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Induction of apoptosis in human leukemia cells through an intrinsic pathway by cathachunine, a unique alkaloid isolated from Catharanthus roseus.


Background: Catharanthus roseus (L.) G. Don consists of a range of dimeric indole alkaloids with significant antitumor activities. These alkaloids have been found to possess apoptosis-inducing activity against tumor cells in vitro and in vivo mediated by nuclear factor kappa-light-chain-enhancer of activated B cells (NF-[kappa]B) and c-Jun N-terminal kinase (JNK) pathways, in which DNA damage and mitochondrial dysfunction play important roles. In this study, a unique bisindole alkaloid named cathachunine, along with five known dimeric indole alkaloids, was obtained from C. roseus and investigated in vitro.

Purpose: The aim of this study was to investigate the antitumor activity of isolated alkaloids and the mechanism through which cathachunine exerts its antitumor effect.

Study design and methods: Cell growth inhibition was assessed by WST-1 and lactate dehydrogenase (LDH) assays in HL60, K562 leukemia cells and EA.hy926 umbilical vein cells. Induction of apoptosis in HL60 cells was confirmed by observation of nuclear morphology, a caspase-3 activity assay and annexin V-fluorescein isothiocyanate/propidium iodide (FITC/PI) double staining. The intrinsic apoptotic pathway induced by cathachunine was evidenced by B-cell lymphoma 2/Bd-2-associated X protein (Bcl-2/Bax) dysregulation, loss of mitochondrial membrane potential, translocation of cytochrome c, and cleavage of caspase-3 and poly-ADP ribose polymerase (PARP). Reactive oxygen species (ROS) production after cathachunine treatment was determined by 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA) staining. Cell cycle arrest of the S phase was also observed in HL60 cells after cathachunine treatment.

Results: The WST-1 and LDH assays showed that Catharanthus alkaloids were cytotoxic toward human leukemia cells to a greater extent than toward normal human endothelial cells, and the anti-proliferation and pro-apoptosis abilities of cathachunine were much more potent than other previously reported alkaloids. The induction of apoptosis by cathachunine occurred through an ROS-dependent mitochondria-mediated intrinsic pathway rather than an extrinsic pathway, and was regulated by the Bcl-2 protein family.

Conclusion: An unprecedented bisindole alkaloid cathachunine which lost C-18' and C-19' was isolated from C. roseus. It exerted a potent antitumor effect toward human leukemia cells through the induction of apoptosis via an intrinsic pathway. Thus, this study provides evidence for a new lead compound from a natural source for anti-cancer investigations.




Intrinsic pathway



Catharanthus roseus (L.) G. Don is a pantropical plant widely studied by pharmacognosists that contains about 130 terpenoid indole alkaloids (van Der Heijden et al. 2004). Extracts from this plant have been used to treat numerous diseases, including diabetes, malaria and cancer. The alkaloids have been noted to exert an apoptosis-inducing ability towards tumor cells in vitro and in vivo mediated by the nuclear factor kappa-light-chain-enhancer of activated B cells (NF-[kappa]B) and c-Jun N-terminal kinase (JNK) pathways, in which DNA damage and mitochondrial dysfunction play important roles (Chiu et al. 2012; Huang et al. 2004, 2012). Among the dimeric indole alkaloids with significant antitumor activity, vinblastine and vincristine are typical compounds that have been used extensively for the clinical treatment of human cancers, such as testicular carcinoma, acute leukemia, rhabdomyosarcoma and breast cancer. Vinblastine induces apoptosis in melanoma cells through mitochondrial and non-mitochondrial pathways mediated by the Rho A protein (Selimovic et al. 2013). Meanwhile, vincristine induces neuroblastoma cell death through mitotic arrest and mitochondria-dependent apoptosis (Tu et al. 2013). Previous investigations, including our own work, have attempted to identify diversiform dimeric indole alkaloids. These efforts have led to the identification of new alkaloids with significant cytotoxicity to human hepatocellular carcinoma (HepG2), human colorectal carcinoma (Lovo) and human breast carcinoma (MCF-7) cell lines (Wang et al. 2012, 2014; Zhang et al. 2013a, 2013b). However, further investigation is necessary for the chemical isolation of such alkaloids, as well as biological studies in order to understand more about their antitumor mechanisms.

