• antiangiogenesis;
  • Tripterygium wilfordii;
  • triptolide;
  • Tie2;
  • VEGFR-2


  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Triptolide is a key anti-inflammatory compound of the Chinese herbal medicine Tripterygium wilfordii Hook. f. (Celastraceae). It also possesses potent antitumor activity. In this study, we show that triptolide is an angiogenesis inhibitor based on various angiogenesis assays. The IC50 in in vitro assays was 45 nM, which was much lower than the plasma concentrations of triptolide in the rat or human administered with T. wilfordii extracts for treating inflammation. When dosed in vivo, triptolide potently inhibited angiogenesis at 100 nM in Matrigel plug assay. Triptolide at 0.75 mg/kg/day significantly blocked tumor angiogenesis and tumor progression in murine tumorigenesis assay. The underlying mechanism of triptolide correlated with downregulation of proangiogenic Tie2 and VEGFR-2 expression in human umbilical vein endothelial cell by semiquantitative RT-PCR and western blot analysis. Although Tie2 inhibition appeared to be a later event as compared with VEGFR-2, Tie2 overexpression significantly attenuated the inhibitory effect of triptolide on endothelial proliferation and network formation. By contrast, Tie2 knockdown mimicked the inhibitory effect of triptolide on endothelial network formation. Our findings suggest that antitumor action of triptolide is partly via inhibition of tumor angiogenesis by blocking 2 endothelial receptor-mediated signaling pathways, and triptolide can be a promising antiangiogenic agent.

Tripterygium wilfordii Hook. f. (Celastraceae), also known as Thunder God Vine, is a popular herb in China for the treatment of immune-inflammatory diseases including rheumatoid arthritis, systemic lupus erythematosus, nephritis and asthma.1 Extracts from T. wilfordii has entered clinical trials for the treatment of rheumatoid arthritis.2–5 Characterization of the active components present in this plant identified triptolide, a diterpenoid triepoxide, as a key component responsible for most of the immunosuppressive, anti-inflammatory and antiproliferative effects.6 The content of triptolide was thus used as the quality control marker for most of the extracts and preparations made from T. wilfordii.7–9 Triptolide itself has also been tested in clinical trials for the treatment of psoriasis vulgaris,10 diabetic nephropathy11 and nephritic syndrome12 in China.

Triptolide was first found to have antileukemic activity in 1972.13 In addition to its potent immunosuppressive and anti-inflammatory activities, its antitumor activity has attracted much interest since then. Triptolide exerted antiproliferative and proapoptotic effects on tumor cell lines in vitro,14–16 restricted tumor growth or shrank tumor in vivo.17, 18 Furthermore, triptolide sensitized tumor cells to other anticancer agents19 and had synergistic effect with other chemotherapeutic agents in preclinical animal models.15, 18 Although tumor growth and metastasis depend upon angiogenesis, there are only preliminary studies on the effect of triptolide on angiogenesis.20, 21

Antiangiogenic therapy has been increasingly recognized as a promising therapy for cancer treatment. We have previously identified that triptolide from T. wilfordii had potent antiangiogenic activity in zebrafish angiogenic assay through targeting angpt2-tie2 signaling pathway in both time- and dose-dependent manners.22 To further confirm the antiangiogenic activity of triptolide in mammalian systems, we used human umbilical vein endothelial cells (HUVECs) and mouse Matrigel plug model to test its antiangiogenic effect in vitro and in vivo and further investigated its action mechanisms. Finally, a murine tumorigenesis model was used to validate its antitumor activity as a potent angiogenic inhibitor.

Material and Methods

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Cell culture

Primary HUVECs were obtained from fresh human umbilical veins with a slight modification as previously described23 and maintained in endothelial cell-growth medium-2 (EGM-2) (BioWittaker, Walkersville, MD). HUVECs at early passages (passages 2–7) were used in all the experiments. Mouse melanoma B16-F0 cells, human lung cancer A549 cells and human immortalized skin HaCaT keratinocytes were from American Type Culture Collection (Manassas, VA). B16 and HaCaT cells were propagated in DMEM with 10% FBS. A549 cells were maintained in MEM medium supplemented with 1 mM sodium pyruvate, 1% nonessential amino acids (NEAA), antibiotics and 10% fetal bovine serum (Life Technologies, Grand Island, NY). All cells were incubated at 37°C in an atmosphere of 95% air and 5% CO2.

