RNAi-mediated inhibition of cathepsin B and uPAR leads to decreased cell invasion, angiogenesis and tumor growth in gliomas


RNA interference (RNAi) provides a powerful method for gene silencing in eukaryotic cells, including proliferating mammalian cells. Here, we determined whether RNAi could be utilized to inhibit the expression of proteases implicated in the extracellular matrix degradation, which is characteristic of tumor progression. We have previously shown that antisense stable clones of uPAR and cathepsin B were less invasive and did not form tumors when injected intracranially ex vivo. Since antisense-mediated gene silencing does not completely inhibit the translation of target mRNA and high molar concentrations of antisense molecules are required to achieve gene silencing, we used the RNAi approach to silence uPAR and cathepsin B in this study. We found that the expression of double-stranded RNA leads to the efficient and specific inhibition of endogenous uPAR and cathepsin B protein expression in glioma cell lines as determined by Western blotting. We also found the RNAi of uPAR and cathepsin B reduces glioma cell invasion and angiogenesis in in vitro and in vivo models. Intratumoral injections of plasmid vectors expressing hpRNA for uPAR and cathepsin B resulted in the regression of pre-established intracranial tumors. Further, RNAi for uPAR and cathepsin B inhibited cell proliferation and reduced the levels of pERK and pFAK compared to controls. Taken together, our findings indicate for the first time that RNAi operates in human glioma cells with potential application for cancer gene therapy.


Cancer cell invasion involves the degradation of the extracellular matrix (ECM), and the basement membrane (BM). Additionally, the presence of adhesion molecules, which are mediated by a variety of proteolytic enzymes, is necessary for the remodeling of cell–cell and/or cell–matrix attachments. The proteinases primarily responsible for ECM degradation in vivo are cysteine proteases, serine proteases and matrix metalloproteases (MMPs). All these proteinases have the combined ability to break down the ECM components. The proteolytic activity of cathepsin B, a cysteine protease, involves the direct degradation of ECM proteins, including fibronectin, types I and IV collagen and laminin (Creemers et al., 1998). Cathepsin B also indirectly activates other enzymes involved in the proteolytic cascade that mediates ECM degradation, including metalloproteinases and both soluble and receptor-bound urokinase plasminogen activator (uPA) (Schmitt et al., 1992). In addition, cathepsin B has been suggested to increase MMP activity by inactivating tissue inhibitors of matrix metalloproteinases (TIMPs). Cathepsin B, therefore, could be an important upstream regulator in the activation of pro-uPA/plasminogen and pro-MMPs. Cathepsin B has also been shown to contribute to apoptosis by causing cytochrome c release and caspase-9 and -3 activation (key events in the mitochondrial pathway of apoptosis) (Guicciardi et al., 2001). Increase in cathepsin B expression and reductions in its inhibitor levels were associated with tumor growth, vascularization, invasion and metastasis in various cancers (Kos et al., 2000).

uPA is a specific serine protease that converts plasminogen into its active form, plasmin, which is a broad-spectrum protease involved in the degradation of the ECM. Plasmin is able to promote ECM degradation by directly degrading ECM components and by activating latent collagenases and metalloproteases (Andreasen et al., 1997). Besides its proteolytic function, the uPA–uPAR system promotes various cell responses such as tumor cell migration, adhesion, proliferation and differentiation as a result of its interaction with various integrins and vitronectin (Andreasen et al., 1997). Formation of the uPA–uPAR complex at the cell surface is required for efficient activation of plasmin, a protease that can degrade the components of the ECM and release various growth factors (Naldini et al., 1995). Past studies indicate a significant positive correlation between levels of uPAR and cathepsin B in tumors. Consequently, various approaches have been pursued to interfere with the expression/activity of uPAR and cathepsin B including antisense strategies and the development of inhibitors for uPA, uPA/uPAR interaction and cathepsin B.

Malignant gliomas are characterized by this 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, metallo and cysteine proteinases. Our previous work and that of others suggest a direct correlation between the expression of cathepsin B (Rempel et al., 1994; Sivaparvathi et al., 1995) and uPAR (Yamamoto et al., 1994; Gladson et al., 1995).

Further, our previous studies demonstrated that antisense cathepsin B stable clones and sense cystatin C (inhibitor of cathepsin B) clones of SNB19 glioblastoma cells were less invasive and did not form tumors in nude mice (Konduri et al., 2002; Mohanam et al., 2002). Similarly, antisense stable uPAR clones and Ad-uPAR-transfected glioblastoma cells were less invasive and did not form tumors ex vivo (Go et al., 1997; Mohanam et al., 1997; Mohan et al., 1999). This antisense-mediated disruption of cathepsin B and uPAR expression clearly demonstrated the therapeutic potential of targeting these molecules in vivo. Here, we describe RNAi-mediated inhibition of uPAR and cathepsin B with the intent of effectively inhibiting cathepsin B and uPAR expression in gliomas in vivo.

Double-stranded RNA (dsRNA) inhibits gene expression in a sequence-specific manner by triggering degradation of mRNA. This effect is called RNA interference (RNAi) and has many features in common with post-transcriptional gene silencing in plants. RNAi was shown to be superior to antisense since only a few molecules of dsRNA per cell can trigger gene silencing (Zamore, 2001). RNAi technology is currently being evaluated for its potential use in dsRNA-based gene-silencing therapies. RNAi can be induced through transfection or microinjection of long dsRNA. The dsRNA is cleaved into 19–23-nt RNA fragments known as small interfering RNAs (siRNA). In mammalian cells, siRNA molecules are capable of specifically silencing gene expression without the induction of nonspecific interferon response pathway. The activity of siRNA is high even at low concentrations and without any apparent toxicity. In a recent study, siRNA was quantitatively more efficient than antisense oligonucleotides at suppressing cotransfected GFP expression both in vitro and in vivo (Bertrand et al., 2002). In this study, we report that RNAi is feasible in glioma cells and can be used to specifically knock down the expression of cathepsin B and uPAR with significant therapeutic potential.