Apoptosis is a highly regulated process of programmed cell death that can be triggered through either extrinsic or intrinsic pathways (Bai and Wang 2014). The extrinsic (death receptor) pathway begins with stimulation of death receptors on the cell membrane (Wallach et al. 2008). The intrinsic (mitochondrial) pathway is initiated by dysfunction of the mitochondria, which leads to the release of signaling factors, such as cytochrome c (cyto c), to the cytosol. Permeabilization of the mitochondrial membrane relies on the B-cell lymphoma 2 (Bcl-2) protein family, where pro-apoptotic Bcl-2-associated X (Bax) or Bcl-2 homologous killer (Bak) proteins oligomerize to form pores on the outer mitochondrial membrane (OMM; Tait and Green 2010). These two pathways are executed mainly by caspases (a family of cysteine proteases), with caspases-8 and -9 engaging in the extrinsic and intrinsic pathways, respectively (Ola et al. 2011). In addition, reactive oxygen species (ROS), a series of oxygen metabolism byproducts, have been closely related to cancer cell apoptosis induced by natural alkaloids (Kardeh et al. 2014).

In this study, a unique bisindole alkaloid named cathachunine (Fig. 1, F) was obtained from C. roseus and its chemical structure was elucidated by high resolution electrospray ionization mass spectrometry (HR-ES1-MS), ID and 2D nuclear magnetic resonance (NMR), circular dichroism (CD) spectrophotometry, and infrared (IR) and ultraviolet (UV) spectroscopy. Subsequently, the cytotoxic mechanisms of cathachunine towards human leukemia cells were investigated, along with five other previously isolated compounds (leurosine, A; Catharine, B; cycloleurosine, C; 15'-R-hydroxyvinamidine, D; and 17-deacetoxyvinamidine, E; Fig. 1). Growth inhibition in HL60, K562 leukemia and EA.hy926 umbilical vein cells by Catharanthus alkaloids was detected by WST-1 and lactate dehydrogenase (LDH) assays. The influence of the alkaloids on cell apoptosis was demonstrated through nuclear morphology observation, annexin V-fluorescein isothiocyanate/propidium iodide (FITC/PI) double staining and caspase-3 activity. Additional experiments were performed to investigate the apoptosis mechanism, which was found to be an ROS-dependent mitochondria-mediated intrinsic pathway. Cell cycle arrest of the S phase was also found in HL60 cells after treatment with F.

Materials and methods


IR and UV spectra were obtained on JASCO FT/IR-480 Plus and JASCO V-550 UV/VIS spectrophotometers, respectively. Melting points (mp) were recorded on an X-5 micro mp apparatus (uncorrected). NMR spectra were run on a Bruker AV-400 spectrometer. HR-ESI-MS data were detected on an Agilent 6210 ESI/TOF mass spectrometer. CD spectra were obtained on a JASCO J-810 spectropolarimeter at room temperature. High performance liquid chromatography (HPLC) separations were performed on a COSMOSIL [C.sub.18] preparative column (5 [micro]m, 20 mm x 250 mm; Nacalai Tesque, Kyoto, Japan).

Plant material

The whole plants of C. roseus were collected in Weifang, Shandong Province, PR China, in October 2012 and authenticated by Dr. Chunhua Wang. A voucher specimen (no. 20121004CH) was deposited in the Tianjin International Joint Academy of Biotechnology & Medicine, Tianjin, PR China.

Extraction and isolation

The dried whole plants (3.0 kg) of C. roseus were extracted with 95% ethanol and the resulting solution was evaporated in vacuo to a residue (180 g) as a part of our previous work (Wang et al. 2012, 2014). The residue was dissolved in water to form a suspension and extracted with dichloromethane (DCM) to obtain an extract (112 g), which was subjected to silica gel column chromatography (CC; DCM : methanol, 100 : 0 [right arrow] 0 :100) to give nine fractions. In previous work, compounds A-E (Fig. 1) were isolated from fractions 6 and 7, and further investigation of fraction 8 led to the isolation of compound F (25.0 mg) along with compound E (22 mg), which were purified by reverse phase HPLC (RP-HPLC) using a mobile phase of acetonitrile (0.1% diethylamine) and water.

Cell culture and drug treatment

HL60 human acute promyelocytic leukemia cells, K562 human chronic myelogenous leukemia cells and EA.hy926 human umbilical vein cells [American type culture collection (ATCC), USA] were cultured in Roswell Park Memorial Institute medium (RPMI 1640) or Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum, 100U/ml penicillin and 100 [micro]g/ml streptomycin in 25 [cm.sup.2] culture flasks at 37[degrees]C in a humidified atmosphere with 5% C[O.sub.2]. Cells used in the assays were from passage numbers 3-6. For the drug treatment experiments, cells were harvested from the culture during the exponential growth phase and then seeded into multi-well culture plates at 5 x [10.sup.4] to 1 x [10.sup.5] cells/ml in fresh medium. After overnight growth, the cells were treated with the compounds at selected concentrations for a period of 1-3 days.