Proliferation assay

Subconfluent wild-type HUVECs and Tie2 overexpressing HUVECs were used for cell proliferation assay. HUVECs (8,000 per well), A549, B16 and HaCaT (3,500 per well) were seeded in 96-well plate in the appropriate growth medium. Plates for HUVECs were precoated with 0.1% gelatin before cell seeding. At 12-h postseeding, HUVECs were starved with M199 (Life Technologies) supplemented with 0.1% bovine serum albumin (BSA, Sigma-Aldrich, St. Louris, MO) plus 1% heat-inactivated FBS and antibiotics for 12 h. A549 cells were starved with MEM supplemented with 0.1% BSA, 1% nonessential amino acids, 1 mM sodium pyruvate and antibiotics for 36 h. HaCaT cells were starved in DMEM with 0.1%BSA for 36 h. Serum-deprived cells were treated for 24 hr with growth medium containing vehicle (0.1% DMSO), triptolide (10–250 nM) or SU5416 (250 nM). Cell proliferation was measured by CellTiter 96® AQueous One Solution (Promega, Madison, WI). Each treatment was done in quadruplicates.

Migration assay

HUVEC migration assay was performed in a modified Boyden chamber (Neuroprobe, Cabin John, MD). Twenty-seven microliters of starvation medium containing VEGF-A (5 ng/ml) (Pepro Tech, London, UK) or angiopoietin 1 (Ang1) (200 ng/ml) (R&D Systems, Minneapolis, MN) together with 0.1% DMSO (vehicle) or 25–250 nM triptolide was added to the lower compartment of each chamber. Gelatin-precoated membrane was used to separate each chamber into upper and lower compartments. HUVEC suspension (4 × 104 cells per well) in M199 medium with 0.1% BSA and antibiotics was added to the upper compartments. After 9-h incubation at 37°C, the nonmigrated cells from the upper face of the membrane were removed by phosphate buffer saline-soaked cotton swab. The migrated cells on the bottom face of the membrane were fixed with methanol and stained with Giemsa solution (Merck, Germany). Each treatment was done in quadruplicates and then randomly selected fields under 100× magnification of each well were scored for migrated cells.

Network formation assay

Green fluorescent protein-positive (GFP+), wild-type HUVECs, Tie2 overexpressing HUVECs and Tie2 knockdown HUVECs (2 × 104 cells/well) in EGM-2 medium were seeded in 48-well plate precoated with Matrigel (100 μl/well, 8 mg/ml) (BD Biosciences, Bedford, MA) and exposed to triptolide (25–1,000 nM) or 0.1% DMSO for 9–12 h. The network formation was visualized and imaged under fluorescent or inverted microscope (Olympus, Tokyo, Japan) at 100 × magnification. The network length was quantified by using Image software developed by Dr. Sun YN at National Cheng Kung University.

Apoptosis assay

HUVECs in EGM-2 medium were seeded at 1.6 × 105 cells per 6-cm dish. Two days after incubation at 37°C, cells were exposed for 24 h to 0.1% DMSO, 10–250 nM triptolide or 250 nM SU5416 in EGM-2 medium. Both floating and adherent cells were harvested for detecting apoptotic cells using Annexin V-FITC kits (Bender MedSystems, Burlingame, CA). Annexin V-labeled apoptotic cells were analyzed by flow cytometry.

Genetic manipulation of Tie2 expression in HUVEC

For Tie2 overexpression, Tie2-bearing adenovirus at multiplicity of infection (MOI) of 5–40 was used to infect HUVEC as described.24 For knocking down Tie2 expression in HUVECs, 2 shRNA-bearing lentiviral vectors, respectively, targeting at the coding sequence (TRCN0000000414) or 3′-noncoding sequence (TRCN00000000412) of Tie2 mRNA were used to infect HUVECs. Lentiviral infected cells were enriched for 3 days with 2.5 μg/ml puromycin before subsequent assays. The altered expression of Tie2 protein was confirmed by western blot analysis.

