Affiliations of authors: Department of Oncology, Hadassah-Hebrew University Medical Center, Jerusalem, Israel (EE, ME, EZ, TP); Cancer and Vascular Biology Research Center, Bruce Rappaport Faculty of Medicine, Technion, Haifa, Israel (IV)
Correspondence to: Israel Vlodavsky, PhD, Cancer and Vascular Biology Research Center, Bruce Rappaport Faculty of Medicine, Technion, Haifa, 31096, Israel (e-mail: vlodavsk{at}cc.huji.ac.il)
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ABSTRACT |
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INTRODUCTION |
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In addition, heparan sulfate moieties in the ECM are responsible for specific binding of members of the heparin-binding family of growth factors (i.e., bFGF, VEGF, KGF, HGF) and serve as the extracellular reservoir for these factors (46). Thus, within the ECM, heparan sulfatebound growth and angiogenic factors are protected, stabilized, and sequestered from their site of action, but under appropriate conditions, they can be readily mobilized to induce growth factordependent processes (i.e., neovascularization and tumor growth). Heparan sulfate molecules are also associated with the cell surface via their core protein and thus are important mediators of cell adhesion (5). Cleavage of heparan sulfate, therefore, results in disassembly of extracellular barriers, promoting cell invasion and releasing heparan sulfatebound bioactive angiogenic and growth-promoting factors.
The mammalian endoglycosidase heparanase is the predominant enzyme responsible for the degradation of heparan sulfate (710). Heparanase activity may therefore play a decisive role in fundamental biologic processes associated with ECM remodeling and cell migration, such as development and morphogenesis, inflammation, angiogenesis, and cancer metastasis (6,7,1115). Heparanase mRNA and protein are preferentially expressed in metastatic cell lines and human tumor tissues (7,8,1416). Moreover, increased heparanase mRNA expression correlates with reduced postoperative survival of cancer patients (17,18). Overexpression of heparanase cDNA in tumor cells with low metastatic potential confers a high metastatic potential after the cells are injected into experimental animals (7). Heparanase has also been shown to elicit an angiogenic response by releasing heparan sulfatebound angiogenic factors sequestered in the ECM (19). A pronounced correlation between heparanase expression and tumor microvessel density has been reported (17,20,21). Heparin, other sulfated polysaccharides, and heparin-mimicking molecules that inhibit heparanase enzymatic activity also reduce the incidence of metastasis in experimental animals (12,14,22,23). However, the use and mode of action of these compounds remains questionable because they lack specificity (24,25).
This study was undertaken to explore alternative strategies for heparanase inhibition by using gene-silencing technologies to specifically suppress heparanase expression in vitro and in vivo and to determine whether heparanase has a role in cancer invasion, metastasis, and angiogenesis. We used a hammerhead ribozyme approach, which is known to be highly effective in gene silencing (26,27). This method uses the homology of a complementary RNA to specifically target an mRNA. To generate a hammerhead ribozyme, a catalytic ribozyme core is flanked by sequences complementary to the target mRNA, which brings the catalytic region into proximity with the targeted mRNA and leads to its cleavage (26). The advantage of this approach over other types of antisense technology is that one ribozyme molecule can cleave many copies of the target mRNA, and thus relatively low levels of ribozyme can achieve efficient inactivation of the target gene. This method is particularly suitable for reducing the amount of abundant mRNAs (26,27). We have designed a hammerhead anti-heparanase ribozyme and explored whether ribozyme-mediated inhibition of heparanase expression affects invasive and adhesive properties of mouse and human cancer cells in vitro, as well as their metastatic and angiogenic potentials in vivo. We also used RNA interfering (RNAi) gene-silencing because this approach is characterized by increased stability and specificity of targeting and has potential therapeutic applications (28,29). We designed a highly specific anti-heparanase siRNA and constructed a vector that enabled its stable expression in cancer cells. The inhibitory effect of siRNA-mediated silencing of endogenous heparanase on mouse B16-BL6 melanoma cell invasion in vitro and lung colonization in vivo was then investigated.
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MATERIALS AND METHODS |
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Methylcholanthrene-induced non-metastatic Eb (L5178Y) T lymphoma cells were provided by Dr. V. Schirrmacher (DKFZ, Heidelberg, Germany) (30,31); cHpaEb cells were obtained by stable transfection of Eb lymphoma cells with the chimeric form of heparanase (cHpa), composed of the human enzyme and the chicken heparanase signal sequence (32). Rat glioma C-6 cells were kindly provided by Dr. Eli Keshet (Hebrew University, Jerusalem, Israel). All cells were grown in RPMI-1640 supplemented with 10% fetal calf serum (FCS), L-glutamine, and antibiotics. Human breast carcinoma MDA-MB-435 and mouse B16-BL6 melanoma cells were purchased from the American Type Culture Collection (Manassas, VA). Cells were cultured in Dulbecco's Modified Eagle Medium (DMEM) containing glucose at 4.5 g/L supplemented with 10% FCS, L-glutamine, and antibiotics.
