RhoC is essential for angiogenesis induced by hepatocellular carcinoma cells via regulation of endothelial cell organization

Cell culture and transfection.  HMEC‐1, a human dermal microvascular endothelial cell line, was purchased from the American Type Culture Collection (Rockville, MD, USA). HCCLM3, a HCC cell line with high metastatic potential, was obtained from the Liver Cancer Institute and Zhong Shan Hospital of Fudan University, Shanghai, China. The PT67 retroviral packaging cell line was purchased from BD Biosciences Clontech (Palo Alto, CA, USA). The HCCLM3 and PT67 cell lines were cultured in high‐glucose Dulbecco's modified eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and incubated in 5% CO₂ at 37°C, whereas the HMEC‐1 cell line was cultured in low‐glucose DMEM supplemented with 10% FBS and incubated under the same conditions. For transfection of packaging cells, the PT67 retroviral packaging cells, which express the 10A1 viral envelope for production of amphotropic virus, were transfected with the relevant reconstructed pSUPER.retro.neo retroviral expression plasmid at 70% confluence with the use of Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. Clones expressing retrovirus at 1 × 10⁵ colony‐forming units (cfu)/mL were selected for subsequent experiments. Multiple rounds of infection, which can increase the number of infected cells as well as increase the number of copies per cell, were carried out during the transfection of HMEC‐1 cells. Briefly, media containing retrovirus was added to the cells supplied with Polybrene (Sigma‐Aldrich, St Louis, MO, USA) (8 µg/mL). After 24 h, the medium was replaced with fresh medium and the retrovirus was added again. Infecting the cells three times resulted in an 80–90% transfection efficiency.

Generation of siRNA constructs.  Reconstructed pSUPER.retro.neo retroviral expression plasmid, containing siRNA directed against human RhoC plasmid, was prepared previously in our laboratory. Briefly, the 60‐bp oligonucleotides, containing the unique 19‐bp RhoC siRNA sequence in both sense and antisense orientations separated by a 9‐bp spacer sequence, were synthesized in the forward and reverse orientation, annealed, and ligated into the pSUPER.Retro.neo vector excised by the HandIII and BglII incision enzymes. The presence of the insert was determined by sequencing. The effective sequence and control sequence were as follows: effective sequence sense, 5′‐GATCCCCCATGTGCTTCTCCATCGACTTCAAGAGAGTCGATGGAGAAGCACATGTTTTTA‐3′; effective sequence antisense, 5′‐AGCTTAAAAACATGTGCTTCTCCATCGACTCTCTTGAAGTCGATGGAGAAGCACATGGGG‐3′; control sequence sense, 5′‐GATCCCCCAGTCAGGAGGATCCAAAGTTCAAGAGACTTTGGATCCTCCTGACTGTTTTTA‐3′; and control sequence antisense, 5′‐AGCTTAAAAACAGTCAGGAGGATCCAAAGTCTCTTGAACTTTGGATCCTCCTGACTGGGG‐3′.

Western blotting.  Cells were lysed in a lysis buffer (Pierce, Rockford, IL, USA). Extract equivalent to 100 µg total protein was separated on 12% polyacrylamide gels and transferred to polyvinylidene fluoride membrane (Invitrogen). The membrane was incubated with primary antibody (goat anti‐RhoC antibody, diluted at 1:500; goat anti‐VEGF antibody, diluted at 1:2000), followed by incubation with a 1:1000 dilution of horseradish peroxidase‐linked rabbit antigoat antibody (Santa Cruz Biotechnology). The membrane was then washed and treated with western blotting luminal reagent (Pierce) to visualize the bands.