Semiquantitative RT-PCR

Subconfluent HUVECs were used for this experiment. HUVECs (8 × 104) were seeded in 6-well plate. Cells were incubated for the indicated time with 5–100 nM triptolide or 0.1% DMSO at 48-h postseeding. Total RNA was extracted using single-step RNA isolation by acid-phenol quanidium thiocyanate. One microgram of total RNA was used for reverse transcription. The primers for each gene were from MDbio (Taipei, Taiwan). PCR was carried out in a volume of 20 μl using the Thermal Cycler (Takara, Japan) with an annealing temperature of 55°C for the housekeeping gene GAPDH; 54°C for VEGFR-1; 59°C for VEGFR-2, Tie1, Tie2; 60°C for VEGF-A and 64°C for Angiopoietin2 (Ang2). The cycle numbers used were validated using the same method as the report.25 Briefly, PCR products from different cycles of amplification were visualized on a UV-transilluminator after electrophoresis on 1.5% agarose gel containing ethidium bromide, and the signal intensity was quantified with the Gel-Doc 1000 system and Quantity One software (Bio-Rad). The cycle number with half-maximal amplification was used for semiquantitative analysis of gene expression, and they were 21 cycles for GAPDH; 30 cycles for Tie1; 31 cycles for VEGFR-2, VEGF-A and 32 cycles for Ang2, VEGFR-1, Tie2.

Western blot analysis

Following treatment, cells were lysed with a lysis buffer (10% glycerol, 50 mM HEPES, PH 7.0, 150 mM NaCl, 1.25% Triton X-100, 1.5 mM MgCl, 1 mM EGTA, 50 mM NaF, 10 mM sodium pyrophosphate, 2 mM sodium orthovanadate, 5 mM EDTA, 2 mM phenylmethyl sulfonyl fluoride, 10 μg/ml approtonin, 10 μg/ml leupeptin, 1 μg/ml pepstatin A; all reagents from Sigma-Aldrich) plus protease inhibitor cocktail (Roche, Switzerland). Equal amounts of total protein (30 μg) were size-fractionated by SDS–PAGE and transferred to PVDF membranes (Millipore). The blots were washed with Tris-buffered saline plus 0.1% (vol/vol) Tween-20 (TBST) and blocked with 5% nonfat milk solution at room temperature for 1 h. Then the blots were probed with specific primary antibodies for 16 h at 4°C. After extensive TBST washes, the blots were probed by appropriate secondary antibodies at room temperature for 1 h. Then the blots were washed extensively with TBST and developed by chemiluminescence detection system. All the antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA).

In vivo mouse Matrigel plug assay and immunohistochemical staining

The mouse Matrigel plug assay was performed as previously described.26 Heparin (64 U/ml) and basic fibroblast growth factor (bFGF) (100 ng/ml), dissolved in PBS; triptolide (100 and 500 nM), dissolved in DMSO (0.1%) were mixed with 200 μl Matrigel (8 mg/ml) prior to subcutaneous injections into abdominal tissues of 6-weeks-old male BALB/c mice (Laboratory Animal Center, College of medicine, National Cheng Kung University). Four to 5 mice were in each group. Ten days after injection, Matrigel plugs were removed, fixed in 10% buffered formalin, embedded in paraffin and sectioned. Sections were immunostained with CD31 (Santa Cruz Biotechnology). CD31-positive vessels in each 200 × magnification were scored and quantified.

In vivo murine tumorigenesis assay and immunohistochemical staining

Triptolide was dissolved in a mixture of 60% ethanol, 30% DMSO and 10% phosphate buffer (pH 6.0) as recommended.27 Mouse melanoma B16 cells (106 cells/injection, 6 mice/group) were injected s.c. into 8-week-old syngenic CL57/B6 mice. The next day, triptolide at 0.25, 0.5 and 0.75 mg/kg were injected i.p. into the mice on a daily basis. Three weeks postinjection, tumor tissues were harvested and weighted, then fixed with 10% formalin and paraffin-embedded. Thin sections (5 μm thick) were deparaffinized in xylene, rehydrated then treated with 3% hydrogen peroxide to block endogenous peroxidase. Before incubation with primary antibody, an antigen retrieval method was used to enhance immunodetection. The tissue sections were incubated overnight with antialpha smooth muscle actin antibody (1:300, Abcam) at 4°C and then incubated with the secondary antibody. Bound antibody was detected using an AEC chromogen substrate kit (Zymed Laboratories). The same sections were then counter-stained with 1% hematoxylin and the slide mounted with glycerol gelatin (Sigma-Aldrich). All animal studies were approved by the Institutional Animal Care and Use Committee at National Cheng Kung University.

Statistical analysis

All experiments were repeated at least 3 times. Values are given as means and standard errors of the mean (SEM). Data were analyzed using Graph Pad Prism 4.0 software. Statistical significance was assessed by Student's t-test or one-way ANOVA. p values less than 0.05 were considered significant.


