 at 37°C. SNB19 and 4910 cells were transfected with plasmid constructs [empty vector (EV), scrambled vector (SV), puPAR, puPA, or pU2] using LipofectAMINE as per the manufacturer's instructions (Life Technologies).

PCR and reverse transcription-PCR. Total RNA was isolated from SNB19 and 4910 control cells and the cells were transfected with EV, SV, puPAR, puPA, or pU2. RNA was also isolated from cells transfected with antisense expression vectors for uPAR and uPA, and from cells transfected with a plasmid vector expressing siRNA-targeting GFP. Reverse transcription-PCR (RT-PCR) was done as per standard protocol for uPAR and uPA. To determine whether these siRNA-expressing plasmids induce an IFN response, RT-PCR for OAS1 was done. Cells were also treated with IFN{alpha} (0.5 ng/mL) to visualize OAS1 mRNA expression as positive control. Total RNA was isolated from fresh or paraffin-embedded brain tissue (Ambion) from control or mice injected i.p. with EV, SV, puPAR, puPA or pU2, or IFN{alpha} (0.5 ng). RT-PCR was done to determine OAS1 expression. PCR was done using primers specific for PCDNA3 vector amplifying CMV to BGH sequence regions using deparaffinized and protease-treated intracranial tumors. These primers had no significant homology to mouse or human sequences. Primers used for PCR and RT-PCR are listed in Table 1 .


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  		Table 1. Primers used for PCR and RT-PCR

 
Western blotting. SNB19 cells were transfected with mock, EV/SV, puPA, puPAR, or pU2. After 48 h, cells were collected and total cell lysates were prepared in extraction buffer containing Tris (0.1 mol/L; pH 7.5), Triton X-114 (1.0%), EDTA (10 mmol/L), aprotinin, and phenylmethylsulfonyl fluoride as described previously (27). The extracts were incubated at 37°C for 5 min and centrifuged to separate the lower (detergent) phase that contains mainly hydrophobic membrane proteins, including the glycosylphosphatidylinositol-anchored uPAR. Subsequently, 20 µg of protein from these samples were separated under nonreducing conditions by 12% SDS-PAGE and transferred to nitrocellulose membranes (Schleicher & Schuell). The membranes were probed for 2 h with antibodies against uPAR. The membranes were subsequently washed thrice with PBS to remove excess of primary antibodies, incubated with secondary antibodies as required, and then developed according to enhanced chemiluminescence protocol (Amersham). For loading control, the membranes were stripped and probed with monoclonal antibodies for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as per standard protocol. Similarly, 4910 EGFR-overexpressing xenograft cells were transfected with mock, EV/SV, puPA, puPAR, or pU2, and proteins were extracted. Western blot analysis was done for EGFR and vascular endothelial growth factor (VEGF) per standard protocol.

Fibrin zymography. The enzymatic activity and molecular weight of electrophoretically separated forms of uPA were determined in conditioned medium of SNB19 cells transfected with mock, EV/SV, puPA, puPAR, or pU2 by SDS-PAGE as described previously (24, 29). Briefly, the SDS-PAGE gel contains acrylamide to which purified plasminogen and fibrinogen were substrates before polymerization. After polymerization, equal amounts of proteins in the samples were electrophoresed and the gel was washed and stained as described previously (24, 29).

Immunocytochemical analysis. 4910 cells (1 x 104) were seeded on vitronectin-coated eight-well chamber slides, incubated for 24 h and transfected with mock, EV, puPAR, puPA, or pU2. After 72 h, cells were fixed with 3.7% formaldehyde and incubated with 1% bovine serum albumin (BSA) in PBS at room temperature for 1 h for blocking. After the slides were washed with PBS, either IgG anti-VEGF (mouse) or IgG anti-EGFR (mouse) was added at a concentration of 1:200. The slides were incubated at room temperature for 1 h and washed thrice with PBS to remove excess primary antibody. Cells were then incubated with anti-mouse FITC-conjugated IgG (1:500 dilution) for 1 h at room temperature. The slides were then washed thrice, covered with glass coverslips with 4',6-diamidino-2-phenylindole–containing mounting medium, and fluorescent photomicrographs were obtained.

In vitro angiogenic assay. 4910 and SNB19 cells (2 x 104 per well) were seeded in eight-well chamber slides and transfected with mock, EV, puPAR, puPA, or pU2. After a 24-h incubation period, the conditioned medium was removed and added to a 4 x 104 human dermal endothelial cell monolayer in eight-well chamber slides and the human dermal endothelial cells were allowed to grow for 72 h. Cells were then fixed in 3.7% formaldehyde, blocked with 2% BSA, and the endothelial cells were incubated with factor VIII primary antibody (DAKO Corp.). Cells were washed with PBS and incubated with a FITC-conjugated secondary antibody for 1 h. The slides were then washed and the formation of capillary-like structures was observed by fluorescent microscopy. Endothelial cells were also grown in conditioned medium of either 4910 or SNB19 cells transfected with mock, EV, puPAR, puPA, or pU2. Endothelial cells were allowed to grow for 72 h and H&E stained to visualize capillary network formation. The degree of angiogenesis was quantified based on the numerical value for the product of the number of branches and number of branch points as an average of 10 fields.