Synthesis of Anti-Heparanase Hammerhead Ribozymes
Six anti-heparanase hammerhead ribozymes (HpaRz 16) and a control ribozyme (ContRz) that contains the hammerhead catalytic unit but lacks specific heparanase mRNA recognition motifs were generated by in vitro transcription of a single-stranded oligonucleotide template encoding the T7 RNA polymerase promoter followed by a specific ribozyme coding sequence. Two micrograms of each template was transcribed by using 10 U of T7 RNA polymerase in a buffer containing 40 mM TrisHCl (pH 7.5), 10 mM MgCl2, 5 mM dithiothreitol, 400 µM dNTP, and 50 µg/mL bovine serum albumin (BSA). The transcription reaction was performed at 37 °C for 30 minutes and was stopped by heating at 75 °C for 10 minutes. Reaction products were subjected to electrophoresis in a denaturing 15% polyacrylamide gel, visualized by ultraviolet illumination on a thin-layer chromatography screen, excised from the gel, and purified by ethanol precipitation as described (33).
Synthesis of Labeled Heparanase RNA Substrate for Ribozyme Cleavage
To produce the template for heparanase RNA substrate transcription, a 1487-base-pair (bp) fragment was amplified from the full-length heparanase cDNA and subcloned into the pcDNA3 plasmid (Invitrogen, Carlsbad, CA) by using polymerase chain reaction (PCR). The following two primers were used: upper primer, containing the T7 RNA polymerase promoter sequence (5'-GTAATACGACTCACTATAGGTGAGCCCCTCGTTCCTGTCCGTCACCAT-3') and lower primer (5'-TTTTATTTTCAGATGCAGCAGC-3'). The PCR conditions included an initial denaturation at 94 °C for 2 minutes and 30 cycles of denaturation at 94 °C for 15 seconds, annealing at 55 °C for 45 seconds, and extension at 72 °C for 1 minute. Aliquots (15 µL) of the amplified cDNA were separated by electrophoresis through a 1.5% agarose gel, and the cDNA bands were visualized by ethidium bromide staining (7). The 1487-bp PCR product was isolated and used as a template for the in vitro transcription of the heparanase RNA substrate used in the ribozyme cleavage assay. Transcription was performed with 10 U of T7 RNA polymerase in a buffer containing 40 mM TrisHCl (pH 7.5), 10 mM MgCl2, 5 mM dithiothreitol, 400 µM [32P]-labeled dNTP, and 50 µg/mL BSA at 37 °C for 30 minutes. The reaction was stopped by heat inactivation at 75 °C for 10 minutes. The substrate was subjected to electrophoresis in a denaturing 5% polyacrylamide gel and visualized after subsequent autoradiography.
In Vitro Ribozyme Cleavage Reaction
Analysis of ribozyme cleavage in a cell-free system was performed as described elsewhere (33). Briefly, the radioactively labeled heparanase RNA substrate, prepared as described above, was incubated with each one of the six anti-heparanase ribozymes (HpaRz 16) and control ribozyme (ContRz) in a molar ratio of 1 : 50 (ribozyme : substrate), at 37 °C for 15 and 60 minutes. Cleavage products were separated by electrophoresis in a denaturing 5% polyacrylamide gel and visualized by autoradiography.
Construction of Ribozyme Expression Vector
Vectors for the expression of the anti-heparanase ribozyme (HpaRz2) and the control ribozyme (ContRz) were constructed by subcloning DNA fragments that encode for HpaRz2 or ContRz into the expression vector pcDNA3, which contains a gene for hygromycin B resistance (Invitrogen). The oligonucleotide 5'-AGCTTGGCTTCTTCTGATGAGGCCGAAAGGCCGAAAGTAGGTGC-3' and the complementary oligonucleotide 5'-GGCCGCACCTACTTTCGGCCTTTCGGCCTCATCAGAAGAAGCCA-3' were used to generate the HpaRz2 ribozyme, and the oligonucleotides 5'-AGCTTGCGAAGAACTGATGAGGCCGAAAGGCCGAAACATCCAG-3' and 5'-GATCCTGGATGTTTCGGCCTTTCGGCCTCATCAGTTCTTCGCA-3' were used to generate the ContRz ribozyme. Each oligonucleotide was annealed to its complement by mixing equal molar amounts, heating to 80 °C, and cooling to 30 °C slowly. The double-stranded DNA was then subcloned into the multiple cloning site (at HindIII and NotI) of pcDNA3. For both HpaRz2 and ContRz, the sequence of the insert and the region flanking the insert was confirmed by DNA sequencing.