  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Inhibition of HUVEC, A549, B16 and HaCaT proliferation by triptolide

Triptolide at 10–250 nM potently inhibited HUVEC proliferation in a dose-dependent manner (Fig. 1a). The IC50 of triptolide for HUVEC proliferation is 45 nM. Triptolide had higher inhibitory effect on HUVEC proliferation than SU5416 at 250 nM, a commercially available VEGFR-2 tyrosine kinase inhibitor (67% vs. 28% inhibition). Triptolide at the same dose had less inhibitory effect on A549 cell and HaCaT cell proliferation (<30% and 20%, respectively). Mouse melanoma B16 cells were resistant to triptolide treatment at all concentrations. In contrast to endothelial-specific effect of SU5416 (Fig. 1a), triptolide had higher inhibitory effect on endothelial cells than that on skin keratinocytes and cancer cells.

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Figure 1. Antiproliferative and proapoptotic effect of triptolide. (a) Inhibitory effect of triptolide on HUVECs, A549, B16 and HaCaT proliferation. (b) Proapoptotic effect of triptolide on HUVECs. Each point represents mean + SEM (n = 4) from 3 independent experiments. *p < 0.05; **p < 0.01; ***p < 0.001. The data were analyzed by one-way ANOVA followed by the Dunnett's test for comparisons of all treated groups with control group.

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Proapoptotic effects of triptolide on HUVECs

We next examined by Annexin V-FITC labeling if the antiproliferative effect of triptolide on HUVECs were through apoptosis induction. Triptolide at 10–250 nM dose-dependently induced HUVECs to undergo apoptosis (Fig. 1b). Over 25% of HUVECs died through apoptosis after triptolide treatment at 250 nM. By contrast, less than 3% of HUVECs were killed by SU5416 at the same dose. Triptolide was 10 times more potent than SU5416 in inducing apoptotic HUVEC. Together, triptolide at the same concentration manifested a higher potency than SU5416 in inducing HUVEC apoptosis.

Inhibition of HUVEC migration by triptolide

Migration is a key step required for angiogenesis. VEGF-A and Ang1 are potent chemoattractants for HUVECs and thus were used as chemoattractants in separate experiments. VEGF-A and Ang1 potently induced HUVECs migration in the control group (Figs. 2a and 2b). Triptolide dose-dependently inhibited both VEGF-A and Ang1 induced HUVEC migration, and the compound at 250 nM achieved 78% and 75% inhibition, respectively, for VEGF-A and Ang1 mediation. Consistent with VEGFR-2 being the primary receptor for VEGF-A signaling and SU5416 being a selective VEGFR-2 inhibitor, SU5416 at 1,000 nM potently inhibited VEGF-A, but not Ang1-induced HUVEC migration (Figs. 2a and 2b). Triptolide at 250 nM possessed a comparable inhibitory effect with SU5416 at 1,000 nM (Fig. 2a). These results indicate that triptolide inhibited HUVEC migration possibly through blocking both VEGF-A and Ang1-mediated signaling pathways.

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Figure 2. Inhibitory effect of triptolide on HUVECs migration mediated by either VEGF-A (5 ng/ml) (a) or Ang-1 (200 ng/ml) (b). Each point represents mean + SEM (n = 4) from 3 independent experiments. *p < 0.05; **p < 0.01; ***p < 0.001. The data were analyzed by one-way ANOVA followed by the Dunnett's test for comparisons of all treated groups with control group.

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Inhibition of HUVEC network formation by triptolide

HUVECs when seeded on Matrigel are able to become elongated and form capillary-like structure mimicking the in vivo neoangiogenesis process.28 To get high-resolution images, ectopic GFP-expressing HUVECs were used in this assay as a reporter for network forming ability before and after drug treatment. HUVECs at 9-h postseeding had peak network formation in control group (Fig. 3). SU5416 at 1,000 nM marginally inhibited network formation, whereas triptolide at 100–1,000 nM significantly attenuated network formation in a dose-dependent manner. Triptolide at 500 and 1,000 nM almost completely blocked network formation at 9-h postseeding (Fig. 3). This phenomenon provides evidence for triptolide-mediated suppression of HUVEC network formation possibly through targeting certain signaling pathway rather than nonspecific cytotoxicity. Taken together, triptolide dose-dependently inhibited HUVEC network formation in a more potent fashion than SU5416.

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Figure 3. Inhibitory effects of triptolide on GFP+-HUVECs network formation. Images were taken under fluorescent microscope after 9-h incubation. The network length in each treatment conditions was quantified in the lower panel. Each point represents mean + SEM (n = 4) from a representative experiment. *p < 0.05; **p < 0.01; ***p < 0.001. The data were analyzed by one-way ANOVA followed by the Dunnett's test for comparisons of all treated groups with control group.