WST-1 assay for cell viability

Cell viability was determined using a WST-1 cell proliferation and cytotoxicity assay kit (Beyotime Institute of Biotechnology, China), according to the supplier's manual. Briefly, at the end of the drug treatment period, 10 [micro]l WST-1 solution was added to each well of the culture plate (containing 100 [micro]l medium). After 2 h incubation at 37[degrees]C, the optical density (OD) of the assay solution was measured at 450 nm with a spectrophotometer (BioTek, USA). Cell viability was calculated as a percentage with the following equation: % cell viability = ([OD.sub.sample]/[OD.sub.control]) x 100. The 50% inhibition concentration ([IC.sub.50]) values were determined by plotting cell viability versus drug dose.

Assessment of cell number

At the end of the drug treatment period, cells were collected from the 6-well culture plates and the cell number (N) was counted using trypan blue dye exclusion of the dead cells. Cell viability was calculated as a percentage with the following equation; % cell number--([N.sub.sample]/[N.sub.control]) x 100.

LDH assay for plasma membrane damage

LDH leakage from cells into the culture medium was assessed using an LDH cytotoxicity assay kit (Beyotime Institute of Biotechnology, China), according to the supplier's manual. Briefly, the culture medium was aspirated and the supernatant was obtained by centrifugation. Then, the assay buffer was prepared and added to the sample, and the OD of the assay solution was measured at 490 nm with a spectrophotometer (BioTek, USA). The percentage of LDH release was calculated by following the instructions of the manufacturer.

Detection of nuclear morphology

At the end of the drug treatment period, the cells in each well were washed once with phosphate-buffered saline (PBS) and fixed with 4% formaldehyde in PBS at 4[degrees]C for 30 min. The cells were then washed with PBS and stained with 1 [micro]g/ml Hoechst 33258 in PBS at 37[degrees]C for 15 min, and then viewed under a fluorescence microscope (Leica, Germany) for any change in nuclear morphology.

Annexin V-FITC/PI double staining

Annexin V-FITC/PI apoptosis analysis was undertaken using an annexin V-FITC apoptosis detection kit (Beyotime Institute of Biotechnology, China), according to the supplier's manual. Briefly, cells were collected, washed twice with PBS and rewashed with binding buffer. Then, cells were stained with annexin V-FITC/PI at room temperature in the dark for 15 min and the fluorescence was subsequently quantified by flow cytometry (BD Biosciences, USA). Early and late apoptotic events were identified by annexin V+/ PI- and annexin V+/ PI+ staining.

Caspase activity assay

Caspases-3, -8 and -9 activities were determined using caspases-3, -8 and -9 colorimetric assay kits (Biovision, USA), according to the supplier's manual. Briefly, cells were resuspended in cell lysis buffer and centrifuged to obtain the supernatant, which was assessed for the protein concentration. Then, the samples were incubated at 37[degrees]C with reaction buffer and substrate for 1-2h. Finally, the OD of the assay solution was measured at 405 nm with a spectrophotometer (BioTek, USA).

Cell cycle analysis

Cell synchronization was performed using a double thymidine block method (Simoes et al. 2015). Briefly, cells were cultured with 2 mM thymidine for 14 h (first block) and then released from the block for 10 h, followed by an additional 14 h block with thymidine (second block). Cells were treated with the alkaloid after synchronization, and measurement of the cellular DNA content for cell cycle analysis was carried out by PI staining. Cells were washed with ice-cold PBS and fixed in 75% ethanol at -20[degrees]C for at least 1 h. After that, cells were washed twice, incubated with 0.5 mg/ml R-Nase A and stained with 10|ig/ml PL Fluorescence emitted from the PIDNA complex was quantitated by flow cytometry (BD Biosciences, USA).

Mitochondrial membrane potential (MMP) measurement

The MMP was determined using a JC-10 assay kit for microplate readers (Sigma-Aldrich, USA), according to the supplier's manual. Briefly, after apoptosis induction, cells in 96-well plates were treated with 50 [micro]l/well JC-10 dye loading solution and incubated at 37[degrees]C, 5% C[O.sub.2] for 1 h in the dark. Then, 50 [micro]l /well assay buffer was added and the fluorescence intensities were monitored with a fluorescence microplate reader (Berthold, Germany) at excitation and emission wavelengths of 490 and 525 nm (green) or 540 and 590 nm (red), respectively. The ratio of red/green fluorescence represented the MMP of the cells.

Measurement of ROS generation

The ROS level of the cells was examined using 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA; Invitrogen, USA). Cells were washed once with PBS and incubated with DCFH-DA (10 [micro]M final concentration) at 37[degrees]C for 30 min in the dark. Then, the cells were washed twice and maintained in 1 ml culture medium. ROS generation was assessed using a fluorescence microscope (Leica, Germany) at excitation and emission wavelengths of 488 and 530 nm, respectively. The fluorescence intensity was further analyzed by a fluorescence microplate reader (Berthold, Germany) at excitation and emission wavelengths of 488 and 525 nm, respectively.