Dorsal skin-fold chamber model. Athymic nude mice (nu/nu; 18 male/female, 28-32 g) were bred and maintained within a specific-pathogen, germ-free environment. The implantation technique of the dorsal skin-fold chamber model has been described previously (30). Sterile small-animal surgical techniques were followed. Mice were anesthetized by i.p. injection with ketamine (50 mg/kg)/xylazine (10 mg/kg). Once the animal was anesthetized completely, a dorsal air sac was made in the mouse by injecting 10 mL of air. Diffusion chambers (Fisher) were prepared by aligning a 0.45-µm Millipore membranes (Fisher) on both sides of the rim of the "O" ring (Fisher) with sealant. Once the chambers were dry (2-3 min), they were sterilized by UV radiation for 20 min. Twenty microliters of PBS were used to wet the membranes. Glioblastoma cells (2 x 106; 4910 cells transfected with either EV or pU2), suspended in 100 to 150 µL of sterile PBS, were injected into the chamber through the opening of the O ring. The opening was sealed by a small amount of bone wax. A 1.5 to 2 cm superficial incision was made horizontally along the edge of the dorsal air sac and the air sac was opened. With the help of forceps, the chambers were placed underneath the skin and sutured carefully. After 10 days, the animals was anesthetized with ketamine/xylazine and sacrificed by intracardiac perfusion with saline (10 mL) followed by a 10 mL of 10% formalin/0.1 mol/L phosphate solution and 0.001% FITC solution in PBS. The animals were carefully skinned around the implanted chambers and the implanted chambers were removed from the sc. air fascia. The skin fold covering the chambers was photographed under visible light and for FITC fluorescence. The number of blood vessels within the chamber in the area of the air sac fascia was counted and their lengths were measured.

Spheroid migration assay. Migration was assayed as described previously (27) with some modifications. Spheroids of SNB19 and 4910 cells were prepared by seeding a suspension of 2 x 106 cells in DMEM on ultra low attachment 100 mm tissue culture plates and cultured until spheroid aggregates formed. Spheroids measuring ~150 µm in diameter (~4 x 104 cells per spheroid) were selected and transfected with pU2. Three days after infection, single glioma spheroids were placed in the center of each well of a vitronectin-coated (50 µg/mL), 96-well microplate, and 200 µL of serum-free medium was added to each well. Spheroids were incubated at 37°C for 24 h, after which the spheroids were fixed and stained with Hema-3 and photographed. The migration of cells from spheroids to monolayers was quantified using a microscope calibrated with a stage and ocular micrometer and represented graphically.

Matrigel invasion assay. Invasion of glioma cells in vitro was measured by the invasion of cells through Matrigel-coated (Collaborative Research, Inc.) transwell inserts (Costar). Briefly, transwell inserts with 8-µm pores were coated with a final concentration of 1 mg/mL of Matrigel; SNB19 and 4910 cells transfected with mock; EV/SV, puPAR, puPA, or pU2 were trypsinized; and 200 µL aliquots of cell suspension (1 x 106/mL) were added in triplicate wells. After a 24-h incubation period, cells that passed through the filter into the lower wells were quantified as described earlier (27, 31) and expressed as a percentage of the sum of cells in the upper and lower wells. Cells on the lower side of the membrane were fixed, stained with Hema-3, and quantified as percentage invasion.

In vitro spheroid invasion assay. Multicellular SNB19 spheroids were cultured in six-well ultra low attachment plates. Briefly, 3 x 106 cells were suspended in 10 mL of medium, seeded onto the plates, and cultured until spheroids formed. Spheroids, 100 to 200 µm in diameter, were selected and transfected with mock, EV/SV, puPAR, puPA, or pU2. Three days after infection, tumor spheroids were stained with the fluorescent dye DiI and confronted with fetal rat brain aggregates stained with DiO. The progressive destruction of fetal rat brain aggregates and invasion of SNB19 cells were observed by confocal laser scanning microscopy and photographed as described previously (32, 33). The remaining volume of the rat brain aggregates at 24, 48, and 72 h were quantified using image analysis software as described previously (32) and graphically represented (33).

Intracranial tumor growth inhibition. For the intracerebral tumor model, 2 x 106 4910 xenograft tumor cells were intracerebrally injected into nude mice. Ten days after tumor implantation, the mice were treated with i.p. injections of pU2 (150 µg/injection/mouse) every other day three times. Control mice were either injected with PBS alone or with empty plasmid, vector (150 µg/injection/mouse). Five weeks after tumor inoculation, six mice from each group were sacrificed by cardiac perfusion with 3.5% formaldehyde in PBS, their brains were removed, and paraffin sections were prepared. Sections were stained with H&E to visualize tumor cells and to examine tumor volume (32, 33). The sections were blindly reviewed and scored semiquantitatively for tumor size. Whole-mount images of brains were also taken to determine infiltrative tumor morphology. The average tumor area per section integrated to the number of sections where the tumor was visible was used to calculate tumor volume and compared between controls and treated groups. RT-PCR was done on fresh or paraffin-embedded brain tissue for OAS1 pcDNA3 plasmid and GAPDH as previously described.