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Angiogenic gene expression in HUVECs treated by triptolide

In our previous research,22 triptolide dose- and time-dependently inhibited angpt2 and tie2 expression in zebrafish angiogenic assay. To examine if the findings in zebrafish could be recapitulated in mammalian cells, we investigated by RT-PCR analysis the expression of several angiogenic genes in triptolide-treated HUVECs. VEGF-A and Ang2 together with their cognate receptors, VEGFR-1, VEGFR-2, Tie1 and Tie2, are 2 common signaling pathways involved in promoting angiogenesis. We first examined the mRNA expression of these genes in HUVECs treated for 24 h with 5–100 nM triptolide (Fig. 4a). Following 24-h treatment, triptolide at 50 nM suppressed VEGFR-2 and Tie2 mRNA expression by more than 70% compared with the control. In particular, VEGFR-2 expression was completely blocked by triptolide at 100 nM. By contrast, the mRNA expression of VEGFR-1 and Tie1 was only marginally affected by the drug. In addition to the differential effect of triptolide on receptor expression, the expression of both ligands, VEGF-A and Ang2, was significantly reduced by 100 nM triptolide.

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Figure 4. Effects of triptolide on the expression of selected angiogenic genes by semiquantitative RT-PCR analysis. Dose-response effect (a) and time-course effect (b) of triptolide on selected genes expression by semiquantitative RT-PCR analysis. The expression of each target gene was normalized to GAPDH, and then expressed as percentage of control group.

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We then studied the temporal expression of these genes in HUVECs treated with 100 nM triptolide (Fig. 4b). Triptolide at this dose, respectively, decreased by 30, 50 and 40% the mRNA expression of Ang2, VEGF-A and VEGFR-2 at as early as 4-h posttreatment. The peak suppression of Ang2, Tie2, VEGFR-2 and VEGF-A (≥50%) was observed at 18–24-h posttreatment. Among the 4 genes, VEGFR-2 expression was completely suppressed at 18-h posttreatment. By contrast, the expression of VEGFR-1 and Tie1 was relatively resistant to triptolide. Because both VEGFR-2 and Tie2 are 2 key receptors involved in angiogenic mediation, time- and dose-dependent downregulation of both receptors, VEGFR-2 and Tie2, by triptolide was further confirmed by western blot analysis (Figs. 5a and 5b). Consistent with the observation of mRNA expression, VEGFR-2 expression appeared more sensitive to triptolide than that of Tie2. Tie2 downregulation was a later event than VEGFR-2. Tie2 expression at 48-h incubation decreased to the same level, one-fifth of control, as VEGFR-2 at 6-h incubation. These data indicate that triptolide inhibited angiogenesis via differential downregulation of both VEGF-A and Ang2-mediated signaling.

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Figure 5. Inhibitory effect of triptolide on VEGF-A and Tie2 expression by western blotting analysis. Dose-response effect (a) and time-course effect (b) of triptolide on VEGF-A and Tie2 expression. Data were normalized to β-Actin and then expressed as percentages of control group.

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Tie2 overexpression enhanced HUVECs resistance to antiproliferative and antinetwork forming effect of triptolide

Although triptolide suppressed the expression of both VEGFR-2 and Tie2, the effect of natural products on Tie2 has been rarely studied.29 We thus investigate the effect of Tie2 overexpression on antiproliferative and antinetwork formative effect of triptolide. Infection of HUVECs with Tie2-bearing adenovirus at MOI of 5–40 dose-dependently increased Tie2 expression (Fig. 6a). MOI at 5 was used for subsequent cell proliferation assay. As shown in Figure 6b, increased Tie2 expression significantly enhanced the resistance of HUVEC proliferation to triptolide inhibition even at 2,000 nM. The IC50 of triptolide on wild-type HUVEC proliferation was 45 nM, whereas the IC50 for Tie2-overexpressing HUVECs was elevated to be higher than 2,000 nM (>60-fold increase). Antinetwork formative effect of triptolide at 1,000 nM was also dose-dependently attenuated by Tie2 overexpression. At 9 h, Tie2 overexpression at MOI 5–20 gradually rescued triptolide treatment. HUVECs with Tie2 overexpression at MOI 40 were almost completely resistant to high concentration of triptolide at 1,000 nM (Fig. 6c). Together, Tie2 overexpression alone dramatically attenuated the antiangiogenic effect of triptolide, indicating a critical role of Tie2 in the antiangiogenic effect of triptolide.