Quantitative real-time polymerase chain reaction (qRT-PCR)

At the end of the drug treatment period, the cells in each well were lysed in TRIzol solution. RNA was extracted with an RNAiso Plus kit (Takara, Japan) according to the manufacturer's protocol and quantitated spectrophotometrically. Total RNA was used as a template for reverse transcription using the following protocol: each 20 [micro]l reaction contained 1 x M-MLV buffer, 125 [micro]M deoxynucleotide (dNTP), 100pmol oligo [dT.sub.18] primer, 100 units of M-MLV reverse transcriptase, diethylpyrocarbonate- (DEPC-) treated water and 2 [micro]g total RNA. Briefly, RNA and oligo [dT.sub.18] primer were incubated at 70[degrees]C for 10 min and then immediately placed on ice, after which the other components were added and incubated at 42[degrees]C for 1 h and then at 70[degrees]C for 15 min. RT-PCR was performed on a CFX96 RT-PCR Detection System (Bio-Rad, USA) using SYBR Premix Ex Taq (Takara, Japan), according to the manufacturer's protocol. Primer sequences are shown in Table 1 (Bcl-F and Bd-R for the Bcl-2 gene; Bax-F and Bax-R for the Bax gene; GAPDH-F and GAPDH-R for the glyceraldehyde-3-phosphate dehydrogenase gene). The RT-PCR reaction mixture contained 10 [micro]l SYBR Premix Ex Taq, 0.2 [micro]M each of the forward and reverse primers, 2 [micro]l cDNA and nuclease-free water to a total volume of 20 [micro]l. The program used for all genes consisted of a denaturing cycle of 30 s at 95[degrees]C, 40 cycles of PCR (95[degrees]C for 5 s and 60[degrees]C for 30 s) and a settled melting cycle in order to confirm the specificity of the RT-PCR products. The product sizes were confirmed by agarose gel electrophoresis and ethidium bromide staining. Results were analyzed using the [2.sup.-[DELTA][DELTA]CT] method to compare the transcriptional levels of target genes normalized to the GAPDH gene in each sample relative to the untreated control.

Extraction of mitochondrial proteins

Mitochondrial proteins were extracted using a cell mitochondria isolation kit (Beyotime Institute of Biotechnology, China), according to the supplier's manual. Briefly, cells (2 x [10.sup.7]) were collected by centrifugation, washed with ice-cold PBS and resuspended with mitochondria extraction buffer on ice for 10 min. Then, cells were homogenized in an ice-cold dounce tissue grinder and the homogenate was centrifuged at 700 x g for 10 min to remove the pellet. After further centrifugation at 10,000 x g for 30 min at 4[degrees]C, the supernatant was collected as the cytosolic fraction, with the pellet being the intact mitochondria. Following resuspension with mitochondria lysis buffer, the mitochondrial fraction of the cell protein lysate was obtained.

Western blotting

After drug treatment, the cells in each well were disrupted with cell lysis buffer (Beyotime Institute of Biotechnology, China). The resulting suspension was centrifuged at 12,000 x g, 4[degrees]C for 5 min and the protein content of the supernatant was measured with a bicinchoninic acid (BCA) assay. Equal amounts of protein sample were loaded onto a 10% sodium dodecyl sulfate-polyacrylamide gel (SDS-PAG) and then transferred to a microporous polyvinylidene difluoride (PVDF) membrane. Western blotting was performed using rabbit anti-human Bcl-2 monoclonal antibody, Bax monoclonal antibody, cyto c monoclonal antibody, caspase-3 polyclonal antibody, poly-ADP ribose polymerase (PARP) polyclonal antibody, voltage-dependent anion channel (VDAC) monoclonal antibody or [beta]-actin monoclonal antibody (1 : 1000 dilution; Cell Signaling Technology, USA) and horseradish peroxidase-conjugated anti-rabbit secondary antibody (1 : 1500 dilution; Cell Signaling Technology, USA). Protein bands were visualized using an enhanced chemiluminescence (ECL) substrate (Pierce, USA).

Statistical analysis

Data were expressed as the means [+ or -] standard deviation (SD) for the indicated number of independently performed experiments. Student's t test was used for the determination of statistical significance. The difference was considered to be statistically significant when P < 0.05.