Construction of siRNA Expression Vectors
We used the pSUPER vector (kindly provided by Dr. R. Agami, Division of Tumor Biology, The Netherlands Cancer Institute, Amsterdam, The Netherlands) in which siRNA expression is driven by the H1 RNA promoter to produce small and size-defined RNA transcripts that lack poly-A tails. We generated two different siRNAs. The oligonucleotides 5'-GATCCCCTCTCAAGTCAACCATGATATTCAAGAGATATCATGGTTGACTTGAGATTTTTGGAAA-3' and 5'-AGCTTTTCCAAAAATCTCAAGTCAACCATGATATCTCTTGAATATCATGGTTGACTTGAGAGGG-3' were used to generate anti-heparanase siRNA Si1, and oligonucleotides 5'-GATCCCCACTCCAGGTGGAATGGCCCTTCAAGAGAGGGCCATTCCACCTGGAGTTTTTTGGAAA-3' and 5'-AGCTTTTCCAAAAAACTCCAGGTGGAATGGCCCTCTCTTGAAGGGCCATTCCACCTGGAGTGGG-3' were used to generate anti-heparanase siRNA Si2. Each oligonucleotide pair (100 pmol) was annealed by incubation at 95 °C for 5 minutes and cooled slowly. One microliter of this mixture was then ligated into the pSUPER vector (34), which had been digested with BglII and HindIII. As a result, two vectors, pSi1 and pSi2, expressing Si1 and Si2 siRNA were generated. The empty pSUPER vector was used as a control.
Transfection
Eb lymphoma (0.5 x 106 cells per milliliter) or MDA-MB-435 breast carcinoma (0.3 x 106 cells per milliliter) cells were incubated (4872 hours at 37 °C) with a total of 12 µg of DNA and 6 µL of FuGene transfection reagent (Boehringer, Mannheim, Germany) in 94 µL of Optimem (Gibco-BRL, Gaithersburg, MD) (35,36). Transfected Eb cells were selected with 200 µg/mL hygromycin B (Sigma) and 350 µg/mL G418 (Gibco-BRL). Transfected MDA-MB-435 cells were selected with 200 µg/mL hygromycin B. Stable transfected cells were obtained and routinely maintained in selection medium to avoid overgrowth of non-transfected cells. B16-BL6 melanoma cells were electroporated with pSi1, pSi2, or empty pSUPER constructs (4 x 106 cells in 400 µL of medium containing 10 µg of plasmid DNA) by using a single 70-ms pulse at 140 V and an ECM 830 Electro Square porator and disposable cuvettes (model 640, 4-mm gap; BTX, San Diego, CA). After electroporation, the transfected cells were plated at a density of 0.4 x 106 cells per 100-mm dish and allowed to grow for 2448 hours. Efficiency of transfection (80%) was evaluated 48 hours after electroporation of a vector containing the gene encoding green fluorescent protein by fluorescence microscopy.
RNA Isolation and Reverse TranscriptionPCR
RNA was isolated from 5 x 106 tumor cells with TRIzol (Invitrogen), according to the manufacturer's instructions, and quantified by spectrophotometry. After oligo(dT)-primed reverse transcription of 500 ng of total RNA, the resulting single-stranded cDNA was amplified using TaqDNA polymerase and buffer (Promega, Madison, WI). The primers used were HPU-355 (5'-TTCGATCCCAAGAAGGAATCAAC-3') and HPL-229 (5'-GTAGTGATGCCATGTAACTGAATC-3') for human heparanase; 431-U (5'-ATGCTCTACAGTTTTGCCAAGTG-3')and 876-L (5'-CAGAATTTTTTGCACAGAGAGAA-3') for mouse heparanase; glyceraldehyde 3-phosphate dehydrogenase (GAPDH) S (5'-CCACCCATGGCAAAATTCCATGGCA-3') and GAPDH-AS (5'-TCTAGACGGCAGGTCAGGTCCACC-3') for human GAPDH; L-19-U (5'-ATGCCAACTCTCGTCAACAG-3') and L-19-L (5'-GCGCTTTCGTGCTTCCTT-3')for mouse L19; and HygR-U (5'-AGACCTGCCTGAAACCGAAC-3') and HygR-L (5'-CCGCTCGTCTGGCTAAGAT-3') for the hygromycin B resistance (hygromycin phospho-transferase) gene expressed by both pHpaRz2 and pContRz vectors. PCR conditions for the expression of human genes included an initial denaturation of 4 minutes at 94 °C followed by 26 cycles of denaturation for 45 seconds at 94 °C, annealing for1 minute at 60 °C, and extension for 1 minute at 72 °C. PCR conditions for the expression of mouse genes included an initial denaturation of 2 minutes at 95 °C followed by 24 cycles of denaturation for 15 seconds at 96 °C, annealing for 70 seconds at 58 °C, and extension for 80 seconds at 72 °C (24 cycles). Aliquots (10 µL) of the amplification products were separated by electrophoresis through a 1.5% agarose gel and visualized by ethidium bromide staining. The intensity of each band was quantified using Scion Image software (Scion, Frederick, MD). Only RNA samples that gave completely negative results in PCR without reverse transcriptase were used to rule out the presence of genomic DNA contamination.