Immunohistochemical analysis. Brains of control and pU2 i.p. treated mice implanted with 4910 tumors were fixed in formaldehyde and embedded in paraffin as per standard protocol. Sections were deparaffinized as per standard protocol. Sections were blocked in 1% BSA in PBS for 1 h, and the sections were subsequently transferred to primary antibody (uPAR and uPA) diluted in 1% BSA in PBS (1:500). Sections were allowed to incubate in the primary antibody solution for 2 h at 4°C in a humidified chamber, followed by a wash in 1% BSA in PBS, and placed in a solution with the appropriate (anti-mouse and anti-rabbit FITC) secondary antibody. The sections were allowed to incubate with the secondary antibody for 1 h and visualized using a confocal microscope. Images were obtained for FITC. Transmitted light images were also obtained after H&E staining as per standard protocol to visualize the morphology of the sections. A control study was done using a normal rabbit immunoglobulin fraction as the primary antibody (control antibody) instead of uPAR and uPA.

In situ hybridization. Xenograft tumor cells (2 x 106) were intracerebrally injected into nude mice. Ten days after tumor implantation, the mice were treated with i.p. injections of mock, EV/SV, puPAR, puPA, or pU2 (150 µg/injection/mouse) every other day three times. Control mice were either injected with PBS alone or with empty plasmid vector (150 µg/injection/mouse). Five weeks after tumor inoculation, six mice from each group were killed by cardiac perfusion with 3.5% formaldehyde in PBS, their brains were harvested, and paraffin sections were prepared. Sections were deparaffinized and probed for PCDNA3-CMV promoter using specific alkaline phosphatase labeled (Alkaphos, Amersham) DNA oligonucleotide (CTGGTGTCGACCTGCTTCCGCGATGTACGGGC) as per standard protocol. Hybridization was observed using nitroblue tetrazolium alkaline phosphatase substrate Western Blue (Promega) as per manufacturer's instructions.

Animal survival analysis. Nude mice were implanted with intracranial 4910 xenograft tumors and their survival ability was determined based on symptoms of intracranial pressure, arched back, and dehydration. If the animals exhibited excessive pain, they were euthanized. Two sets of animals were used (six mice per group). Both sets were implanted with intracranial xenograft tumors as described previously. Ten days after tumor implantation, the mice were treated with i.p. injections of pU2 (150 µg/injection/mouse) every other day three times. Control mice were either injected with PBS alone or EV/SV (150 µg/injection/mouse). The mice were maintained in clean room conditions and monitored every day for 112 days after which the experiment was artificially terminated. Brains were collected from the control and treated mice, paraffin-embedded, sectioned, and H&E stained per standard protocols. The survival curve was plotted as per standard methods and graphically represented as percentage survival.


   Results
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 Abstract
 Materials and Methods
 Results
Discussion
 References
 
Effect of pU2 plasmid on uPA and uPAR mRNA levels in total cell extracts. Total RNA was isolated from control cells and cells transfected with EV, SV, puPAR, puPA, and pU2. RNA was also isolated from cells transfected with antisense expression vectors for uPAR and uPA, and from cells transfected with plasmid vector expressing siRNA-targeting GFP. RT-PCR was done as per standard protocol for uPAR and uPA. To determine whether these siRNA-expressing plasmids induce an IFN response, RT-PCR for OAS1 was done. As a positive control, cells were treated with IFN{alpha} (0.5 ng/mL) to visualize OAS1 mRNA expression. Figure 2A shows RT-PCR analysis of total RNA isolated from cells treated with IFN{alpha}, control, antisense uPAR (asuPAR), antisense uPA (asuPA), and siRNA expressing vectors for uPAR (puPAR), uPA (puPA), and both upAR and uPA (pU2). As RNAi controls, siRNA targeting GFP was used in non-GFP cells, EV (pEV), and SV (pSV). RT-PCR for GAPDH served as an internal control. IFN-treated cells did not show a change in the expression levels of uPA or uPAR; OAS1 induction was observed and served as control for OAS1 RT-PCR. Control, pGFP, pEV, and pSV-treated cells did not show changes in uPAR or uPA expression and no detectable levels of OAS1 were observed. Antisense for uPAR and uPA did not show a change in the mRNA levels of uPAR or uPA, whereas the protein levels showed a 20% to 50% decrease (not shown). Cells treated with puPAR showed a 20% decrease in uPA expression and a 50% decrease in uPAR expression. Cells treated with puPA did not show appreciable change in uPAR expression levels, whereas uPA levels showed a 30% to 60% decrease. Cells treated with pU2 showed a 75% decrease in uPAR mRNA levels and 60% decrease in uPA mRNA levels. GAPDH levels did not change and served as controls.