Invasion and migration assays.  For the invasion assay, a transwell with an 8‐µm diameter pore membrane (Costar, Cambridge, MA, USA) was coated with 200 µL matrigel at 200 µg/mL (Becton‐Dickinson) and incubated overnight. Twenty thousand HMEC‐1 cells were seeded into the upper chamber of the transwell and the lower chamber was filled with 0.8 mL low‐glucose DMEM supplemented with VEGF (10 ng/mL). After 24 h of incubation at 37°C, the cells were fixed in methanol and stained with hematoxylin–eosin and the cells that invaded through the pores to the lower surface of the filter were counted under a microscope. Three invasion chambers were used per condition. The values obtained were calculated by averaging the total number of cells from three filters. For the migration assay, a wound‐healing assay was carried out. HMEC‐1 cells (1 × 10⁶) were seeded on 6‐cm plates coated with 10 µg/mL type I collagen. The cells were incubated for 24 h before the monolayer was disrupted with a cell scraper (1.2‐mm width). Photographs were taken at 0 and 24 h using a phase‐contrast microscope (ELWD 0.3; Nikon, Tokyo, Japan). Experiments were carried out in triplicate, and four fields were recorded for each point.

FACS analysis.  For fluorescence‐activated cell sorter (FACS) analysis, after incubation in serum‐free media for 24 h, cells were washed twice with phosphate‐buffered saline at 4°C, and the concentration was adjusted to 1 × 10⁶/mL. Annexin V/Cy5 and propidium iodide (PI) were added and incubated for 30 min. FACS analysis was carried out using a FACS caliber instrument (Becton Dickinson). Experiments were repeated three times with similar results.

Immunohistochemistry and immunofluorescence staining.  For im‐munohistochemistry, briefly, tissue sections of 4 µm in thickness were cut and baked at 60°C overnight. After heat‐induced antigen retrieval in ethylenediaminetetraacetic acid buffer, the sections were incubated at 37°C for 1 h with specific primary antibodies used at a 1:50 dilution, followed by incubation with the secondary antibody at 37°C for 45 min. The sections were then visualized by applying 3,3‐diaminobenzidine tetrahydrochloride. A negative control omitted the primary antibody, whereas overexpression of the target protein confirmed by western blotting was used as a positive control. Diffuse, moderate to strong cytoplasmic staining characterized positive expression, whereas negative expression was devoid of any cytoplasmic staining or contained faint, equivocal staining. For immunofluorescence, cells were grown on matrigel in DMEM containing 10% FBS for 18 h, then F‐actin filaments were visualized in cells using rhodamine‐conjugated phalloidin (1:40 dilution in phosphate‐buffered saline) for 20 min at room temperature.

In vivo.  A RhoC‐RNAi (RNA interference) retroviral gene‐delivery BALB/c nude mouse model was established with important modifications by including the PT67 retroviral packaging cells as a constant source of retrovirus.^{( 17} , ^{18 )} Briefly, after transfecting the reconstructed pSUPER.retro.neo retroviral expression plasmid into the PT67 packaging cell line, stable virus‐producing PT67 cells, named PT67^RhoCRNAi+ and PT67^RhoCRNAi– (control group), respectively, were produced and clones expressing retrovirus at 1 × 10⁵ cfu/mL were selected. After 1 week, a total of 1 × 10⁷ HCCLM3 cells in 0.2 mL DMEM were transplanted subcutaneously into the right flank of BALB/c nude mice, and 5 × 10⁶ retroviral packaging cells (PT67^RhoCRNAi+ or PT67^RhoCRNAi–), with a high titer of approximately 10⁵, were inoculated subcutaneously into the left flank as a constant source of retrovirus. Mice were killed 45 days after implantation and the tumors were removed and fixed in 10% formalin, embedded in paraffin and cut into 4 µm‐thick slices for standard histopathological study. Paraffin sections embedded with the subcutaneous tumor were stained with hematoxylin–eosin for histological examination and stained with anti‐green fluorescent protein (GFP) antibody or anti‐RhoC antibody for evaluating infection efficiency or suppression efficiency in vivo. For evaluating microvessel density (MVD), paraffin sections were stained with antimouse CD31 antibody, then three areas with the most vessels within each section were selected for quantitation of angiogenesis, and vessels labeled with the antimouse CD31 antibody were counted under a light microscope at ×200 magnification. The histopathological examination was carried out by a pathologist who was masked from the experimental group designation. For the angiogenesis assay in vivo, based on the RhoC‐RNAi retroviral gene‐delivery BALB/c nude mice model, we developed a neovascularization model in vivo, in which PT67 packing cells were inoculated subcutaneously as a constant source of retrovirus, and matrigel mixed with VEGF was inoculated subcutaneously to drive angiogenesis. One week after implantation, matrigel (including skin) was dissected and photographed under a visible‐light microscope (×20). New blood vessels on the matrigel were counted and their lengths were determined.^{( 19 )} Sterile small‐animal surgical techniques were followed during the whole procedure. All experimental protocols were approved by the Institutional Animal Care and Use Committee, Xiangya Hospital, Central South University.