Effect of pCU vector on cathepsin B and uPAR protein levels in total cell extracts

RNAi targeted against proteolytic degradation could be an important intervention to prevent cancer cell invasion. Cathepsin B and uPAR have been shown to play significant roles in ECM degradation. Transfection of SNB19 cells with the vector expressing siRNA for cathepsin B and uPAR (pCU) strongly inhibited the expression of both protein as compared to mock/EV and scramble vectors controls (Figure 1a, c). The levels of β-actin determined that equal quantities of protein were loaded in the gel (Figure 1). Quantitative analysis of cathepsin B and uPAR bands by densitometry revealed a significant (P<0.001) decrease in cathepsin B (14–16-fold) and uPAR protein (10–12-fold) and in pCU-transfected cells compared to mock- and EV-transfected cells (Figure 1b, d). Cells transfected with pU and pC vectors inhibited the levels for uPAR and cathepsin B, respectively (Figure 1a, c).

Figure 1

Western blot analysis for uPAR and cathepsin B. SNB19 cells were transfected with mock/EV, SV and a vector encoding siRNA for cathepsin B and uPAR (pCU). Western blot analysis of cathepsin B (a) uPAR (c) protein levels in cell lysates from SNB19 cells transfected with mock/EV, SV and pCU was performed using an antibody specific for cathepsin B and uPAR as described in Materials and methods. β-Actin was simultaneously immuno-probed for loading control. Quantitation of cathepsin B (b) and uPAR protein (d) was obtained by scanning the autoradiograms with densitometry as described in Materials and methods. Data were shown as mean values±s.d. of five different experiments from each group (P<0.001)

Inhibition of tumor cell-induced capillary network formation by pCU vector

Emerging tumors are dependent on the formation of new blood vessels that fuel tumor growth. Because cathepsin B and uPAR have been reported to regulate angiogenesis, we next assessed the effect of pCU on tumor cell-induced angiogenesis. We performed immunohistochemical analysis using factor VIII antigen to evaluate tumor-induced vessel formation in an in vitro coculture system and stained hematoxylin & eosin (H&E) for endothelial cells grown in the presence of conditioned media of SNB 19 cells after transfection with mock/EV, pC, pU or pCU. The results demonstrate that endothelial cells form capillary-like structures in the presence of mock/EV and SV-transfected cells within 48 h; whereas, the pCU vector significantly inhibited tumor cell-induced capillary-like network formation (Figure 2a). The quantification of the branch points and number of branches were undetectable in pCU-transfected cocultures compared to mock and EV (Figure 2b). Further, the effect was less than 50% in pC or pU-treated cocultures when compared to pCU vector in relation to capillary-like structures. To confirm the in vitro coculture experiments, we examined whether the pCU vector can inhibit tumor angiogenesis in vivo as assessed by the dorsal chamber model. Implanted chambers containing mock/EV and SV-transfected SNB19 cells resulted in the development of microvessels (as indicated by arrows) with curved thin structures and many tiny bleeding spots. In contrast, implanted chambers of SNB19 cells transfected with the pCU vector did not result in the development of any additional microvessels (Figure 2c).

Figure 2

RNAi inhibits tumor cell-induced capillary network formation. SNB19 cells were transfected with mock/EV, SV, pC, pU and pCU for 24 h. Then, cells were cocultured with human dermal endothelial cells for 48 h. After incubation, cells were fixed and blocked with 2% bovine serum albumin for 1 h and endothelial cells were probed with antibody for factor VIII antigen (factor VIII antibody, DAKO Corporation, Carpinteria, CA, USA) or H&E staining and examined under a laser scanning confocal microscope (a). Quantification of angiogenesis in cocultures infected with mock, EV or pCU vector as described in Materials and methods (b). Inhibition of tumor angiogenesis in SNB19 cells infected with pCU vector by mouse dorsal window assay as described in Materials and methods (c). PV, pre-existing vasculature; TN, tumor-induced vasculature

Inhibition of migration of SNB19 spheroids by siRNA

To determine whether cathepsin B and uPAR siRNA expression is capable of influencing tumor cell migration and proliferation, we transfected SNB19 spheroids with the pCU vector. As shown in Figure 3a, there was much higher cell migration from spheroids transfected with mock and EV/SV and up to 50% inhibition of migration was observed with single construct transfected spheroids. However, cell migration from tumor spheroids was completely inhibited in spheroids transfected with the pCU vector. The migration of the mock- and EV/SV-transfected spheroids was significantly higher (P<0.001) compared to pC-, pU- and pCU-transfected spheroids as quantitated by the number of cells migrating out from the spheroids (Figure 3b). The migration of cells from the spheroids were inhibited with bicistronic construct compared to single RNAi constructs for these molecules.

Figure 3

RNAi inhibits glioma cell migration and invasion. SNB19 GFP spheroids were infected with mock, EV/SV, pC, pU and pCU. After 72 h, single glioma spheroids were placed in the center of a vitronectin-coated well in a 96-well plate and cultured for 48 h. At the end of the migration assay, spheroids were fixed and photographed (a). The migration of cells from the spheroids was measured using a microscope calibrated with a stage and ocular micrometer (b) as described in Materials and methods. The data shown were the mean value±s.d. of the results from four independent experiments from each group. SNB19 cells were trypsinized 72 h after transfection with mock, EV/SV, pC, pU and pCU, washed with PBS and resuspended in serum-free medium. Invasion assays were carried out in a 12-well transwell unit (Costar, Cambridge, MA, USA) on polycarbonate filters with 8-μm pores coated with Matrigel. After a 24 h incubation period, the cells that had passed through the filter into the lower wells were stained, counted and photographed under a light microscope (c). The percentage of invasion was quantitated as described in Materials and methods (d). Values are mean±s.d. from five different experiments (P<0.001). Spheroids of SNB19 cells were transfected with mock, EV/SV, pC, pU and pCU and stained with DiI and cocultured with DiO-stained fetal rat brain aggregates. Progressive destruction of fetal brain aggregates by tumor spheroids was observed (e). Quantification of remaining fetal brain aggregates by SNB19 spheroids infected with EV/SV, pC, pU or pCU vector as described in Materials and methods (f)