Figure 2
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  		Fig. 2. Determination of uPAR and uPA mRNA levels using semiquantitative RT-PCR. Total RNA was isolated from control cells and cells transfected with EV, SV, puPAR, puPA, or pU2. RNA was also isolated from cells transfected with antisense expression vectors for uPAR and uPA, and from cells transfected with a plasmid vector expressing siRNA for GFP. RT-PCR was done per standard protocol for uPAR and uPA. To determine whether these siRNA-expressing plasmids induce an IFN response, RT-PCR for OAS1 was done. As a positive control, cells were also treated with IFN{alpha} (0.5 ng/mL) to visualize OAS1 mRNA expression (A). SNB19 cells were transfected with mock, EV/SV, puPA, puPAR, or pU2. After 48 h, cells were collected and total cell lysates were prepared in extraction buffer containing Tris (0.1 mol/L; pH 7.5), Triton X-114 (1.0%), EDTA (10 mmol/L), aprotinin, and phenylmethylsulfonyl fluoride as described previously. Subsequently, 20 µg of protein from these samples were separated under nonreducing conditions by 12% SDS-PAGE and transferred to nitrocellulose membranes (Schleicher & Schuell). The membranes were probed for 2 h with antibodies against uPAR (B). The membranes were subsequently washed thrice with PBS to remove excess primary antibody, incubated with a secondary antibody as required, and then developed per standard protocol. For loading control, the membranes were stripped and probed with monoclonal antibodies for GAPDH. The enzymatic activity and molecular weight of electrophoretically separated forms of uPA were determined from the conditioned medium of SNB19 cells transfected with mock, EV/SV, puPA, puPAR, or pU2 by SDS-PAGE as described previously (B). Western blot analysis was also done using cell lysates of 4910 EGFR-overexpressing 4910 xenograft cells transfected with mock, EV/SV, puPA, puPAR, or pU2. Western blots were immunoprobed for EGFR and VEGF per standard protocols (C).

 
Effect of pU2 on uPA and uPAR enzymatic activity in SNB19 cells and EGFR and VEGF protein levels in 4910 xenograft cells. RNAi targeted against proteolytic degradation could be an important intervention to prevent cancer cell invasion. uPA and uPAR have been shown to play significant roles in extracellular matrix degradation. As shown by Western blotting, transfection of SNB19 cells with a vector expressing siRNA for uPAR and uPA (pU2) strongly inhibited the protein expression of uPAR levels when compared with mock and EV/SV–transfected cells (Fig. 2B). Fibrin zymography revealed that uPA enzymatic activity significantly decreased in SNB19 cells transfected with puPAR, puPA, and pU2 compared with mock and EV/SV transfection (Fig. 2B). GAPDH protein levels served as a loading control (Fig. 2B). Quantitative analysis of uPAR and uPA bands by densitometry revealed a significant (P < 0.001) decrease in uPAR protein (13- to 16-fold) and uPA enzymatic activity (10- to 12-fold) in pU2-transfected cells compared with mock and EV/SV–transfected cells. Cells transfected with puPAR and puPA inhibited uPAR and uPA levels in almost the same manner as pU2, but down-regulation of the target molecules was much more pronounced with the bicistronic construct compared with either of the single constructs. 4910 xenograft cells exhibited similar down-regulation of uPA and uPAR as SNB19 cells (data not shown). Because EGFR levels are overexpressed in 4910 cells, we determined the effect of pU2 transfection. Simultaneous down-regulation of uPAR and uPA decreased EGFR levels in 4910 cells at least 15-fold, whereas VEGF levels decreased 5-fold (Fig. 2C). GAPDH levels did not change and served as a loading control. In situ levels of VEGF and EGFR were also determined and those results correlated with Western blot analysis (Fig. 3A ).


Figure 3
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  		Fig. 3. To visualize VEGF and EGFR expression in EGFR-overexpressing 4910 cells, 1 x 104 cells were seeded on vitronectin-coated eight-well chamber slides, incubated for 24 h, and transfected with mock, EV, and a vector-expressing siRNA for uPAR (puPAR), uPA (puPA), or both (pU2). After 72 h, cells were fixed with 3.7% formaldehyde and incubated with 1% BSA albumin in PBS at room temperature for 1 h for blocking. After the slides were washed with PBS, either IgG anti-VEGF (mouse) or IgG anti-EGFR (mouse) was added at a concentration of 1:200. The slides were incubated at room temperature for 1 h and washed thrice with PBS to remove excess primary antibody. Cells were then incubated with anti-mouse FITC conjugated IgG (1:500 dilution) for 1 h at room temperature. The slides were washed thrice, covered with glass coverslips with 4',6-diamidino-2-phenylindole–containing mounting medium, and fluorescent photomicrographs were obtained. A, expression of EGFR and VEGF in control and EV/SV–, puPAR-, puPA-, and pU2-transfected 4910 cells. To determine in vitro angiogenesis, 4910 or SNB19 cells (2 x 104 per well) were seeded in eight-well chamber slides and transfected with mock, EV, and a vector-expressing siRNA for uPAR (puPAR), uPA (puPA), or both (pU2). After a 24-h incubation period, the conditioned medium was removed and added to 4 x 104 human dermal endothelial cell monolayer in eight-well chamber slides. The human dermal endothelial cells were allowed to grow for 72 h. Cells were then fixed in 3.7% formaldehyde, blocked with 2% BSA, and incubated with factor VIII primary antibody (DAKO Corp.). The cells were then washed with PBS and incubated with a FITC-conjugated secondary antibody for 1 h. The slides were washed and the formation of capillary-like structures was observed using fluorescent microscopy (B). Endothelial cells were also grown in conditioned medium of 4910 or SNB19 cells transfected with mock-, EV-, and a vector-expressing siRNA for uPAR (puPAR), uPA (puPA), or both (pU2). Endothelial cells were allowed to grow for 72 h, and H&E stained for visualization of network formation (B). As determined by in vitro angiogenesis quantification, similar results were obtained for SNB19 and 4910 cells. The degree of angiogenic induction was quantified for both SNB19 and 4910 cells based on the numerical value for the product of the number of branches and number of branch points (*, P = 0.005; C). D, in vivo angiogenic assay using the dorsal skin fold model as described in Materials and Methods. Briefly, the animals were implanted with diffusion chambers containing control or pU2-transfected 4910 cells in a dorsal cavity. Ten days after implantation, the animals were sacrificed and vasculature flushed with FITC solution. The skin fold covering the diffusion chamber was observed for FITC fluorescence and in visible light for the presence of tumor-induced neovasculature (TN) and preexisting vasculature (PV).