In vitro angiogenesis: endothelial cell network formation assays.  To analyze in vitro angiogenesis, a 24‐well tissue culture plate was coated with 0.2 mL matrigel (8.8 mg/mL). HMEC‐1 cells (1 × 10⁵ cells/mL) were seeded on matrigel and incubated at 37°C in a 5% CO₂ humidified atmosphere for 18 h. The network formation was observed under an inverted phase‐contrast microscope (Nikon), and the images were captured with a video graphic system (Retiga 1300 Digital Output Camera; QIMAGING, Surrey, BC, Canada). The number of network projections in 10 × 40 fields was counted for three independent experiments in each group, and the data were obtained using the following formula:

where a = the number of branch points per ×40 field and b = the total number of branches per point.^{( 19 )}

In vitro angiogenesis: endothelial cell sprout formation assays.  HMEC‐1 cells were mixed with Cytodex‐3 microcarriers (Amersham Pharmacia Biotech) at a concentration of 300 HMEC‐1 cells per bead in 1 mL DMEM medium. Beads with cells were shaken gently every 20 min for 4 h at 37°C and 5% CO₂. After incubation, beads with cells were transferred to a six‐well tissue culture plate (BD Biosciences, Bedford, MA, USA) and left for 12–16 h in 1.5 mL DMEM at 37°C and 5% CO₂. The following day, beads with cells were washed three times and resuspended at a concentration of 200 cell‐coated beads/mL in 4 mg/mL matrigel. Next, 0.8 mL matrigel mixed with cell‐coated beads was add to one well of a 24‐well tissue culture plate and culture for 1 week. The sprouting of HMEC‐1 cell was count as described previously.^{( 20 )} Briefly, high‐resolution images of beads were captured on an IX70 microscope (Olympus, Tokyo, Japan) with a ×4 objective. Capturing images at low magnification has the advantage of a high depth of field, which enables inclusion of all sprouts in focus. Images were then magnified in Adobe Photoshop (Adobe Systems, San Jose, CA, USA) and analyzed using ImageJ (National Institutes of Health, Bethesda, MD, USA). The number of sprouts per bead was determined, where a sprout was defined as a vessel of length equal to the diameter of a bead. For statistical analysis, the number of sprouts/10 beans was counted and 100 beads were assessed for each condition.

Statistical analysis.  Statistical analysis was carried out using SPSS software (version 11.0; SPSS, Chicago, IL, USA). The results of the quantitative data in the present study are expressed as mean ± SD. Statistical differences between groups were compared using two‐tailed ANOVA and t‐test. A P‐value of less than 0.05 was considered significant.

RhoC is essential for angiogenesis induced by VEGF.  Expression of VEGF in the HCC tumor was downregulated in mice^RhoCRNAi+, indicating that RhoC regulates angiogenesis probably through downregulation of VEGF expression in HCC tumors. However, it is not known whether there is another pathway through which RhoC can regulate angiogenesis. Therefore, based on the RhoC‐RNAi retroviral gene‐delivery BALB/c nude mouse model, we developed a neovascularization model in vivo, in which PT67 packing cells were inoculated subcutaneously as a constant source of retrovirus and matrigel mixed with VEGF was inoculated subcutaneously to drive angiogenesis (Fig. 2a,b). As anticipated, neovascularization was markedly reduced in RhoC‐deficient mice. The total length of angiogenesis in matrigel was 24532 ± 6783 µm and 34869 ± 4396 µm in mice^RhoCRNAi+ and mice^RhoCRNAi–, respectively (P < 0.05) (Fig. 2c–e), indicating that RhoC critically regulates angiogenesis driven by VEGF in vivo. Meanwhile, this result also indicates that RhoC regulation of angiogenesis induced by HCC cells did not exclusively depend on downregulation of VEGF expression.