Compounds isolated from C. roseus

Six dimeric indole alkaloids were isolated from C. roseus as shown in Fig. 1 (compounds A-F). Compound F was obtained as a white amorphous powder and its molecular formula was determined to be [C.sub.41] [H.sub.48] [N.sub.4] [O.sub.7] via HR-ES1-MS (m/z 709.3605, [[M+H].sup.+]). The IR spectrum of F showed absorptions due to OH (3452 [cm.sup.-1]), C = O (1737 [cm.sup.-1]), O = C-N< (1688 [cm.sup.-1]) and an aromatic ring (1616, 1456, 1244 [cm.sup.-1]). The [sup.1]H NMR spectrum showed four aromatic protons of the indole moiety [[delta] 7.53 (1H, d, J = 7.6 Hz), 7.16 (1H, m), 7.13 (1H, m) and 7.11 (1H, d, J = 7.6 Hz)], four protons typical of a dihydroindole moiety [[delta] 6.52 (1H, s), 6.08 (1H, s), 5.67 (1H, dd, J = 9.6, 4.0 Hz) and 5.46 (1H, d, J = 9.6 Hz)] and five methyl groups [[delta] 3.78 (6H, s, 2 x OC[H.sub.3]), 3.58 (3H, s), 2.48 (3H, s) and 0.89 (3H, t, J = 7.6Hz)]. Accordingly, the [sup.13]C NMR and distortionless enhancement by polarization transfer (DEPT-135) spectra of F showed the characteristic signals ascribed to a dihydroindole unit [[delta] 158.1 (C), 153.2 (C), 123.9 (C), 122.7 (CH), 119.7 (C) and 93.6 (CH)], typical signals due to the indole moiety [[delta] 134.8 (C), 132.6 (C), 129.1 (C), 123.0 (CH), 119.4 (CH), 118.3 (CH), 114.9 (C) and 110.6 (CH)], two olefinic carbons at [delta] 136.0 and 121.9, and three carbonyls at [delta] 179.0, 174.3 and 173.6. This data indicated that compound F was a Catharanthus alkaloid and an analogue of 17-deacetoxyvinamidine (E), which was also isolated in this study.

Comparison of the NMR data of compound F with that of 17-deacetoxyvinamidine revealed that the NMR data of the dihydroindole moiety was almost the same as 17deacetoxyvinamidine, with the aldehyde group of N-CHO and an ethyl at the C-18' and 19' positions of 17-deacetoxyvinamidine absent in the indole group of compound F. Moreover, an extra amide bond ([delta] 179.0) in the ring was observed. This data indicated that the ethyl at C-18' and 19' was lost, and that C-20 and N-4 were linked via an amide bond. The key heteronudear multiple bond correlations (HMBCs) between H-3'a ([delta] 3.99)/H-5'b ([delta] 2.99)/H-14' ([delta] 1.54) and C-18 ([delta] 179.0) confirmed the above deduction (Fig. 2). The relative configuration was established by the rotating frame overhauser effect spectroscopy (ROESY) correlations (Fig. 2), which were the same as the relative configuration of 17-deacetoxyvinamidine. Furthermore, the absolute configuration was established by CD analysis (Fig. 3), which showed peaks ([[lambda].sub.max]) at 228, 261.5 and 315 nm (positive Cotton effect) and troughs at 195, 201 and 215.5 nm (negative Cotton effect). The data was similar to that of 17-deacetoxyvinamidine, as reported in the literature (Wang et al., 2012), which indicated that compound F had the same absolute configuration as 17-deacetoxyvinamidine. Thus, compound F was identified and named cathachunine.

Cathachunine (F): white amorphous powder, mp 208-210[degrees]C, [[alpha]]20D +22.5 (c 0.2, CH[C.sub.l3]). UV (CHCN) [[lambda].sub.max] in nm (log [epsilon]): 220 (0.68), 260 (0.19). IR (KBr) in [cm.sup.-1]: 3452, 1688, 1616, 1456, 1244, 1144, 741. [sup.1]H NMR and [sup.13]C NMR data, see Table 2. HR-ESI-MS m/z: 709.3605 [[M+H].sup.+] (calcd for [C.sub.41] [H.sub.49] [N.sub.4] [O.sub.7], 709.3601).

Alkaloids reduced leukemia cell viability and caused membrane damage

Cell growth inhibition after treatment with the alkaloids for 48 h was determined by a WST-1 assay, as shown in Fig. 4A-C. All of the compounds exhibited a dose-dependent increase of growth inhibition in the range of 0.1-100 [micro]M. Much lower inhibitory effects were demonstrated in EA.hy926 human umbilical vein cells (a normal human endothelial cell line) than in HL60 cells and K562 human leukemia cells. The relative cytotoxicity potency of compounds A-F in HL60 and K562 cells was determined and it was found that compound F was the most potent, while D was the least potent. This finding was consistent with the IC5o values of these compounds, as shown in Table 3. Cell numbers were also counted in HL60, K562 and EA.hy926 cells after treatment with compound F for 48 h (Fig. 4D). The results confirmed the findings of the WST-1 assay.