 
Inhibition of tumor cell–induced capillary network formation by pU2. Emerging tumors are dependent on the formation of new blood vessels to fuel tumor growth. Because uPA and uPAR have been reported to regulate angiogenesis, we next assessed the effect of pU2 on tumor cell-induced angiogenesis. We did immunohistochemical analysis using factor VIII antigen to evaluate tumor-induced vessel formation in an in vitro coculture system and H&E staining of endothelial cells cultured in conditioned medium of SNB19 and 4910 xenograft cells after transfection with mock, EV/SV, puPA, puPAR, or pU2. The results showed that endothelial cells formed capillary-like structures in the presence of mock and EV/SV–transfected cells within 48 h. In contrast, the pU2 vector significantly inhibited tumor cell-induced capillary-like network formation. SNB19 and 4910 xenograft cells exhibited similar behavior although 4910 xenograft cells were slightly more aggressive in terms of angiogenesis when compared with SNB19 cells (Fig. 3B). The quantitation of the branch points and number of branches was extremely low in pU2-transfected cocultures compared with mock and EV/SV–transfected cells. In the case of xenograft cocultures, rudimentary network-like structures were seen in the pU2 treatment group (Fig. 3B). The effect was <50% to 60% in puPA- and puPAR-transfected cocultures, respectively, when compared with mock and EV/SV–transfected cocultures in relation to capillary-like structure formation (Fig. 3C). To confirm the in vitro coculture experiments, we examined whether pU2 could inhibit tumor angiogenesis in vivo as assessed by the dorsal window model. Implantation of a chamber containing mock- and EV-transfected 4910 xenograft cells resulted in microvessel development with thin, curved structures and numerous tiny bleeding spots [as indicated by arrows (tumor-induced neovasculature and preexisting vasculature)]. In contrast, implantation of 4910 xenograft cells transfected with pU2 did not result in the development of any additional microvessels (Fig. 3D). Similar results were obtained with SNB19 cells (data not shown).

siRNA against uPA and uPAR inhibits glioma migration and invasion. To determine whether uPAR and uPA siRNA expression is capable of influencing tumor cell migration, we transfected SNB19 and 4910 xenograft spheroids with the puPAR, puPA, and pU2. As shown in Fig. 4A , there was much higher cell migration in the control and EV/SV–transfected spheroids than in the pU2-transfected spheroids. As shown by the quantitation of the distance of migration out from the spheroids, cell migration of the control and EV/SV–transfected spheroids was significantly higher (P < 0.05) compared with pU2-transfected spheroids. Moderate cell migration from tumor spheroids transfected with puPAR and puPA was observed. Further, 4910 xenograft cells seemed to be more migratory than SNB19 cells. The effect of pU2 was similar in both SNB19 and 4910 xenograft cells (Fig. 4A).


Figure 4
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  		Fig. 4. Migration was assayed as previously described. Spheroids of SNB19 or 4910 cells were prepared by seeding a suspension of 2 x 106 cells in DMEM on ultra–low attachment 100-mm tissue culture plates and cultured until spheroid aggregates formed. Spheroids measuring ~150 µm in diameter (~4 x 104 cells per spheroid) were selected and transfected with mock-, EV-, and a vector-expressing siRNA for uPAR (puPAR), uPA (puPA), or both (pU2). Three days after infection, single glioma spheroids were placed in the center of each well in vitronectin-coated (50 µg/mL) 96-well microplates and 200 µL of serum-free medium was added to each well. Spheroids were incubated at 37°C for 24 h, after which the spheroids were fixed and stained with Hema-3 and photographed. Cell migration from spheroids to monolayers was quantified using a microscope calibrated with a stage and ocular micrometer and represented graphically (A). In vitro invasion of SNB19 and 4910 cells was determined by measuring the cells that invaded through Matrigel-coated (Collaborative Research) transwell inserts (Costar). Briefly, transwell inserts with 8-µm pores were coated with a final concentration of 1 mg/mL of Matrigel; SNB19 cells transfected with mock, EV/SV, puPAR, puPA, or pU2 were trypsinized; and 200 µL aliquots of cell suspension (1 x 106 cells/mL) were added to the wells in triplicate. After a 24-h incubation period, cells that passed through the filter into the lower wells were quantified as described earlier and expressed as a percentage of cells in the lower wells. Cells on the lower side of the membrane were fixed, stained with Hema-3, and quantified as percentage invasion (B). SNB19 and 4910 spheroids were cultured in six-well ultra–low attachment plates. Briefly, 3 x 106 cells were suspended in 10 mL of medium, seeded onto the plates, and cultured until spheroids formed. Spheroids, 100 to 200 µm in diameter, were selected and transfected with mock, EV/SV, puPAR, puPA, or pU2. Three days after infection, tumor spheroids were stained with the fluorescent dye DiI and confronted with fetal rat brain aggregates stained with DiO. The progressive destruction of fetal rat brain aggregates and invasion of SNB19 cells were observed by confocal laser scanning microscopy and photographed as described previously (32, 33). The remaining volume of the rat brain aggregates at 24, 48, and 72 h were quantified using image analysis software as described previously and graphically represented (C). S, SNB19; X, 4910 xeno.