siRNA against cathepsin B and uPAR inhibits tumor cell invasion

To evaluate the impact of siRNA-mediated inhibition of cathepsin B and uPAR on glioma invasiveness, we utilized a two-model system. SNB19 cells transfected with mock and EV/SV extensively invaded the Matrigel-coated transwell inserts as observed by the intense staining of the cells. In contrast, the pC, pU and pCU-transfected cultures had markedly less invasiveness through the reconstituted basement membrane, compared to mock- and EV/SV-transfected cells (Figure 3c). Quantitative determination of invasion confirmed that SNB19 cells transfected with the pC, pU and pCU vector invaded only 55%, 40% and 6%, respectively, as compared to mock- and EV/SV-transfected controls (Figure 3d). Inhibition of the invasive behavior of these cells as determined by Matrigel invasion assay was much higher in the bicistronic construct transfected cells when compared to the single construct.

We further examined the extent of effect of pCU in spheroid invasion assay. In the spheroid coculture assay, control glioma spheroids and spheroids transfected with EV/SV progressively invaded fetal rat brain aggregates and resulted in partial to almost complete inhibition of invasion of spheroids transfected with pCU (Figure 3e). Quantitation of the fetal rat brain aggregates revealed that 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 pCU vector did not invade the fetal rat brain aggregates. At 72 h, the rat brain aggregates revealed invasion approximately 90%, 85%, 55% and 35% in the mock- EV/SV, pC- and pU-transfected cocultures, but only 2–3% invasion in the pCU-transfected cocultures (Figure 3f). Taken together, these findings provide strong evidence that RNAi-mediated silencing of cathepsin B and uPAR strongly inhibits glioma cell invasion in both in vitro models, and that the effect was much higher with bicistronic construct compared to single constructs.

siRNA-mediated downregulation of cathepsin B and uPAR reduces the proliferation of SNB 19 cells

We used the standard MTT assay to assess the effect of the siRNA vectors (mock/EV, SV, pC, pU and pCU) on proliferation of cells cultured on vitronectin-coated microplates (Figure 4). At 72 h after infection, the pC, pU and pCU vector-infected SNB19 cells showed a decrease in proliferation relative to that of SNB19 and vector controls. pCU-transfected cell did not show any appreciable growth even after 7 days of transfection. No floating cells or cell derby was seen in any of the transfected cells even after 7 days of assay indicating the absence of apoptosis.

Figure 4

RNAi-mediated downregulation of uPAR and cathepsin B reduces SNB 19 glioma cell proliferation. Briefly, 5 × 104 SNB19 cells transfected with SV, pU, pC and pCU were seeded in VN-coated 96-well microplates under serum-free conditions. The number of viable cells was assessed by the MTT assay. Shown are the mean (±s.d.) values from three separate experiments

siRNA-mediated downregulation of uPAR and cathepsin B inhibits ERK1/2 and FAK phosphorylation

To determine the effect of downregulation of uPAR and cathepsin B on signaling pathway molecules, we assayed for the phosphorylation of ERK and FAK by Western blotting both of which are directly involved in tumor cell survival, migration and proliferation. From Figure 5 it is clearly evident that RNAi mediated simultaneous downregulation of uPAR and cathepsin B retards the phosphorylation of ERK1/2 and FAK and the effect was much less with single constructs.

Figure 5

RNAi-mediated downregulation of uPAR and cathepsin B reduces the phosphorylation of ERK and FAK: Western blot analysis of total and phospho-ERK, FAK, proteins using their specific antibodies after transfection of SNB19 cells with mock, EV, SV, pU, pC and pCU constructs. β-Actin levels served as loading control

Cathepsin B and uPAR siRNA suppresses intracranial tumor growth

We used an intracranial tumor model to assess potential effects of RNAi-mediated inhibition on pre-established tumor growth in vivo. The brain sections of the untreated (mock) and EV/SV-treated control groups were characterized by large spread tumor growth by H&E staining and high GFP fluorescence after a 5-week follow-up period (Figure 6a, b). However, we could not detect GFP fluorescence in the brain sections of mice treated with the pCU vector (Figure 6a, b). Further quantification of H&E-stained brain sections were scored by a neuropathologist who was blind to the treatment, revealed no difference in tumor size between the mock, EV and SV treatment groups and significant regression of tumor growth 55% and 65% in the pC- and pU-treated groups compared to controls. However, total regression of pre-established tumors was revealed in the pCU-treated group (Figure 6). These results demonstrate that RNAi-mediated suppression of cathepsin B and uPAR significantly inhibited intracranial tumor growth.

Figure 6

RNAi-mediated regression of pre-established tumor growth. SNB19 GFP tumor cells were injected intracerebrally with the help of a stereotactic frame into nude mice. After 1 week, either an EV/SV or a vector expressing siRNA for cathepsin B and uPAR (pCU) or separately by pC or pU was injected into the brain using an Alzet mini-osmotic pump. Photomicrographed tumor sections were observed for GFP fluorescence (a) and subsequently stained with hematoxylin and eosin (b). Semiquantification of tumor volume in mock/EV, pU, pC and pCU vector treated groups after 5 weeks as described in Materials and methods. Data shown are the ±s.d. values from six animals from each group (*P<0.001) (c)


RNAi, or post-transcriptional gene silencing, is a relatively novel technology that offers an opportunity to inhibit gene expression. siRNAs are more potent inhibitors of gene expression with less toxicity, making them more robust tools for many investigators (McCaffrey et al., 2002; Xia et al., 2002). Reports indicate that siRNA can also function in vivo (Song et al., 2003). Given the potency and effectiveness of RNAi as a gene silencing tool and the ease with which dsRNA can be introduced into cells and tissues, has significant therapeutic potential for human applications. The feasibility of RNAi-mediated gene silencing as a novel tool to arrest tumor growth and kill cancer cells is being tested in several laboratories and their initial results are promising (Jiang et al., 2002; Wilda et al., 2002). This study demonstrates that RNAi-mediated inhibition of cathepsin B and uPAR inhibits glioma invasion, angiogenesis and growth.