Thus, compound F was chosen for further investigation. It exhibited a time- (24, 48 and 72 h; Fig. 4E) and dose-dependent (0.3, 1, 3, 10, 30, 100 [micro]M; data not shown) inhibitory effect on HL60 cells by WST-1 assay, and an LDH assay showed damage to the plasma membrane after treatment in a similar time- and dose-dependent manner (Fig. 4F). These results suggested that increased concentrations and treatment times led to greater growth inhibition and membrane damage in HL60 cells.

Alkaloids induced apoptosis in HL60 leukemia cells

Caspase-3 is a well-defined execution enzyme of the caspase family that is involved in the process of apoptosis. Accordingly, a caspase-3 activity assay was performed to obtain a quantitative comparison of the compounds with regards to their apoptosis-inducing activities in HL60 cells. After treatment with F (10[micro]M) for 48 h, the caspase-3 activity in cells was significantly increased, while it was hardly affected by the other compounds. When a larger dose (30 [micro]M) was applied to cells for 48 h, the caspase-3 activity increased approximately three fold for compound F (compared to untreated cells) and, with the exception of D, it also increased for the other compounds at this higher dose (Fig. 5A).

Subsequently, Hoechst 33258 staining was used to confirm cell apoptosis through observation of the nuclear morphology. At a dose of 30 [micro]M and a treatment period of 48 h, chromatin condensation could be observed in the majority of cells treated with compound F (Fig. 5B, panel d), and to a lesser extent in cells treated with compounds A-C and E, while cells treated with compound D appeared unaffected (data not shown). At lower doses of 3 [micro]M and 10 [micro]M, the chromatin condensation was less widespread in cultures treated with compound F (Fig. 5B, panels b and c) and barely visible in cells treated with the other compounds. These results were in accordance with the cell viability results. In addition, the caspase-3 activity and Hoechst 33258 staining were assessed in EA.hy926 cells, whereby no distinct apoptotic changes were noted, indicating the specific effects of these compounds on leukemia cells.

Phosphatidylserine (PS) translocation from the inner to the outer side of the cell membrane is a typical event occurring in the early stage of apoptosis. Annexin V is a phospholipid-binding protein with high affinity for PS and is frequently used to examine the translocation of PS across the plasma membrane. Here, annexin V-FITC/PI double staining was applied to confirm the apoptosis of HL60 cells treated with compound F. Annexin V+/P1- (lower right) and annexin V+/PI+ (upper right) cells were designated as early and late apoptotic populations, as shown in Fig. 5C. After 48 h treatment, few untreated cells or cells treated with 3 [micro]M of compound F were apoptotic. However, increased concentrations of compound F resulted in higher percentages of both early and late apoptotic cells (Fig. 5D), which was in accordance with the results of Hoechst 33258 staining. In addition, the percentages of necrotic cells (annexin V-/PI+, upper left) also increased after drug treatment.

Compound F increased intracellular ROS in HL60 cells

DCFH-DA is a common fluorescent probe that can be deacetylated by intracellular esterase to nonfluorescent DCFH. When ROS exist in the cells, DCFH can be further oxidized to fluorescent compound DCF, whose fluorescence intensity is proportional to the levels of ROS. Here, DCFH-DA was used to assess the effect of compound F on ROS generation in HL60 cells after 24 h treatment. As shown in Fig. 6A, the untreated control cells displayed little green fluorescence, while increasing concentrations of compound F resulted in much stronger signals, with 30 [micro]M compound F producing the strongest intensity. Microplate reader analysis quantitatively confirmed this result (Fig. 6B), whereby compound F resulted in a dose-dependent increase of ROS, with 30 [micro]M raising the ROS relative level 2.6 times compared to untreated control cells.

Compound F triggered mitochondria-dependent apoptosis in HL60 cells

Disruption of the MMP is a key event in mitochondria-dependent apoptosis. Although the JC-1 dye has been widely used to study MMP, its poor water solubility often results in precipitation in aqueous buffers when used at higher concentrations. Therefore, we utilized JC-10 in our study, which is a superior alternative to JC-1. In the mitochondria of cells with a polarized mitochondrial membrane, JC-10 is concentrated and forms reversible red fluorescent JC-10 aggregates. In apoptotic cells, MMP loss leads to a failure to retain JC-10 in the mitochondria and it returns to its monomeric, green fluorescent form. The ratio of the red/green fluorescence intensity ([DELTA][[PSI].sub.m]) is used to measure MMP. Our results showed that a dose-dependent reduction of [DELTA][[PSI].sub.m] occurred in HL60 cells after treatment with compound F for 24 h (Fig. 7A).