RhoC participates in invasion and migration of HMEC‐1 cells.  We hypothesized that RhoC plays a critical role in the migration of endothelial cells, which may be responsible for regulation of angiogenesis. We therefore investigated the effect of RhoC on the invasion and migration of HMEC‐1 cells. First, the siRNA approach combining multiple rounds of infection was used to inhibit RhoC expression in HMEC‐1 cells, which resulted in ~80–90% transfection efficiency. The expression of RhoC protein in HMEC‐1^RhoCRNAi+ and HMEC‐1^RhoCRNAi– was 0.33 ± 0.05 and 0.84 ± 0.07, respectively (P < 0.05) (Fig. 3a), suggesting successful inhibition of RhoC in HMEC‐1 cells. Next, a wound‐healing assay was used to determine the migration of HMEC‐1 cells. HMEC‐1^RhoCRNAi+ decreased wound closure by 23% compared with HMEC‐1^RhoCRNAi–, suggesting a role for RhoC in the migration of HMEC‐1 cells (P < 0.05) (Fig. 3b). Meanwhile, a matrigel invasion assay in transwell culture chambers was carried out to determine the effect of RhoC on the in vitro invasion of HMEC‐1 cells. The number of HMEC‐1^RhoCRNAi+ cells that passed through the matrigel was 49 ± 6 compared with 70 ± 3 HMEC‐1^RhoCRNAi– cells (P < 0.05) (Fig. 3c). At the same time, apoptosis in HMEC‐1^RhoCRNAi+ and HMEC‐1^RhoCRNAi– cells under serum‐starvation conditions, determined using FACS, showed no significant difference (P > 0.05) (Fig. 3d). Together, these results support a critical role for RhoC in the invasion and migration but not apoptosis of HMEC‐1 cells.

Knockdown of RhoC inhibits angiogenesis in vitro.  As a critical role for RhoC in the migration of HMEC‐1 cells has been confirmed, we hypothesized that knockdown of RhoC may inhibit angiogenesis through inhibition of HMEC‐1 organization. Therefore, the effect of RhoC on HMEC‐1 cell organizational behavior in matrigel was investigated. As shown in Figure 4a, inhibition of RhoC in HMEC‐1 cells resulted in inhibition of capillary network formation of HMEC‐1 cells in matrigel, with the index of capillary network formation being 489 ± 215 and 2397 ± 726 in HMEC‐1^RhoCRNAi+ and HMEC‐1^RhoCRNAi–, respectively (P < 0.05). Meanwhile, with the help of rhodamine‐conjugated phalloidin, F‐actin filaments of HMEC‐1 cells in matrigel were visualized, as shown in Figure 4b. Reorganized HMEC‐1 cells had more F‐actin filaments than HMEC‐1 cells without reorganization, indicating that RhoC regulates endothelial cell organization through reorganization of the F‐actin filaments. To further confirm the critical role of RhoC in angiogenesis, HMEC‐1 cells were coated on Cytodex‐3 microcarriers and cultured in a three‐dimensional angiogenesis model. As expected, knockdown of RhoC inhibited tube formation of HMEC‐1 in three‐dimensional matrigel, with the number of sprouts per 10 beads being 9 ± 7 and 45 ± 9 in HMEC‐1^RhoCRNAi+ and HMEC‐1^RhoCRNAi–, respectively (P < 0.05) (Fig. 4c). Together, these results demonstrate that knockdown of RhoC inhibits angiogenesis in vitro.