The key features of glioblastoma multiforme are vasculature remodeling and destruction of surrounding normal brain tissue, local infiltration and growth. The invasive behavior of glioblastoma appears to depend partly on the proteolytic destruction of ECM. Several reports have indicated that proteases are involved in tumor growth and invasion both at the primary and metastatic sites (Lochter et al., 1997).

Our research and that of others have previously demonstrated that cathepsin B and uPAR levels are overexpressed during the progression of glioma tumor growth (Rempel et al., 1994, Yamamoto et al., 1994; Gladson et al., 1995; Sivaparvathi et al., 1995; Strojnik et al., 1999). Our previous work demonstrated that antisense stable clones of SNB19 for cathepsin B (Mohanam et al., 2002) and uPAR (Go et al., 1997; Mohanam et al., 1997) were less invasive in vitro and formed very small tumors in vivo. The SNB19 cell line possesses high invasive potential and shows increased protein and mRNA expression of cathepsin B and uPAR. In the present study, we have developed a vector-based siRNA expression system that can induce RNAi in glioma cells. We directed 21-nt siRNA against cathepsin B and uPAR (pCU) in order to simultaneously suppress cathepsin B and uPAR expression. We have shown that the endogenous levels of cathepsin B and uPAR proteins are reduced in SNB19 glioblastoma cell line by transient transfection with the pCU vector using Western blotting. Very few cells migrated from pCU-transfected SNB19 spheroids as compared to that of the mock and EV/SV controls, thereby indicating the role of cathepsin B and uPAR in cell migration.

However, conflicting results have been reported on the effect of cathepsin B on the motility of cancer cells. Exposure of human transitional carcinoma cells to E-64 resulted in a dose-dependent reduction in motility (Redwood et al., 1992). Krueger et al. (1999) reported a 25–50% inhibition of motility by human osteosarcoma cells that expressed an antisense cathepsin B construct. In contrast, cathepsin B expression and cysteine proteinase inhibitors did not alter the motility of murine SCC-VII squamous carcinoma cells (Coulibaly et al., 1999) and B16F1 melanoma cells (Szpaderska and Frankfater, 2001). Similarly, transfection of metastatic human prostate PC3M and melanoma A375M cell lines with antisense human cathepsin B cDNA did not have any effect on cell migration (Szpaderska and Frankfater, 2001). Previous studies have shown that uPAR interacts with the extracellular domain of integrins allowing association with the cytoskeleton, thereby mediating cell adhesion and migration (Kjoller, 2002). It has been reported that uPA and uPAR are involved in cellular migration (Planus et al., 1997). Our results demonstrate that the downregulation of uPAR and cathepsin B induces the downregulation of ERK1/2 and FAK phosphorylation which are directly responsible for cell survival and proliferation. The involvement of uPAR in the ERK–FAK cascade has previously been reported in human carcinoma cells HEp3 (Aguirre Ghiso, 2002), but the role of cathepsin B still remains unclear. Our results demonstrate that a combinational downregulation of uPAR and cathepsin B is more effective in inhibiting phosphorylation of ERK1/2 and FAK. The direct or indirect interaction of cathepsin B and uPAR warrants further investigation. Earlier studies have shown that the binding of uPA and uPAR in McF-7 cells activates ERK1 and 2, which is required in cell motility (Nguyen et al., 1998). It has also been shown that in the absence of EGFR, alternate pathways like uPAR to ERK are activated, in which uPAR is involved in cell motility (Jo et al., 2003). Binding of uPA to uPAR is critical for proliferation adhesion, migration and proteolysis of cancer cells (Aguirre-Ghiso et al., 2001).

The acquisition of tumor cell invasiveness is one of the important aspects of tumor progression. There are several reports to indicate that expression of cathepsin B and uPAR are essential components of the invasion process. Transfection with the pCU vector inhibited the invasiveness of SNB19 cells and spheroids in the Matrigel invasion and spheroid coculture assays. The requirement of cathepsin B for Matrigel invasion could be due to its interaction with a network of proteases. Cathepsin B was shown to activate precursors of serine proteinases to their active forms, such as pro-uPA (Kobayashi et al., 1991) and metalloproteinases, such as pro-stromelysin (Murphy et al., 1992). Invasiveness through Matrigel of transformed human breast epithelial cell lines was related to cathepsin B expression and was inhibited by cysteine proteinase inhibitors (Bervar et al., 2003). In ovarian cancer cells, inhibition of cell surface cathepsin B prevents activation of pro-uPA, and subsequently, invasion of the carcinoma cells through Matrigel (Kobayashi et al., 1993). Cathepsin B activity in human colon cancer is associated with the invasiveness of cancer cells, endothelial cells and inflammatory cells as well as apoptotic and necrotic cell death (Hazen et al., 2000). Stable transfection of human cathepsin B cDNA into a murine squamous cell carcinoma resulted in a nearly three-fold increase in the level of secreted pro-cathepsin B and a 22% increase in invasiveness across a Matrigel-reconstituted basement membrane (Coulibaly et al., 1999). uPA and uPAR are known to be overexpressed in various malignancies including breast, ovarian and gastric cancers, and have been demonstrated to be essential in the maintenance of the invasive and metastatic phenotype (Reuning et al., 1998). Several studies demonstrated that uPA inhibitors, uPA antibodies and antisense stable clones for uPA and uPAR have been shown to inhibit the invasiveness of tumor cells into ECM as well as amniotic and chick chorioallantoic membranes (Holst-Hansen et al., 1996; Mohanam et al., 1997; Alonso et al., 1998; Mohanam et al., 2002).