 
To evaluate the effect of siRNA-mediated inhibition of uPAR and uPA on glioma invasiveness, we used a two-model system: the standard Matrigel invasion assay and the spheroid invasion assay. SNB19 or 4910 xenograft cells transfected with mock and EV/SV extensively invaded the Matrigel-coated transwell insert. In contrast, the pU2-transfected cultures had markedly much less invasiveness through the Matrigel compared with control and EV/SV–transfected cells. Quantitative determination of invasion confirmed that pU2 transfection showed only 6% invasion in SNB19 cells and 8% invasion in 4910 xenograft cells compared with mock and EV/SV–transfected cells (Fig. 4B). Further, quantitative analysis results of invasion of SNB19 and 4910 xenograft cells transfected with puPAR and puPA was 22% (SNB19), 30% (4910), 35% (SNB19), and 40% (4910), respectively (Fig. 4B).

We further examined the extent of the pU2 effect in a spheroid invasion assay. The EV/SV–transfected spheroids progressively invaded fetal rat brain aggregates, and quantitation revealed that the glioma spheroids invaded the fetal rat brain aggregates by 25% within 24 h, >70% within 48 h, and >90% at 72 h. In contrast, the tumor spheroids transfected with the pU2 vector invaded the fetal rat brain aggregates by only 2% (SNB19) and 2.8% (4910 xeno) after 72 h. Invasion of fetal rat brain aggregates by the single constructs were as follows: 20% (SNB19) and 23% (4910 xeno) with puPAR, and 40% (SNB19) and 60% (4910 xeno) with puPA (Fig. 4C). Taken together, these findings provide strong evidence that RNAi-mediated silencing of uPAR and uPA greatly inhibits glioma cell invasion in both in vitro models compared with the single siRNA constructs for uPAR and uPA.

uPA and uPAR siRNA suppresses intracranial tumor growth. We used an intracranial tumor model with 4910 xenograft cells to assess the effects of RNAi-mediated inhibition on preestablished tumor growth in vivo. Paraffin brain sections of the untreated (mock) and EV/SV–treated control groups were characterized by large-spread tumor growth after a 5-week follow-up period as visualized by H&E staining of similar sections (Fig. 5A ). However, we could not detect tumors in mice treated with the pU2 vector (Fig. 5A). Further quantification of H&E-stained brain sections by a neuropathologist (Dr. Gujrati) blinded to treatment condition revealed no difference in tumor size between the control and EV/SV treatment groups. However, total regression of preestablished tumors was revealed in the pU2-treated group (Fig. 5A and B). Preestablished intracranial tumor growth was inhibited by 70% in puPAR-treated mice and 55% in puPA-treated mice. These results showed that RNAi-mediated suppression of uPAR and uPA inhibited preestablished intracranial tumor growth. Paraffin sections were probed for uPA and uPAR protein as previously described (see Materials and Methods). pU2-treated mice did not exhibit uPAR or uPA protein expression above expected background levels (not shown), whereas the control and EV/SV–treated mice showed increased levels of uPAR and uPA expression localized at the tumor region (Fig. 5C). To determine whether this down-regulation of protein expression was caused by pU2 treatment, paraffin brain sections of 4910 cells implanted in untreated mice (mock) and mice treated with SV/EV, puPAR, puPA, and pU2 were probed with an alkaline phosphatase–labeled oligonucleotide for the CMV promoter region of the pcDNA3 plasmid vector. In the mock sections, no alkaline phosphatase activity was detected. In contrast, we observed alkaline phosphatase activity in the EV/SV–, puPAR-, puPA-, and pU2-treated mice, indicating the presence of a pcDNA3 vector in the tissue (Fig. 5D). As a positive control, mice were injected intracranially with IFN{alpha} to determine OAS1 expression. Total RNA from fresh or paraffin-embedded brain sections was used for RT-PCR to determine in vivo induction of IFN response plasmids inducing RNAi (see Materials and Methods). From the RT-PCR analysis, it is clear that no OAS1 induction was observed in the mock, EV/SV–, puPAR-, puPA-, or pU2-treated mice. RT-PCR for GAPDH served as an internal control (Fig. 5E). The presence of a pcDNA3 vector was confirmed by regular PCR analysis (Fig. 5E). To determine the survival rate of mice implanted with 4910 xenograft intracranial tumors, we did a long-term survival study, Thirty days after implantation, all of the control mice died and paraffin sections of these brains revealed the presence of large intracranial tumors (Fig. 5A). In contrast, pU2-treated mice survived much longer. One mouse from the pU2-treated group died after 43 days, but paraffin sections revealed the presence of a hemorrhage and very few xenograft cells, indicating that the death of this animal was not due to an intracranial tumor (not shown). The remaining five animals in the pU2-treated group were sacrificed after 112 days. Paraffin sections of these mice revealed no intracranial tumors, normal brain morphology, and no visible 4910-xenograft tumor cells. The survival rate is represented as percentage survival in Fig. 5F.