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Figure 6. Inhibitory effect of triptolide on the proliferation and network formation of Tie2-over-expressing HUVECs. (a) Western blotting analysis of Tie2-over-expression in HUVECs by different adenovirus MOI. (b) Following adenoviral infection at MOI = 5 (AdNull verus Ad-Tie2), triptolide-treated HUVEC proliferation was expressed as percentage of control cells. Each point represents mean + SEM (n = 4) from 3 independent experiments. *p < 0.05; **p < 0.01; ***p < 0.001. The data were analyzed by one-way ANOVA followed by the Dunnett's test for comparisons of all treated groups with control group. (c) Images showing network formation of Ad-Tie2 infected HUVECs at MOI of 5–40. The network length (indicated by arrow) in each treatment condition was quantified and shown in lower panel. Each point represents mean + SEM (n = 4) from a representative experiment. *p < 0.05; **p < 0.01. The data were analyzed by Student's t test for comparisons of all treated groups with respective control group.

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Tie2 expression was critical for HUVECs to form networks on Matrigel

Tie2 plays an important role in modulating network formation, maturation and stability.30 We knocked down the expression of Tie2 to examine the knockdown effect on wild-type HUVEC network formation assay. First, 2 different Tie2-targeting shRNA clones (412 and 414) were, respectively, used to generate partial and complete Tie2 knockdown (Fig. 7a). ShRNA to Luc served as a negative control. Because 9–12 h incubation was the peak time for network formation, we recorded peak network formation at 12 h in this assay. There was no difference in the network-forming abilities between Luc and parental HUVECs (Fig. 7b). Both cells formed networks at 12 h, and the networks remained stable for at least 24 h on Matrigel (data not shown). By contrast, only a few networks were observed at 12-h postseeding in the partial knockdown cells. Only residual, fragmented networks remained at 12 h in the complete knockdown cells (Fig. 7b). Together, HUVECs with reduced Tie2 expression had less potential to form and maintain network structures than wild-type HUVEC. Moreover, the network forming ability of Tie2-knockdown HUVECs resembled the inhibitory effect of triptolide on wild-type HUVECs, suggesting a role of Tie2 in triptolide-mediated inhibition of network formation.

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Figure 7. Inhibitory effect of triptolide on the network formation of Tie2-knockdown HUVECs. (a) Western blotting analysis of Tie2 knockdown in HUVEC by lentivirus. (b) Representative images depicting formation of capillary like structures by different types of HUVECs following 12-h treatment. The network length in each treatment condition was quantified and shown in the lower panel. Each point represents mean + SEM (n = 4) from a representative experiment. a: p < 0.5 compared with control; b: p < 0.05 compared with Luc; c: p < 0.01 compared with control; d: p < 0.01 compared with Luc and e: p < 0.01 compared with 412 clone. The data were analyzed by one-way ANOVA followed by the Newman–Keuls test for comparisons of all pairs of columns. [Color figure can be viewed in the online issue, which is available at]

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Antiangiogenic effect of triptolide on In vivo Matrigel plug model

Triptolide was a potent angiogenesis inhibitor in vitro. We further assessed the in vivo antiangiogenic effect of triptolide using Matrigel plug assay. We tested 2 doses of triptolide, 100 and 500 nM, together with vehicle control. BFGF (100 ng/ml) together with heparin (64 U/ml) induces new blood vessel formations in plug from nearby tissues at 10-day postinjection (positive control). DMSO as a vehicle control had no inhibition on the vessel formation in the plug (Fig. 8a and Supporting Fig. 1). Triptolide at 100 nM potently inhibited angiogenesis in the plug induced by bFGF and heparin. Triptolide at 500 nM almost completely blocked angiogenesis and the Matrigel plugs looked transparent (Fig. 8a). The vessel formation in plugs was further confirmed by immunohistochemical staining of CD31, an endothelial maker. The CD31-positive vessel density decreased with the increase of triptolide concentration (Fig. 8b). Together, triptolide potently blocks angiogenesis in vivo.