Our data showed that local intracranial delivery of pCU using mini-osmotic pumps effectively inhibited human malignant glioma growth. Mini-osmotic pumps maintain a well-defined and consistent pattern of drug exposure for a significant period of time and can be used successfully to deliver agents to the brain (Kisker et al., 2001). Our study also establishes that downregulation of cathepsin B and uPAR results in inhibition of tumor-induced angiogenesis. We used a coculture assay in vitro to test the effect of pCU on angiogenesis. The results demonstrate that cathepsin B and uPAR play important roles in stimulating angiogenesis, suggesting a possible mechanism of action for the in vivo antitumor activity of pCU in the intracranial tumor model. Intense staining for cathepsin B is present in endothelial cells of neovessels but not in pre-existing microvasculature in prostate (Sinha et al., 1995) and human gliomas (Mikkelsen et al., 1995). Likewise, strong immunostaining of cathepsin B was observed in rat brain microvascular endothelial cells as they formed capillary tubes in vitro (Keppler et al., 1996). Since cathepsin B was shown to be an inhibitor of TIMPs (Kostoulas et al., 1999) and TIMPs are inhibitors of angiogenesis, cathepsin B could also stimulate angiogenesis, which has an important role in tumor spread. It has been reported in the host results in the enhanced adhesion of uPAR-bearing endothelial cells to the ECM protein vitronectin due to the loss of PAI-1, which adversely affects cell motility and resultant neovascularization (Gutierrez et al., 2000). Previous studies demonstrated that several antagonistic peptides identified by bacteriophage display as blocking uPA binding have been shown to inhibit angiogenesis and primary tumor growth in syngenic mice (Min et al., 1996). Similar observations were reported that an octameric peptide derived from the nonreceptor-binding region of uPA was shown to inhibit tumor growth and angiogenesis in breast cancer (Guo et al., 2000) and in combination with cisplatin in glioblastoma (Li et al., 1998). The antiangiogenic effects of pCU are also in agreement with previous studies showing that inhibition of uPAR activity suppressed retinal neovascularization, possibly through a reduction in cell-associated proteolytic activity, cell signaling or cell–matrix adhesion necessary for cell migration during angiogenesis (McGuire et al., 2003). Synthetic cysteine proteinase inhibitors, selective for cathepsin B, have been shown to significantly reduce the invasiveness of MCF10AT cells (Bervar et al., 2003). Several studies using selective inhibitors of uPA or small, synthetic cyclic, competitive uPA antagonists derived from the binding sites of uPAR resulted in highly significant reduction of tumor burden (Evans et al., 1998; Sato et al., 2002). Very recent studies have also shown that fusion of diphtherotoxin with the ATF-uPA caused a statistically significant regression of glial tumors and was highly potent and selective at killing uPAR expressing glioblastoma cell lines (Vallera et al., 2002).

In conclusion, we have demonstrated that the RNAi-mediated targeting of cathepsin B and uPAR suppressed pre-established intracranial tumor growth, possibly by inhibiting angiogenesis and invasiveness. These results also strongly suggest that the siRNA-mediated downregulation of target gene expression is sufficiently stable within the brain microenvironment. Consequently, our data raise hope for future application of siRNA in glioma patients.

Materials and methods

Construction of a vector expressing siRNA for cathepsin B and uPAR

pcDNA 3 was used for the construction of a vector expressing siRNA for both cathepsin B and uPAR downstream of the cytomegalovirus (CMV) promoter (Scheme 1). The uPAR sequence from +77 to +98 was used as the target sequence and for convenience, a self-complimentary oligo was used. The uPAR sequence 21 bases in length with a nine base loop region with BamHI sites incorporated at the ends (IndexTermgatcctacagcagtggagagcgattatatataataatcgctctccactgctgtag) was used. The oligo was self-annealed in 6 × SSC using standard protocols and ligated on to the BamHI site of a pcDNA-3 vector plasmid. Similarly, a cathepsin B complimentary sequence from +732 to +753 (IndexTermtcgaggtggcctctatgaatcccaatatataattgggattcatagaggccacc) with XhoI sites incorporated at the ends was ligated into the XhoI site of the vector containing the siRNA sequence for uPAR. This finally resulted in a siRNA expression plasmid for cathepsin B and uPAR designated pCU. Single siRNA expression vectors for uPAR (pU) and cathepsin B (pC) were also constructed. The orientation of either insert in the single or bicistronic did not matter since the oligos were self-complimentary and had bilateral symmetry. BGH poly A terminator served as a stop signal for RNA synthesis for all three constructs.

Scheme 1

Schematic representation showing the plausible formation of hpRNA molecules from a single CMV-driven dual-inverted repeat construct for cathepsin B and uPAR. The powerful CMV viral promoter drives the formation of an RNA molecule that possesses self-complementary inverted repeats for cathepsin B and uPAR as described in Materials and methods

Cell culture and transfection conditions

The human glioblastoma cell line SNB19 was maintained in DMEM F-12 (Sigma Chemical Co., St Louis, MO, USA) supplemented with 10% FCS, 100-μg/ml streptomycin and 100-U/ml penicillin (Invitrogen, Carlsbad, CA, USA) at 37°C in a humidified 5% CO2 atmosphere. Cells were transfected with pC pU or pCU plasmid expressing siRNA using the Lipofectamine reagent (Invitrogen Grand Island, NY, USA) as per the manufacturer's instructions. After transfection, cells were incubated in serum-containing medium for 48 h.