Heparanase Activity
Tumor cell lysates prepared from 1 x 106 cells by three cycles of freezing and thawing in reaction buffer (20 mM phosphatecitrate buffer containing 1 mM dithiothreitol, 1 mM CaCl2, and 50 mM NaCl) were incubated (5 hours, 37 °C [pH 6.6]) with 35S-labeled ECM, prepared as described (37). The incubation medium was centrifuged (20 000g, 4 °C, 15 minutes), and the supernatant containing 35S-labeled heparan sulfate degradation fragments was analyzed by gel filtration on a Sepharose CL-6B column (0.9 x 30 cm). Fractions (0.2 mL) were eluted with phosphate-buffered saline (PBS), and the amount of radioactivity in each fraction was counted in a beta scintillation counter (7,12,30). Each experiment was performed at least three times, and the variation in elution positions (Kav values) did not exceed 15%. Reaction buffer with or without recombinant human heparanase (1 ng/mL) was routinely used as a positive or negative control, respectively.
Immunohistochemistry
MDA-MB-435 cells transfected with pHpaRz2 or pContRz were seeded on round glass coverslips in 4-well plates. After 24 hours, the cells were washed twice with PBS and fixed with chilled (20 °C) 100% methanol for 3 minutes. The cells were then washed five times with PBS, and the intrinsic fluorescence was blocked by treating the cells with 50 mM NH4Cl for 5 minutes at room temperature. The cells were then washed three times with PBS, incubated in PBS with 5% goat serum and 0.1% Triton X-100 for 30 minutes at room temperature (24 °C), and washed twice with PBS. Slides were incubated for 2 hours at 24 °C with polyclonal anti-heparanase antibodies (10 µg/mL pAb 733 in PBS with 5% goat serum) (38), washed five times with PBS, and incubated with Cy-3-conjugated goat anti-rabbit immunoglobulin G (1 : 100 diluted in PBS; Jackson, Bar Harbor, ME) for 1 hour at 24 °C. Slides were then washed eight times with PBS, mounted with 90% glycerol in PBS, and examined microscopically with a Zeiss LSM 410 confocal microscope (32,36).
Tissue sections for immunostaining were obtained from formalin-fixed and paraffin-embedded primary tumors produced in mice by subcutaneously injected cHpaEb lymphoma cells transfected with pHpaRz2 or pContRz vectors. Immunostaining was performed with anti-human heparanase monoclonal antibody (mAb 130, kindly provided by InSight Biopharmaceuticals, Rehovot, Israel) and horseradish peroxidaseconjugated goat anti-mouse antibodies (Histostatin-PLUS Bulk Kit; Zymed Laboratories, San Francisco, CA), as described (7). Vascular endothelial cells in tumor tissue sections were stained with anti-Von Willebrand polyclonal antibodies (Dako, Golstrup, Denmark) as described (39). Negative controls without addition of primary antibody showed low background staining in all cases.
Proliferation Assay
MDA-MB-435 breast carcinoma cells (5 x 105 cells per 100-mm dish) and cHpaEb lymphoma cells (2 x 105 cells per 75-mL flask) that were stably transfected with pHpaRz2 or control vectors and B16-BL6 melanoma cells (3 x 105 cells per 100-mm dish) that were transiently transfected with anti-heparanase pSi2 or control vectors were seeded in complete media in triplicate. The cells were harvested and counted in triplicate with a Coulter counter (Coulter Electronics, Luton, U.K.) every other day for 7 days, as described (22). The breast carcinoma and melanoma cells were dissociated into single-cell suspensions with trypsin and EDTA before counting.
Matrigel Invasion Assay
Tumor cells (3 x 105 cells in 1 mL of DMEM containing 0.1% BSA) were assayed in triplicate for Matrigel invasion at 37 °C in a humidified incubator (95% air, 5% CO2) for 6 hours, by using Boyden blind-well chemotaxis chambers (NeuroProbe, Gaithersburg, MD) and polycarbonate filters (13 mm in diameter, 8-µm pore size; Costar Scientific, Cambridge, MA) coated with Matrigel, as described (40,41). Medium conditioned by NIH3T3 fibroblasts was used as a chemoattractant and placed in the lower compartment of the Boyden chamber (40). For a negative control, serum-free DMEM containing 0.1% BSA was placed in the lower compartment of the Boyden chamber instead of the chemoattractant. After 6 hours, the filters were removed and the cells on the lower surface of the filter were stained with reagents from the Diff-Quik kit (American Scientific Products, McGaw Park, IL). The total number of cells from five microscopic fields selected at random (x100 magnification) was counted for each well.