Western blotting was then used to determine the changes in location and content of the proteins related to mitochondria-dependent apoptosis, with the results shown in Fig. 7B and C. The release of cyto c from the mitochondria to the cytosol, due to the loss of [DELTA][[PSI].sub.m]. is a limiting factor in the mitochondrial pathway. After 48 h treatment with increasing concentrations of compound F, cyto c expression gradually increased in the cytosol and decreased in the mitochondria of HL60 cells. Caspase-3 is usually present in cells as a zymogen (procaspase) with an inactive form of 35 KDa. Activation of caspase-3 requires proteolytic processing of the zymogen into 17kDa and 12kDa subunits. PARP (113 kDa) is a nuclear enzyme involved in the DNA repair process but it is specifically cleaved into 89 kDa and 24 kDa fragments by activated caspase-3. Here, polyclonal antibodies were used to detect the levels of full-length (35 kDa or 113 kDa) caspase-3 and PARP, and their large fragments (17 kDa or 89 kDa). After 48 h treatment with 30 [micro]M compound F, cleavage of caspase-3 and PARP was clearly visible via western blotting, while the contents of full-length caspase-3 and PARP declined to 49.2% and 41.8%, respectively, compared to the untreated control. Lower concentrations of compounds F (10 [micro]M) also had effects on caspase-3 activation, but to a lesser extent.

Caspases are a family of cysteine proteases that play essential roles in apoptosis. They can be divided into two types: initiator caspases and effector caspases. Caspase-3 is a typical effector caspase, while caspases-8 and -9 are typical initiator caspases with roles in the extrinsic and intrinsic pathways of apoptosis, respectively. Here, the activities of caspases-3, -8 and -9 were determined to fully understand the mechanism of apoptosis induced by compound F in HL60 cells. As shown in Fig. 7D, dose-dependent increases in the activities of caspases-3 and -9 were revealed after 48 h treatment, while 30 [micro]M of compound F increased caspases-3 and -9 by 3.0 and 2.2 fold compared with the untreated control. However, caspase-8 activity remained constant regardless of treatment.

Bcl-2 protein family in the apoptosis of HL60 cells induced by compound F

The Bcl-2 of family proteins are important factors in mitochondria-dependent apoptosis, playing important roles in the regulation of MMP, where Bax and Bcl-2 are representative pro-apoptotic and anti-apoptotic members, respectively. Here, we used qRT-PCR and western blotting to assess the transcription and translation levels of Bax and Bcl-2 genes in HL60 cells after treatment with compound F for 48 h. Treated cells displayed a dose-dependent increase of Bax expression and a decrease of Bcl-2 expression, with similar results for both transcription and translation (Fig. 8A-C). For example, 30|iM of compound F increased the mRNA and protein levels of the Bax gene by 3.4 and 1.9 fold, and suppressed the mRNA and protein levels of Bcl-2 by 48.5% and 59.8%, compared with the untreated control. Consequently, the Bcl-2/Bax ratio decreased gradually as the concentration of compound F increased, as shown in Fig. 8D.

Cell cycle regulation in HL60 cells via compound F

To give a more concrete comprehension of the mechanism of HL60 cell growth inhibition by compound F, the cell cycle was analyzed by flow cytometry after PI staining. After treatment with compound F for 48 h, HL60 cells displayed a dose-dependent increase in the percentage of cells in the sub-G, phase (data not shown), confirming cell apoptosis. The phases of the cell cycle were assessed in more detail by eliminating the sub-[G.sub.1] phase through gating during flow cytometry. From Fig. 9A and B, it is clear that the S phase was significantly enhanced in HL60 cells as a result of treatment with compound F, whereby 30 [micro]M of compound F for 48 h increased the percentage of cells in the S phase by 1.8 fold compared to untreated control cells. Meanwhile, cells in the [G.sub.0]/[G.sub.1] and [G.sub.2]/M phases decreased. These results suggested that compound F delayed cell cycle progression by arresting cells in the S phase.