Western blotting

SNB19 cells were transfected with mock, EV, SV, pC, pU or pCU and cultured for 48 h. At the end of incubation, cells were harvested, washed twice with cold PBS and lysed in buffer (150 mM NaCl, 50 mM Tris-HCl, 2 mM EDTA, 1% NP-40, pH 7.4), containing protease inhibitors. Equal amounts of protein (30 μg/lane) from supernatants or cells were electrophoresed under nonreducing conditions on 10% acrylamide gels. After SDS–PAGE, proteins were transferred to a polyvinylidene difluoride membrane (Bio-Rad). To block nonspecific binding, the membrane was incubated for 2 h in PBS with 0.1% Tween-20 [T-PBS] containing 5% nonfat skim milk for 2 h. Subsequently, the membrane was incubated for 2 h with antibody against cathepsin B, uPAR, ERK, pERK, FAK or pFAK, respectively, in T-PBS + 5% nonfat milk. After washing in T-PBS, protein on the membrane was visualized using the ECL™ detection kit with a peroxidase-labeled antirabbit antibody (Amersham Pharmacia Biotech, Amersham, UK) as per the manufacturer's instructions. For loading control, the membranes were stripped and probed with monoclonal antibodies for β-actin, as per standard protocols.

Cell proliferation assay

Cell growth was assessed by MTS assay. To detect the effect of these constructs on the growth of the SNB19 cells in vitro, we measured viable cell mass using the Cell Titer 96™ colorimetric assay. Glioblastoma cells (5 × 103) were seeded in triplicate into 96- or 24-well plates and allowed to grow for 24 h before transfection with culture medium alone (mock), EV, SV, pC, pU and pCU vectors for 48 h. These cells were then changed to serum containing medium and allowed different time intervals. Before each time point, we added MTS reagent and continued incubation for an additional 2 h to permit color development. A490 was measured in each well using an ELISA plate reader. Absorbance readings for short-term vs long-term cell cultures was compared, and the effects of these constructs were interpreted with respect to the growth of corresponding untreated/control groups. Percent inhibition of growth due to the siRNA constructs was calculated relative to the growth rate of the same cells in the same medium minus these contructs.

In vitro angiogenic assay

SNB19 cells (2 × 104) were seeded in eight-well chamber slides and transfected with mock, EV, SV, pU, pC and pCU as per standard protocols. After a 24 h incubation period, the medium was removed and 4 × 104 human dermal endothelial cells were seeded and allowed to coculture for 72 h. After fixation in 3.7% formaldehyde, endothelial cells were immunoprobed for factor VIII antigen. Factor VIII antibody was purchased from the DAKO Corporation (Carpinteria, CA, USA). Cells were washed with PBS and incubated with FITC-conjugated secondary antibody for 1 h and were then washed and examined under a fluorescent microscope. Similar slides of endothelial cells grown in the presence of conditioned media from the SNB 19 mock-, EV-, SV-, pU-, pC- or pCU-transfected cell were stained with H&E to visualize network formation. Image Pro software was used for quantification of angiogenesis, the degree of angiogenesis was measured by the following method: number of branch points and the total number of branches per point were counted at random (per 10 fields), with the product indicating the degree of angiogenesis compared to the controls.

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 (Leunig et al., 1992). Sterile small-animal surgical techniques were followed. Mice were anesthetized by i.p. injection with ketamine (50 mg/kg) zylazine (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. A volume of 20 μl of PBS was used to wet the membranes. SNB 19 cells (2 × 106) (mock-, EV- SV- or pCU-transfected), suspended in 100–150 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 ½ 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 were anesthetized with ketamine/xylazine and killed by intracardiac perfusion with saline (10 ml) followed by 10 ml of 10% formalin/0.1 M phosphate solution and followed by 0.001% FITC solution in PBS. The animals were carefully skinned around the implanted chambers and the implanted chambers were removed from the s.c. air fascia. The skin fold covering the chambers were photographed under visible light and for FITC fluorescence. The numbers of blood vessels within the chamber in the area of the air sac fascia were counted and their lengths measured.


A suspension of 2 × 106 cells in Dulbecco's modified Eagle medium of a GFP-expressing variant of SNB19 cells was seeded on ultra-low attachment 100 mm tissue culture plates and cultured until spheroid aggregates formed. Spheroids measuring 150 μm in diameter (about 4 × 104 cells/spheroid) were selected, transfected with mock, EV, SV, pC, pU and pCU and cultured for 48 h. At 72 h after transfection, a single glioma spheroid was placed in each well of a vitronectin-coated (50 μg/ml) 96-well microplate and cultured with 200 μl of serum-free medium. 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 measured using a microscope calibrated with a stage and ocular micrometer and used as an index of cell migration.

Boyden chamber invasion assay

The in vitro invasiveness of SNB19 cells in the presence of the vector expressing siRNA for cathepsin B and uPAR was assessed using a modified Boyden chamber assay. SNB19 cells were transfected with mock, EV, SV, pU, pC or pCU vector expressing siRNA for cathepsin B and uPAR single or together for 48 h. Cells (1 × 106) were suspended in 600 μl of serum-free medium supplemented with 0.2% BSA and placed in the upper compartment of the transwell chambers (Corning Costar Fischer Scientific Cat No. 07-200-158, Pittsburg, PA, USA) coated with Matrigel (0.7 mg/ml). The lower compartment of the chamber was filled with 200 μl of serum-free medium and the cells were allowed to migrate for 24 h. After incubation, the cells were fixed and stained with Hema-3 and photographed. Quantification of the invasion assay was performed as described previously (Mohanam et al., 1993; Mohan et al., 1999).

Spheroid assay

SNB19 glioblastoma cells (3 × 106) were seeded in 100 mm tissue culture plates (Corning, Corning, NY, USA) precoated with 0.75% agar prepared in DMEM and cultured until spheroid aggregates formed. Spheroids of 100–200 μm in diameter were selected and transfected with mock, EV, SV, pC, pU and pCU for 48 h. At 3 days after infection, SNB19 spheroids were stained with the fluorescent dye DiI and placed in contact 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 (Go et al., 1997). The remaining volume of brain aggregates or tumor spheroids during cocultures was determined as described previously (Go et al., 1997).