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Figure 8. Inhibitory effects of triptolide in in vivo mouse Matrigel Plug assay. Animals were injected with: Matrigel plus 64 U/ml heparin and 100 ng/ml bFGF (positive control); Matrigel plus 64 U/ml heparin, 100 ng/ml bFGF and 0.1% DMSO (vehicle control); Matrigel plus 64 U/ml heparin, 100 ng/ml bFGF and 100 nM triptolide (100 nM) and Matrigel plus 64 U/ml heparin, 100 ng/ml bFGF and 500 nM triptolide (500 nM). (a) Images of Matrigel plug removed from respective animals. (b) CD31 staining of the blood vessels in the Matrigel from respectively treated animals. Scale bars: 1 cm (a), 40 μm (b). Each point represents mean + SEM (n = 4 or 5) from a representative experiment. *p < 0.05; **p < 0.01. The data were analyzed by one-way ANOVA followed by the Dunnett's test for comparisons of all treated groups with control group. [Color figure can be viewed in the online issue, which is available at]

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Antitumor and antiangiogenic effects of triptolide on in vivo murine tumorigenesis model

Based on our present data, triptolide is a potent angiogenesis inhibitor. To minimize the direct inhibitory activity of triptolide on tumor cell, B16 melanoma cells were selected and injected into syngenic CL57/B6 mice for murine tumorigenesis model. Angiogenesis occurs when tumor reaches 2–3 mm in diameter and goes on to supply nutrients for further tumor progression.31, 32 To test if triptolide could be used as an antitumor agent through blocking tumor angiogenesis, we administered triptolide to the mice right after tumor cell implantation in this tumor model. A group of researchers reported that a water-soluble form of triptolide ranged from 0.25 to 0.75 mg/kg/day potently inhibited tumor progression without any observable side effect.18 The same concentration range of triptolide was used in this study. Triptolide potently inhibited tumor progression in a dose-dependent manner (Figs. 9a and 9c). At 0.75 mg/kg/day, tumor tissues shrank to one-fifth of control tumors by weight (Fig. 9c). Immunohistological analysis of the tumor vessels (indicated by arrow) showed that the vessel density in tumor tissue decreased with the increase of triptolide dosage (Figs. 9b and 9d). The blood vessel formation in the tumor tissue was significantly blocked by triptolide at 0.75 mg/kg/day treatment. These data further underscore the potency of triptolide to suppress tumor growth in vivo and correlate its effect on the inhibition of angiogenesis in vitro.

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Figure 9. In vivo murine tumorigenesis assay. Animals were treated with control, 0.25, 0.50 and 0.75 mg/kg/day. (a) Tumor tissues removed from respective animals. (b) Immunohistochemical analysis of blood vessels (arrow) in tumor tissue from respectively treated animals. (c) Dose-response effect of triptolide on tumor weight. (d) Dose-response effect of triptolide on tumor vascular density. Each point represents mean + SEM (n = 6) from a representative experiment. *p < 0.05; **p < 0.01; ***p < 0.001. Scales bars: 1 cm (a), 100 μm (b). [Color figure can be viewed in the online issue, which is available at]

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  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

In this study, we were able to demonstrate that triptolide potently inhibited angiogenesis in vitro and in vivo, manifested by endothelial cell proliferation, migration, network formation and tumor angiogenesis. The inhibition was in part through potent inhibition of Tie2 and VEGFR-2 production at both transcriptional and posttranscriptional levels, especially the production of VEGFR-2. This is a very exciting result since both VEGFR-2 and Tie2 are primarily expressed in endothelial cells.33 Moreover, a multitargeted antiangiogenic druglike Sutent, a multityrosine kinase inhibitor, has been recently approved by FDA to treat advanced cancer.34 Triptolide may thus be used as a lead compound for the development of a potential multitarget drug for cancer treatment.

Although triptolide inhibited both the production of VEGFR-2 and Tie2, triptolide blocked VEGFR-2 expression and production at earlier time points than that of Tie2 (Figs. 4b and 5b). This phenomenon agrees very well with the notion that 2 ligand-receptor pairs, VEGF-A-VEGFR-2 and Angs-Tie2, play complementary and coordinated roles during vascular development.35, 36 In early stages of angiogenesis, VEGFR-2 and VEGF-A play a crucial role in vessel spouting and new vessel initiation through induction of proliferation, migration and survival of endothelial cells.37 By contrast, Tie2 and Angs play a critical role in stabilizing the immature endothelial cell network, attracting pericytes and maintaining vessel integrity, which are thought to be implemented in the later stage of blood vessel formation.38–40 These 2 pathways are considered as 2 independent mediators essential for the process of in vivo angiogenesis,41 because the inhibition of VEGFR-2 pathway cannot be compensated by the Tie2 pathway, nor vice versa.41 Thus, a dual inhibitor of VEGFR-2 and Tie2 would be expected to demonstrate synergistic effects through inhibition of both critical stages of blood vessel formation and offers a potential for the development of new antiangiogenic agents.42 Triptolide targeted both at VEGFR-2 and Tie2 in the cascades of angiogenesis, revealing triptolide as a promising angiogenesis inhibitor.