Invasiveness of the Eb lymphoma cells was determined by using a modified invasion assay. Because staining Eb cells with the reagents from the Diff-Quik kit is not effective, the cells were first labeled with [3H]thymidine (1 x 106 cells, 48 hours, 37 °C, 1 µCi/mL [3H]thymidine, Amersham, Buckinghamshire, U.K.). Cell invasion was performed as described above and quantified by counting triplicate Matrigel-coated filters in a beta scintillation counter (35). Before the radioactivity was counted, cells from the upper side of the filter were removed with a cotton swab. Each experiment was repeated three times.
Cell Adhesion
Eb cells were grown (1 x 106 cells per milliliter, 48 hours, 37 °C) in RPMI-1640 medium supplemented with 10% FCS (complete medium) in the presence of [3H]thymidine (1 µCi/mL). The labeled cells were washed three times in complete medium to remove unincorporated thymidine, added to ECM-coated wells (1 x 106 cells per well in four-well plates) in triplicate, and incubated at 37 °C in complete medium (pH 7.2) for 15 minutes (35). In some experiments, adhesion of transfected cHpaEb cells to dishes coated with a naturally produced ECM, prepared as described in (37), was tested. After the incubation, the wells were washed three times with serum-free medium and the remaining firmly adhered cells were lysed in 0.2 M NaOH for 2 hours at 37 °C. The lysates were counted in a beta scintillation counter. Each experiment was repeated three times.
Experimental and Spontaneous Metastasis
For the experimental metastasis studies, the lateral tail vein of 6-week-old male C57BL/6 mice was injected with 0.4 mL of a cell suspension containing 0.4 x 106 B16-BL6 melanoma cells transiently transfected with anti-heparanase siRNA-expressing vector (pSi2) or empty pSUPER vector (34). Fifteen days after cell injection, mice were killed and their lungs were removed, fixed in Bouin's solution, and scored under a dissecting microscope for the number of metastatic nodules on the lung surface (12). Five mice were used per group, and three independent experiments were performed.
For the spontaneous metastasis studies, 2-month-old male CD1 nude mice (10 mice per group) were injected subcutaneously in the lower back with 1 x 106 Eb lymphoma cells expressing the secreted chimeric heparanase protein (cHpaEb) (36) or Eb lymphoma cells stably transfected with either anti-heparanase ribozyme (pHpaRz2) or control ribozyme (pContRz) expression plasmids. Five mice from each group were killed on day 11 (at this time point, live mice from the control group were still available) and examined for primary tumor size, vascularity, and liver metastasis (36). Metastatic colonization of the liver was evaluated by gross examination, wet weight measurements, and microscopic inspection of tissue sections. The remaining five mice from each group were monitored for survival. All animal experiments were approved by the Animal Care Committee of the Hebrew University, Jerusalem, Israel.
Statistical Analysis
InStat statistical software, version 3.05 (GraphPad Software, San Diego, CA), was used to calculate statistical differences between groups (tested by the unpaired Student's t test). All P values were two-sided.
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RESULTS |
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We produced six different hammerhead ribozymes (HpaRz 16) by in vitro transcription using double-stranded DNA oligonucleotides containing the T7 promoter, the conserved catalytic domain, and two flanking sequences designed to recognize specific motifs along the human heparanase mRNA, as shown for HpaRz2 (Fig. 1, A). To associate with and cleave its specific target, the hammerhead ribozyme must fold into a typical three-dimensional structure, with the folded catalytic core domain connected to the flanking complementary sequences, as shown for HpaRz2 in Fig. 1, B.
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Inhibition of Heparanase Activity, Cell Invasion, and Adhesion In Vitro Associated With Stable Expression of Anti-Heparanase Ribozyme
We constructed a vector to constitutively express the anti-heparanase ribozyme HpaRz2 (pHpaRz2) and tested its ability to inhibit the synthesis of endogenous heparanase. We stably transfected MDA-MB-435 breast carcinoma cells, which are known to express high levels of heparanase (7), with the pHpaRz2 vector and assayed the transfected cells for heparanase mRNA expression and enzymatic activity (Fig. 2, A). To control for possible effects of the hammerhead unit itself, MDA-MB-435 cells were transfected with the pContRz vector, which encodes for a ribozyme with an identical catalytic core but that lacks the specific heparanase mRNA recognition motifs (Fig. 2, A). Compared with MDA-MB-435 cells stably transfected with pContRz, MDA-MB-435 cells stably transfected with pHpaRz2 displayed a marked decrease in heparanase mRNA levels when evaluated by reverse transcriptionPCR (RT-PCR) (Fig. 2, A, inset) and densitometric analysis (data not shown) and showed no heparanase enzymatic activity (Fig. 2, A). In addition, immunofluorescent staining with anti-heparanase antibodies revealed that MDA-MB-435 cells stably transfected with pHpaRz2 had a marked decrease in heparanase protein levels compared with MDA-MB-435 cells stably transfected with pContRz (Fig. 2, B). There was no difference in the rate of proliferation in vitro between cells transfected with pHpaRz2 and cells transfected with pContRz (data not shown).