Animal experiments

SNB19 GFP cells (2 × 106) were injected into the brains of nude mice using a stereotactic frame. After 8–10 days, the mice were treated with mock, EV/SV, pC, pU and pCU. The in vivo intracranial delivery of vectors was performed using Alzet (Direct Corp. Cupertion, CA, USA) mini-osmotic pumps at the rate of 0.25 μl/h, mock (PBS) of 150 μg vector DNA, 150 μg pC, 150 μg pU and 150 μg pCU. All experiments were performed in compliance with institutional guidelines set by the Institutional Animal Control Users Committee that approves experiments at the University of Illinois College of Medicine at Peoria. After 5 weeks, or when the control mice started showing symptoms, mice were euthanized by cardiac perfusion with formaldehyde. The brains were then removed and paraffin embedded as per standard protocols. Sections were prepared and observed for GFP expression or were stained with H&E. The sections were blindly reviewed and scored semiquantitatively for tumor size in each case. The average tumor area per section was used to calculate tumor volume and compared between controls and treated groups.



RNA interference


urokinase-type plasminogen activator


uPA receptor




simian virus type 40


polymerase chain reaction


phosphate-buffered saline






3,3'-dioctadecyloxacarbocyanine perchlorate


green fluorescent protein


extracellular matrix


  1. Aguirre Ghiso JA . (2002). Oncogene, 21, 2513–2524.

  2. Aguirre-Ghiso JA, Liu D, Mignatti A, Kovalski K and Ossowski L . (2001). Mol. Biol. Cell, 12, 863–879.

  3. Alonso DF, Tejera AM, Farias EF, Bal de Kier JE and Gomez DE . (1998). Anticancer Res., 18, 4499–4504.

  4. Andreasen PA, Kjoller L, Christensen L and Duffy MJ . (1997). Int. J. Cancer, 72, 1–22.

  5. Bertrand JR, Pottier M, Vekris A, Opolon P, Maksimenko A and Malvy C . (2002). Biochem. Biophys. Res. Commun., 296, 1000–1004.

  6. Bervar A, Zajc I, Sever N, Katunuma N, Sloane BF and Lah TT . (2003). Biol. Chem., 384, 447–455.

  7. Coulibaly S, Schwihla H, Abrahamson M, Albini A, Cerni C, Clark JL, Ng KM, Katunuma N, Schlappack O, Glossl J and Mach L . (1999). Int. J. Cancer, 83, 526–531.

  8. Creemers LB, Hoeben KA, Jansen DC, Buttle DJ, Beertsen W and Everts V . (1998). Matrix Biol., 16, 575–584.

  9. Evans DM, Sloan-Stakleff K, Arvan M and Guyton DP . (1998). Clin. Exp. Metast., 16, 353–357.

  10. Gladson CL, Pijuan-Thompson V, Olman MA, Gillespie GY and Yacoub IZ . (1995). Am. J. Pathol., 146, 1150–1160.

  11. Go Y, Chintala SK, Mohanam S, Gokaslan Z, Venkaiah B, Bjerkvig R, Oka K, Nicolson GL, Sawaya R and Rao JS . (1997). Clin. Exp. Metast., 15, 440–446.

  12. Guicciardi ME, Miyoshi H, Bronk SF and Gores GJ . (2001). Am. J. Pathol., 159, 2045–2054.

  13. Guo Y, Higazi AA, Arakelian A, Sachais BS, Cines D, Goldfarb RH, Jones TR, Kwaan H, Mazar AP and Rabbani SA . (2000). FASEB J., 14, 1400–1410.

  14. Gutierrez LS, Schulman A, Brito-Robinson T, Noria F, Ploplis VA and Castellino FJ . (2000). Cancer Res., 60, 5839–5847.

  15. Hazen LG, Bleeker FE, Lauritzen B, Bahns S, Song J, Jonker A, Van Driel BE, Lyon H, Hansen U, Kohler A and Van Noorden CJ . (2000). J. Histochem. Cytochem., 48, 1421–1430.

  16. Holst-Hansen C, Johannessen B, Hoyer-Hansen G, Romer J, Ellis V and Brunner N . (1996). Clin. Exp. Metast., 14, 297–307.

  17. Jiang Y, Goldberg ID and Shi YE . (2002). Oncogene, 21, 2245–2252.

  18. Jo M, Thomas KS, O'Donnell DM and Gonias SL . (2003). J. Biol. Chem., 278, 1642–1646.

  19. Keppler D, Sameni M, Moin K, Mikkelsen T, Diglio CA and Sloane BF . (1996). Biochem. Cell. Biol., 74, 799–810.

  20. Kisker O, Becker CM, Prox D, Fannon M, D'Amato R, Flynn E, Fogler WE, Sim BK, Allred EN, Pirie-Shepherd SR and Folkman J . (2001). Cancer Res., 61, 7669–7674.

  21. Kjoller L . (2002). Biol. Chem., 383, 5–19.

  22. Kobayashi H, Moniwa N, Sugimura M, Shinohara H, Ohi H and Terao T . (1993). Jpn. J. Cancer Res., 84, 633–640.

  23. Kobayashi H, Schmitt M, Goretzki L, Chucholowski N, Calvete J, Kramer M, Gunzler WA, Janicke F and Graeff H . (1991). J. Biol. Chem., 266, 5147–5152.

  24. Konduri SD, Yanamandra N, Siddique K, Joseph A, Dinh DH, Olivero WC, Gujrati M, Kouraklis G, Swaroop A, Kyritsis AP and Rao JS . (2002). Oncogene, 21, 8705–8712.

  25. Kos J, Krasovec M, Cimerman N, Nielsen HJ, Christensen IJ and Brunner N . (2000). Clin. Cancer Res., 6, 505–511.

  26. Kostoulas G, Lang A, Nagase H and Baici A . (1999). FEBS Lett., 455, 286–290.

  27. Krueger S, Haeckel C, Buehling F and Roessner A . (1999). Cancer Res., 59, 6010–6014.

  28. Leunig M, Yuan F, Menger MD, Boucher Y, Goetz AE, Messmer K and Jain RK . (1992). Cancer Res., 52, 6553–6560.

  29. Li H, Lu H, Griscelli F, Opolon P, Sun LQ, Ragot T, Legrand Y, Belin D, Soria J, Soria C, Perricaudet M and Yeh P . (1998). Gene Therapy, 5, 1105–1113.