In spite of various approaches used to reduce tumor growth and tumor vascularization via blockage of VEGF-A/VEGFR-2 signaling, there were still tumors remaining unaffected.43 Moreover, definitive preclinical studies have shown that blockage of VEGF-A signaling led to the development of alternative VEGF-A-independent proangiogenic signaling pathways in the tumor cells resistant to anti-VEGF-A treatment.44 These findings have drawn researchers' attention to alternative signaling pathways involved in tumor angiogenesis.41 We, therefore, focused on the role of Tie2 in triptolide-mediated antiangiogenesis. Both proliferation and network formation assays using Tie2-genetically manipulated HUVECs and migration assay using Ang1 as a chemoattractant revealed that Tie2 plays an important role in angiogenesis. Tie2 overexpression alone was sufficient to attenuate triptolide-mediated inhibition of HUVEC proliferation and network formation. These observations were in agreement with 1 earlier study using xenografts derived from tumor cells expressing soluble Tie2 (sTie2). Tumor cells expressing sTie2, behaving as a sink for Ang1, reached only 4–8% of the size of the control tumor, and the vessel density was statistically significant reduced by a factor of 4–5 in these tumors.41 Taken together, the inhibition of Tie2 signaling pathway plays a crucial role in triptolide-mediated angiogenesis inhibition.

Although the antitumor activity of triptolide has been previously investigated in cancer cells,17 we are the first to demonstrate in vitro and in vivo antitumor action of triptolide partly via inhibition of tumor angiogenesis. Moreover, the potent antiangiogenic action exerted by triptolide was through inhibiting the propagation of 2 key angiogenic pathways mediated by VEGFR-2 and Tie2. The IC50 of triptolide in our in vitro HUVEC assays was at 45 nM, which was much lower than the plasma concentrations of triptolide detected in the rat or human when extracts of T. wilfordii was administered as antisystemic lupus and antiinflammatory agents.42, 45, 46 Triptolide also more potently inhibited the proliferation of HUVECs than other normal cells including skin keratinocytes HaCaT cells in our study and liver cells L-02.47 These indicate that triptolide should have less side effects when used as antiangiogenic agent at lower doses although triptolide was reported to have male reproductive toxicity48 and hepatic injury.47

Because angiogenesis is a rate limiting step for tumor progression and the prevention of angiogenic switch in the early phase of tumor progression can effectively delay tumor progression,32, 49 we used a prevention protocol to evaluate the antiangiogenic potential of triptolide in murine tumorigenesis. A significant delay of tumor progression was observed in the tumor-bearing mice treated daily with triptolide at 0.25–0.75 mg/kg partly via antiangiogenesis. Although our study was not able to completely address clinical utility of using this compound in vivo, a long-term use of triptolide at low dose is anticipated to be well tolerated. The reasons are several folds. First, the parental herb, T. wilfordii, has been traditionally used as 1 medicinal herb. Second, a 71% response rate was detected in leukemia patients treated bi-daily with a dose as low as 30 μg/kg. Assuming a water to weight conversion, these patients were treated with a equivalent dose of 10–40 ng/ml (28–112 nM) without suffering severe side effects.50 Third, triptolide shows great promise in the phase II clinical trial for treating arthritis in the United States.50 Fourth, triptolide was reported to be effective for treating psoriasis vulgaris with daily oral intake of 30–60 mg triptolide tablet (equivalent to 12–24 μg triptolide) for 2–6 months.10 More evidence is, however, needed to support whether triptolide can be used as a chemopreventive agent. Taking into the consideration of the immunosuppressive, hepatoxic and even antimale reproductive effects of triptolide in clinical and preclinical settings, treating neoplastic diseases with a long-term, low dose of triptolide to normalize blood vessels (metronomic therapy49) combined with other chemotherapeutic regimens should be more feasible in the clinical setting for this drug. Finally, the use of triptolide as a lead compound for the development of antiangiogenic drug should hold great potential of developing novel anticancer drug for cancer treatment.


  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

We thank Miss MZ Chang for her help in immunohistochemical staining of Matrigel plugs and tumor sections. ShRNA reagents were obtained from the National RNAi Core Facility at the Institute of Molecular Biology/Genomic Research Center, Academia Sinica.


  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Additional supporting information may be found in the online version of this article.

IJC_24694_sm_suppinfofig1.tif4842KSupplementary Figure 1.

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