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Next, we used Eb lymphoma cells, which lack endogenous heparanase activity (30). Eb lymphoma cells were engineered to express a readily secreted chimeric form of heparanase (cHpa), composed of the human enzyme and the chicken heparanase signal sequence (32). We previously demonstrated that Eb cells expressing cHpa (cHpaEb) degraded heparan sulfate in ECM to a much greater extent than non-transfected Eb cells or cells transfected with the non-secreted human enzyme (32). Moreover, cHpaEb cells had increased invasiveness and metastatic potential in mice (36) and are therefore a useful experimental model for the study of heparanase involvement in tumor progression. In this study, cHpaEb cells stably transfected with pHpaRz2 or pContRz were tested for heparanase activity (Fig. 3, A) and their ability to invade Matrigel in vitro (Fig. 3, B). Compared with cHpaEb cells stably transfected with pContRz, cHpaEb cells stably transfected with pHpaRz2 had a marked decrease of more than 70% in heparanase-mediated degradation of heparan sulfate, demonstrating that the secreted form of heparanase was efficiently targeted by the ribozyme. Compared with cHpaEb cells stably transfected with pContRz, the amount of radioactivity associated with cHpaEb cells stably transfected with pHpaRz2 that invaded the Matrigel was statistically significantly reduced by 54% from 112.2 x 103 cpm (95% CI = 108.6 x 103 to 115.8 x 103 cpm) to 50.9 x 103 cpm (95% CI = 49.1 x 103 to 52.7 x 103 cpm; P<.001) (Fig. 3, B). There was no difference in the rate of proliferation in vitro between cHpaEb cells transfected with pHpaRz2 and cHpaEb cells transfected with pContRz (data not shown).
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Effect of Anti-Heparanase Ribozyme on Lymphoma Primary Tumor Vascularization, Metastasis, and Mortality of Tumor-Bearing Mice
We next investigated the effect of ribozyme-mediated heparanase gene silencing on the metastatic potential of cHpaEb lymphoma cells stably transfected with either pHpaRz2 or pContRz injected subcutaneously into nude mice. The mice were monitored for survival time and liver metastases. All mice injected with cells transfected with pHpaRz2 survived during the first 3 weeks of the experiment (Fig. 4, A). By contrast, all mice injected with cells transfected with pContRz, the control inactive ribozyme, were dead by day 14 (Fig. 4, A). On day 11, livers of five randomly selected mice from each group were removed, weighed, and processed for histological examination (Fig. 4, B). Gross macroscopic examination of the livers revealed numerous lymphoma metastases in all of the mice injected with pContRz-transfected cells but few or no visible metastatic nodules in the livers of mice injected with pHpaRz2-transfected cells (Fig. 4, B, top). The liver weights of mice injected with pContRz-transfected cHpaEb cells (mean weight = 4.7 g, 95% CI = 4.5 to 5.9 g) were statistically significantly higher than the liver weights of mice injected with pHpaRz2-transfected cHpaEb cells (mean weight = 2.0 g, 95% CI = 1.9 to 2.1 g; P<.001) (Fig. 4, B, middle). In fact, the liver weight of mice injected with pHpaRz2-transfected cHpaEb cells was similar to that of control mice (mean weight = 1.7 g, 95% CI = 1.5 to 1.7 g). Microscopic examination of liver tissue sections revealed a massive infiltration of the liver by pContRz-transfected cells but little or no liver infiltration by pHpaRz2-transfected cells (Fig. 4, B, bottom).