  30. Lochter A, Galosy S, Muschler J, Freedman N, Werb Z and Bissell MJ . (1997). J. Cell. Biol., 139, 1861–1872.

  31. McCaffrey AP, Meuse L, Pham TT, Conklin DS, Hannon GJ and Kay MA . (2002). Nature, 418, 38–39.

  32. McGuire PG, Jones TR, Talarico N, Warren E and Das A . (2003). Invest. Ophthalmol. Vis. Sci., 44, 2736–2742.

  33. Mikkelsen T, Yan PS, Ho KL, Sameni M, Sloane BF and Rosenblum ML . (1995). J. Neurosurg., 83, 285–290.

  34. Min HY, Doyle LV, Vitt CR, Zandonella CL, Stratton-Thomas JR, Shuman MA and Rosenberg S . (1996). Cancer Res., 56, 2428–2433.

  35. Mohanam S, Chandrasekar N, Yanamandra N, Khawar S, Mirza F, Dinh DH, Olivero WC and Rao JS . (2002). Oncogene, 21, 7824–7830.

  36. Mohan PM, Chintala SK, Mohanam S, Gladson CL, Kim ES, Gokaslan ZL, Lakka SS, Roth JA, Fang B, Sawaya R, Kyritsis AP and Rao JS . (1999). Cancer Res., 59, 3369–3373.

  37. Mohanam S, Chintala SK, Go Y, Bhattacharya A, Venkaiah B, Boyd D, Gokaslan ZL, Sawaya R and Rao JS . (1997). Oncogene, 14, 1351–1359.

  38. Mohanam S, Sawaya R, McCutcheon I, Ali-Osman F, Boyd D and Rao JS . (1993). Cancer Res., 53, 4143–4147.

  39. Murphy G, Atkinson S, Ward R, Gavrilovic J and Reynolds JJ . (1992). Ann. NY Acad. Sci., 667, 1–12.

  40. Naldini L, Vigna E, Bardelli A, Follenzi A, Galimi F and Comoglio PM . (1995). J. Biol. Chem., 270, 603–611.

  41. Nguyen DH, Hussaini IM and Gonias SL . (1998). J. Biol. Chem., 273, 8502–8507.

  42. Planus E, Barlovatz-Meimon G, Rogers RA, Bonavaud S, Ingber DE and Wang N . (1997). J. Cell. Sci., 110, 1091–1098.

  43. Redwood SM, Liu BC, Weiss RE, Hodge DE and Droller MJ . (1992). Cancer, 69, 1212–1219.

  44. Rempel SA, Rosenblum ML, Mikkelsen T, Yan PS, Ellis KD, Golembieski WA, Sameni M, Rozhin J, Ziegler G and Sloane BF . (1994). Cancer Res., 54, 6027–6031.

  45. Reuning U, Magdolen V, Wilhelm O, Fischer K, Lutz V, Graeff H and Schmitt M . (1998). Int. J. Oncol., 13, 893–906.

  46. Sato S, Kopitz C, Schmalix WA, Muehlenweg B, Kessler H, Schmitt M, Kruger A and Magdolen V . (2002). FEBS Lett., 528, 212–216.

  47. Schmitt M, Janicke F, Moniwa N, Chucholowski N, Pache L and Graeff H . (1992). Biol. Chem. Hoppe Seyler, 373, 611–622.

  48. Sinha AA, Wilson MJ, Gleason DF, Reddy PK, Sameni M and Sloane BF . (1995). Prostate, 26, 171–178.

  49. Sivaparvathi M, Sawaya R, Wang SW, Rayford A, Yamamoto M, Liotta LA, Nicolson GL and Rao JS . (1995). Clin. Exp. Metast., 13, 49–56.

  50. Song E, Lee SK, Wang J, Ince N, Ouyang N, Min J, Chen J, Shankar P and Lieberman J . (2003). Nat. Med., 9, 347–351.

  51. Strojnik T, Kos J, Zidanik B, Golouh R and Lah T . (1999). Clin. Cancer Res., 5, 559–567.

  52. Szpaderska AM and Frankfater A . (2001). Cancer Res., 61, 3493–3500.

  53. Vallera DA, Li C, Jin N, Panoskaltsis-Mortari A and Hall WA . (2002). J. Natl. Cancer Inst., 94, 597–606.

  54. Wilda M, Fuchs U, Wossmann W and Borkhardt A . (2002). Oncogene, 21, 5716–5724.

  55. Xia H, Mao Q, Paulson HL and Davidson BL . (2002). Nat. Biotechnol., 20, 1006–1010.

  56. Yamamoto M, Sawaya R, Mohanam S, Rao VH, Bruner JM, Nicolson GL and Rao JS . (1994). Cancer Res., 54, 5016–5020.

  57. Zamore PD . (2001). Nat. Struct. Biol., 8, 746–750.

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We would like to thank Noorjehan Ali for technical assistance, Tina Wilson for preparing the manuscript and Sushma Jasti and Diana Meister for manuscript review. Supported by National Cancer Institute Grants CA 85216, CA 75557 and CA 92393 (to JSR).

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Correspondence to Jasti S Rao.

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Gondi, C., Lakka, S., Dinh, D. et al. RNAi-mediated inhibition of cathepsin B and uPAR leads to decreased cell invasion, angiogenesis and tumor growth in gliomas. Oncogene 23, 8486–8496 (2004). https://doi.org/10.1038/sj.onc.1207879

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  • gliomas
  • invasion
  • cathepsin and uPAR
  • RNA interference
  • angiogenesis

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