Figure 5
Figure 5
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  		Fig. 5. For the intracerebral tumor model, 2 x 106 4910 xenograft tumor cells were intracerebrally injected into nude mice. Ten days after tumor implantation, the mice were treated with i.p. injections of pU2 (150 µg/injection/mouse) every other day three times. Control mice were either injected with PBS alone or with an empty plasmid vector (150 µg/injection/mouse). Five weeks after tumor inoculation, six mice from each group were sacrificed via cardiac perfusion with 3.5% formaldehyde in PBS, their brains were removed, and paraffin sections were prepared. Sections were stained with H&E to visualize tumor cells and to examine tumor volume as described in Materials and Methods (arrows, approximate site of intracranial implantation site; A and B). To visualize the expression levels of uPAR and uPA in intracranial tumors, mouse brains were fixed in formaldehyde and embedded in paraffin per standard protocols. Sections were deparaffinized, blocked in 1% BSA in PBS for 1 h, and subsequently transferred to primary antibody (uPAR and uPA) diluted in 1% BSA in PBS (1:500). Sections were allowed to incubate in the primary antibody solution for 2 h at 4°C in a humidified chamber, followed by a wash in 1% BSA in PBS, and placed in a solution with the appropriate (anti-mouse and anti-rabbit FITC) secondary antibody. The sections were allowed to incubate with the secondary antibody for 1 h and visualized using a confocal microscope. Images were obtained for FITC. Transmitted light images were also obtained after H&E staining to visualize the morphology of the sections. A control study was done using a normal rabbit immunoglobulin fraction as the primary antibody (control antibody) instead of uPAR or uPA (C). In situ hybridization was done to determine the presence of transfected plasmid intracranially after i.p. injections. Briefly, 10 d after tumor implantation, the mice were treated with i.p. injections of control, EV/SV, puPAR, puPA, or pU2 (150 µg/injection/mouse) every other day three times. Control mice were injected with PBS alone. Five weeks after tumor inoculation, six mice from each group were sacrificed and brains were processed as described in Materials and Methods. Sections were deparaffinized and probed for pcDNA3-CMV promoter using specific alkaline phosphatase–labeled DNA oligonucleotides (CTGGTGTCGACCTGCTTCCGCGATGTACGGGC) per standard protocols. The presence of CMV promoter was determined by the development of a blue precipitate of nitroblue tetrazolium alkaline phosphatase substrate. Arrows, region of localization. D, the presence of plasmids intracranially was also determined by PCR amplification of CMV to BGH construct region of the plasmid using deparaffinized intracranial sections of control, EV/SV–, puPAR-, puPA-, or pU2 i.p. injected mice. To determine if IFN induction was present intracranially, total RNA was isolated from fresh or paraffin-embedded brain tissue from mice injected with control, EV/SV, puPAR, puPA, pU2, or IFN (0.5 ng) intracranially, and RT-PCR was done using primers specific for OAS1 (E). Nude mice were implanted with intracranial xenograft tumors and their survival ability was determined. Two sets of animals were used (six mice per group). Both sets of mice were implanted with intracranial xenograft tumors as described previously. Ten days after tumor implantation, the mice were treated with i.p. injections of pU2 plasmid (150 µg/injection/mouse) three times every other day. Control mice were either injected with PBS alone or with empty plasmid vector (150 µg/injection/mouse). The mice were maintained in clean room conditions and monitored every day for 112 d after which the experiment was artificially terminated. Brains were harvested, paraffin embedded, sectioned, and H&E stained as per standard protocols. Survival curve was plotted per standard methods and graphically represented (F).

 