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To ensure that expression of the HpaRz2 and control ribozymes from the pHpaRz2 and pContRz vectors in the lymphoma cells was preserved throughout the in vivo experiment, total RNA was extracted from subcutaneous tumors produced by pHpaRz2- and pContRz-transfected cHpaEb cells. The tumors were excised 2 weeks after cell injection, and the presence of the hygromycin-resistance gene mRNA, equally expressed by both vectors, was demonstrated by RT-PCR (data not shown). Immunostaining of the tumor tissue with a monoclonal antibody (mAb 130) directed against human heparanase revealed intense staining of lymphoma cells in sections derived from tumors produced by cells transfected with pContRz but very weak or no staining in tumors produced by cells transfected with pHpaRz2 (Fig. 4, D). This result clearly demonstrates that heparanase gene silencing in pHpaRz2-transfected cHpaEb cells persisted throughout the duration of the in vivo experiment.
siRNA-Mediated Heparanase Silencing and Invasive and Metastatic Potential of B16-BL6 Melanoma Cells
To further elucidate the direct involvement of heparanase in tumor progression and the effectiveness of endogenous heparanase gene silencing, we applied the siRNA targeting approach, using highly metastatic B16-BL6 mouse melanoma cells, which contain high levels of endogenous heparanase (12,22). Two siRNA variants (Si1 and Si2) targeting two different regions of the mouse heparanase mRNA were designed and cloned into the pSUPER plasmid (34) to generate pSi1 and pSi2 expression vectors.
B16-BL6 mouse melanoma cells were transiently transfected with pSi1, pSi2, or empty pSUPER (mock) vectors by electroporation, and the cells were tested for heparanase expression and activity 48 hours later. Cells transfected with pSi1 or pSi2 contained 70%80% less heparanase mRNA than mock-transfected cells, as determined by semiquantitative RT-PCR (Fig. 5, A). Heparanase enzymatic activity, measured in cell lysates, was markedly lower (60% and 75%, respectively) in lysates from pSi1- and pSi2-transfected cells than in lysates from mock-transfected cells, further demonstrating effective silencing of the heparanase gene by heparanase-targeted siRNA (Fig. 5, B). The duration of siRNA-mediated heparanase silencing was determined by measuring heparanase enzymatic activity in B16-BL6 cells at various time points after electroporation with the anti-heparanase pSi2 vector. Maximal effect of siRNA silencing was observed 48 hours after electroporation, and it declined thereafter. The difference in levels of heparanase activity between mock-transfected cells and siRNA-transfected cells was less pronounced by 72 hours and even more so by 96 hours (nearly a 50% decrease in heparanase enzymatic activity in siRNA- versus mock-transfected cells). By 89 days after transfection, there was no detectable difference in heparanase activity between siRNA- and mock-transfected cells (data not shown).
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Finally, we tested the effect of siRNA-mediated heparanase silencing on experimental metastasis in vivo. For this purpose, B16-BL6 cells were electroporated with pSi2 or the empty vector. After 48 hours, when silencing of the heparanase gene had reached its maximum, the cells were injected into the tail vein of C57BL/6 mice (0.4 x 106 cells per mouse). In this experimental metastasis model, the invasive properties of the tumor cells are critical for their extravasation, primarily during the first 3 hours after the tumor cells enter into the circulation (12,4244). Therefore, the transient, noncontinuous silencing of heparanase in B16-BL6 cells allows for the identification of parameters directly involved in blood-borne tumor dissemination, such as the contribution of heparanase to cell extravasation as opposed to its possible effects on subsequent survival of the melanoma colonies in the target organ, stromal support, and secondary metastasis. Fifteen days after injection, the mice were killed, and their lungs were evaluated for the number of surface metastatic colonies. B16-BL6 melanoma cells transiently transfected with pSi2 established statistically significantly fewer metastatic colonies (mean number of colonies = 16.0, 95% CI = 13.4 to 18.6) than B16-BL6 melanoma cells transiently transfected with the empty vector (mean number of colonies = 144.3, 95% CI = 95.1 to 193.5; P<.001) (Fig. 5, D).
To ensure that the observed in vitro and in vivo effects were not a result of differences in the proliferative ability of B16-BL6 melanoma cells transfected with the pSi2 and control vectors, we compared the rate of proliferation of the two cell types. Mock- and pSi2-transfected B16-BL6 cells had the same rate of proliferation in vitro (Fig. 5, E). Thus, these data clearly demonstrate that specific silencing of endogenous heparanase gene expression effectively inhibits the invasive and metastatic potential of B16-BL6 melanoma cells.
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DISCUSSION |
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Various polyanionic compounds capable of inhibiting heparanase enzymatic activity, such as heparin, laminaran sulfate (22), and maltohexaose sulfate (23), have antitumor and antimetastatic activities. However, because of the multiple biologic activities of these compounds (24,25), the mechanism of their antitumor activity and its relation to heparanase inhibition are not straightforward. Moreover, the pleiotropic interactions of these compounds with the ECM and the cell surface might produce nonspecific and undesirable effects (24,25). Similarly, studies on the involvement of heparanase in cancer progression are hampered by the lack of effective neutralizing anti-heparanase antibodies. Recently, an attempt to use a more specific antisense approach has been reported (54), although in that animal model, the tumor cells were injected intrathoracically, which is not typical for metastasis studies. Hence, cells injected intrathoracically are not required to undergo intravasation or extravasation and do not encounter extracellular barriers to invade, such as the subendothelial basement membrane. In addition, among the currently available gene-silencing methods, the antisense approach is regarded as being of limited value because of its known sequence-independent effects and limited efficacy (55).
In this study, we used two powerful gene-silencing strategies (ribozyme and siRNA) to functionally inactivate the heparanase gene in diverse cellular and animal tumor models. Ribozyme targeting led to a marked inhibition of in vitro invasive and adhesive potentials of cells that either naturally express increased levels of the endogenous enzyme (i.e., MDA-MB-435 human breast carcinoma) (7) or were genetically engineered to overexpress the human heparanase gene (C-6 glioma, Eb lymphoma) (36). The anti-heparanase ribozyme statistically significantly inhibited the spontaneous metastases of cHpaEb cells to the liver. This effect was reflected in increased survival of mice injected with cHpaEb lymphoma cells expressing the anti-heparanase ribozyme relative to that of mice injected with cells expressing a control ribozyme. Moreover, vascularization of cHpaEb primary tumors was also markedly suppressed in cells that were transfected with the anti-heparanase ribozyme. The observed anti-angiogenic effect of the anti-heparanase ribozyme is primarily the result of an indirect effect of heparanase silencing on the bioavailability of angiogenesis-promoting factors that are sequestered by heparan sulfate in the tumor microenvironment and can be released by the tumor-derived heparanase (13,19).
The biologic and therapeutic relevance of the heparanase-silencing approach was further validated using the highly metastatic B16-BL6 mouse melanoma cells (12) transfected with mouse heparanase-specific siRNA. Knockdown of heparanase expression resulted in an almost complete inhibition (80%) of lung colonization after intravenous injection of siRNA-transfected B16-BL6 cells compared with mock-transfected cells. Previous studies (12,4244) clearly established that in the B16-BL6 experimental metastasis model, invasive properties of the tumor cells are critical for cell extravasation, primarily during the first few hours after the tumor cells enter into the bloodstream. Based on the kinetics of heparanase siRNA silencing in transiently transfected B16-BL6 melanoma cells cultured in vitro, the in vivo experiments were designed to have maximal heparanase silencing occur during the first several hours after cells were injected into the bloodstream. Transient, noncontinuous silencing of heparanase in B16-BL6 cells restricted the suppression of heparanase activity to the critical window of blood-borne cell extravasation, followed by release of the inhibitory effect during the subsequent formation of metastatic colonies in the lungs. Thus, we have specifically analyzed the role of heparanase in cell extravasation rather than in the multiple subsequent events, such as cell proliferation and establishment and survival of colonies in the target organ and angiogenesis and formation of secondary metastases.
To our knowledge, the data described here represent the first successful application of ribozyme- and siRNA-mediated gene silencing to effectively reduce the levels of heparanase. Our results highlight a critical role of heparanase in tumor angiogenesis, growth, and metastasis. In addition to demonstrating the potential promise for cancer treatment, the specific heparanase gene-silencing tools clarify the molecular and cellular mechanisms underlying recently described heparanase-mediated processes such as cell adhesion (35,56) and survival signals (56,57) in vitro, as well as tissue repair, hair growth (39), and bone formation in vivo (Zcharia E, Kram V, Yacobi-Zeevi Y, Bab I, Vlodavsky I: unpublished data). These tools, which target RNA, are especially important in light of the recently discovered nonenzymatic functions of heparanase (35,56)functions that are not sensitive to the currently available heparanase inhibitors (22,23). For example, we found that heparanase-mediated cell adhesion was markedly inhibited in HpaRz-transfected lymphoma cells but not by laminaran sulfate, which efficiently inhibits heparanase enzymatic activity (12,22). Heparanase gene silencing may thus provide novel mechanistic insights into the functions of heparanase (i.e., cell adhesion and signal transduction) unobtainable by other methods of inhibiting heparanase enzymatic activity.
Further investigation is required to identify the most effective delivery strategies for therapy using ribozyme and siRNA gene-silencing approaches. Experiments are underway to test the feasibility of electroporating heparanase-targeted siRNA-expressing vectors directly into the primary tumor in vivo (58). Of particular promise is the development of lentivirus-based vectors that enable sustained expression of heparanase-silencing agents in the target tumor cells (59). Such a vector has already been constructed and will be administered systemically or locally by electroporation into the tumor tissue. The ability of a lentivirus to infect both dividing and nondividing cells and to allow for long-term multilineage gene expression (60) will extend the range of cells and tumor types in which the heparanase-silencing approach may be practical.
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We thank Dr. Yaron Tikochinski (Intelegene) and Dr. Helena Aingorn (Department of Oncology, Hadassah-University Hospital) for their excellent advice and continuous support.
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Manuscript received January 25, 2004; revised June 3, 2004; accepted June 16, 2004.
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