   Discussion
Top
 Abstract
 Materials and Methods
 Results
 Discussion
References
 
In the present study, we provide evidence to suggest that the direct i.p. injections of plasmids expressing siRNA targeting uPAR and uPA inhibit intracranial tumor growth in nude mice. Extracellular matrix destruction is dependent on the expression of proteases, which are overexpressed in gliomas (34–37). Using an adenovirus-mediated antisense construct for uPAR and uPA, we have already shown that the simultaneous down-regulation of uPAR and uPA has a synergistic effect rather than an additive one (38). However, using an adenovirus to deliver therapeutic genes presents significant problems, including accurate targeting and toxicity (39). In recent years, several groups have experimented with various modifications, but thus far, none of these altered methods have been successful in eliminating the aforementioned problems (40). Here, we show that a simple CMV plasmid vector driving the production of hairpin-like RNA molecules can be used therapeutically. As indicated by the RT-PCR results, the use of an mRNA-like molecule possessing a polyadenylic acid tail and having 21-bp inverted repeats, which target uPAR and uPA, did not induce an IFN-like response. Other groups have used lentiviral vectors to successfully target specific proteases, such as plasminogen activator inhibitor-1, but still induced OAS1 (41). Here, we did not observe the induction of OAS1. This might have been due to the presence of a polyadenylic acid tail mimicking cellular mRNA and appearing as "self" to the cell. As expected, the down-regulation of the target mRNA molecule was observed with siRNA, whereas no down-regulation was observed with the use of an antisense sequence. With an antisense approach, equimolar quantities of the antisense molecule are required to silence the target gene. This is not required with RNAi where the RISC behaves like a catalyst and is reused (42). In essence, a small amount of RNAi-inducing molecules, such as siRNA or hpRNA, is sufficient to induce silencing of target genes. Our results show that future therapeutic use of RNAi to treat gliomas is very probable. Urokinase plasminogen activator and its receptor play important roles in invasion and migration of metastatic gliomas (35–37). Using these proteases as targets is very attractive for the treatment of gliomas.

In terms of angiogenesis, the use of antiangiogenic drugs has been met with mixed success (43). The targeting of uPAR and uPA, which are also indirectly involved in angiogenic pathways (44, 45), has revealed the importance of their targeting in cancer therapy. Our results show that the simultaneous down-regulation of uPAR and uPA causes the down-regulation of EGFR and VEGF in EGFR-overexpressing glioma xenograft cells (4910). In situ studies confirm Western blot analysis results in also demonstrating the down-regulation of EGFR and VEGF. In situ angiogenic assays have shown that endothelial cells cocultured with EGFR-overexpressing 4910 cells induce the endothelial cells to form a network-like pattern mimicking tumor angiogenesis. Progressive reduction in the network formation was seen in puPAR- and puPA-transfected cells. In cells transfected with pU2, we observed complete regression of network formation, indicating that the simultaneous down-regulation of uPAR and uPA causes the tumor cells to retard or stop secreting factors necessary for the induction of angiogenesis. The dorsal skin fold assay, an in vivo angiogenic assay, revealed complete inhibition of angiogenesis by pU2-transfected 4910 cells. Although our previous work using an adenovirus-mediated strategy had similar results (38), the major difference is that 100 multiplicity of infection of virus particles were required to achieve the same effect as 6 µg of plasmid. We also observed similar results with the other assays. For example, spheroid migration was significantly inhibited in pU2-treated SNB19 and 4910 xenograft cells. Our invasion studies showed that after transfection with pU2, both SNB19 and 4910 cells exhibited a significant reduction in their invasive ability—only 5% to 8% invasion when compared with the controls. From these spheroid invasion assay results, it is clear that the simultaneous down-regulation of uPAR and uPA retards the invasion of fetal rat brain aggregates.

uPAR is associated with integrins and is known to mediate cellular motility via extracellular matrix components such as vitronectin (46). In addition, association of uPAR with integrins mediates ras signaling pathways (47, 48). uPAR and uPA have also been observed at the leading edge of invading tumors (48). Hence, uPAR is a logical target to inhibit tumor invasion and migration. Our animal studies show that the simultaneous down-regulation of uPAR and uPA causes the regression of intracranial tumors. Nude mice implanted with 4910 xenograft cells intracranially usually die in 4 weeks due to tumor invasion. In contrast, mice injected with pU2 i.p. do not exhibit tumor establishment and survived for over 112 days after implantation. It would be of academic interest to see if similar results can be obtained using antisense vector constructs. Nevertheless, as the in situ hybridization studies indicated, there was translocation of the i.p. injected plasmid to the brain. The plasmids may pass through the blood-brain barrier probably due to the already compromised blood-brain barrier at the tumor site. The presence of the plasmid intracranially in control was seen primarily surrounding vessels (not shown). As such, the potential for using siRNA vectors for future therapy is promising. Agrawal and Iyer (49) have reviewed the advantages and disadvantages of using an antisense strategy for therapeutic purposes. Our results show that, in spite of being injected i.p., siRNA-expressing plasmids localize intracranially and effectively down-regulate uPAR and uPA. RT-PCR of the brain tissue showed that although plasmid localization was observed in the brain, no OAS1 induction was detected. This indicates that the presence of a polyadenylic acid tail probably prevented the induction of an IFN-like response. In conclusion, the RNAi-mediated down-regulation of uPAR and uPA has clear clinical implications for the treatment of gliomas as well as other cancers.


   Acknowledgments
 
We thank Shellee Abraham for manuscript preparation, Diana Meister and Sushma Jasti for manuscript review, and Noorjehan Ali for technical assistance.


   Footnotes
 
Grant support: National Cancer Institute grants CA 75557, CA 92393, CA 95058, and CA 116708; National Institute of Neurological Disorders and Stroke NS47699 and NS57529; Caterpillar, Inc.; and OSF Saint Francis, Inc., Peoria, IL (J.S. Rao).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 12/20/06; revised 2/28/07; accepted 3/ 6/07.


   References
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 Materials and Methods
 Results
 Discussion
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