Leukemia is a group of cancers that exhibit a range of symptoms, including unusual bleeding, bruising and fatigue. The cancers usually begin in the bone marrow and result in large numbers of abnormal white blood cells. Current treatments involve chemotherapy, radiation therapy, targeted therapy and bone marrow transplants. Several promising agents are currently in preclinical or clinical trials for the treatment of leukemia, including monoclonal antibodies, kinase inhibitors, proteasome inhibitors and epigenetic agents Qabbour and Kantarjian 2014; Tasian et al. 2014). Many natural product extracts have also been shown to exert antitumor properties and apoptosis induction in leukemia cells (Al-Salahi et al. 2014; Azadmehr et al. 2013; Guo et al. 2014). The most common four forms of leukemia are acute lymphoblastic leukemia (ALL), acute myeloid leukemia (AML), chronic lymphocytic leukemia (CLL) and chronic myeloid leukemia (CML; Khalade et al. 2010). Consequently, in this study HL60 and K562 cells were used as representatives for AML and CML to assess the cytotoxicity of Catharanthus alkaloids against leukemia cells.

A common problem of chemotherapeutic agents is their indiscriminant toxicity against both cancer and normal cells. Therefore, EA.hy926 human umbilical vein cells, which may be considered as normal cells, were also employed in our cell viability assay. The WST-1 assay results showed that the isolated alkaloids showed growth inhibitory effects against HL60 and K562 cells, but did not affect the growth of EA.hy926 cells, indicating a degree of specificity toward human leukemia cells. Interestingly, compound F was found to be the most potent, while compound D was the least potent, suggesting a structure-activity relationship (SAR). In addition to the WST-1 assay, LDH, which is a stable cytosolic enzyme that catalyzes the oxidation of L-lactate to pyruvate, was used to assess toxicity since it may be released from cells following irreversible cell membrane damage (Furtado et al. 2012). This data confirmed that compound F resulted in the concentration-dependent impairment of leukemia cells, notably by destroying plasma membrane integrity.

Apoptosis or programmed cell death has been recognized as an important physiological event that may be targeted by anticancer agents and cancer therapies (Bai and Wang 2014; Wong 2011). Previous studies have demonstrated that Catharanthus alkaloids induce apoptosis in a variety of malignant cells, including human melanoma, neuroblastoma and breast cancer cells (Huang et al. 2012; Selimovic et al. 2013; Tu et al. 2013). Here, exposure to bisindole alkaloids resulted in typical apoptotic traits in HL60 cells, such as caspase-3 activation and nuclei condensation. Among the alkaloids, compound F exerted the strongest apoptotic effect and acted in a dose-dependent manner, in agreement with the findings from the cytotoxicity assays.

Cell cycle control has also been reported as a major regulatory mechanism for cell growth; therefore, cancer therapies may be used to block the cancer cell cycle (Buolamwini 2000; Malumbres and Barbacid 2005). With regards to Catharanthus alkaloids, vinblastine and vincristine can induce partial arrest of both the [G.sub.0]/[G.sub.1] and [G.sub.2]/M phases in human neuroblastoma cells (Comin-Anduix et al. 2001), and an arrest of the [G.sub.2]/M phase in human breast cancer and epidermoid tumor cells. A prolonged S phase is another common phenomenon of cancer cell cycle delay caused by treatment with naturally occurring alkaloids (Tan et al. 2009; Uadkla et al. 2013). Consequently, in this study we investigated cell cycle progression by flow cytometry after PI staining. Treatment with compound F for 48 h at a concentration of 30 [micro]M induced debris formation in up to 70% of HL60 cells (data not shown), further suggesting the induction of apoptosis. After gating and eliminating the sub-[G.sub.1] phase, S phase induction and [G.sub.0]/[G.sub.1] phase reduction were clearly observed in HL60 cells treated with compound F. Taken together, our findings suggest that compound F inhibits the growth of leukemia cells by the induction of apoptosis, partly due to S phase arrest.

ROS are constantly produced in the human body as a series of oxygen metabolism byproducts, which are removed by antioxidants. Although they play an indispensable role in normal physiological functions, excess concentrations or the overproduction of ROS are harmful due to the damage they inflict on biomacromolecules, such as DNA and proteins (Auten and Davis 2009; Kardeh et al. 2014). They are also believed to function as a double-edged sword in cancer progression and prevention. Moderate levels of ROS maintain essential mechanisms of cancer cell survival, such as proliferation and angiogenesis (Stone and Collins 2002), while high levels of ROS lead to destructive effects on cancer cells through pathways such as apoptosis and autophagy (Chen et al. 2009; Holbrook and Ikeyama 2002). Many alkaloids have been demonstrated to elevate ROS levels and induce apoptosis in human cancer cells (Eom et al. 2010; Ip et al. 2012). Among these, vinorelbine and vincristine promote apoptosis in human lung adenocarcinoma cells and melanoma cells, respectively, through activation of ROS-mediated kinases (Chen et al. 2011; Chiu et al. 2012). In this study, we revealed that treatment with compound F for 24 h increased ROS production in HL60 cells in a dose-dependent manner. Since the stability of the mitochondria is closely related to ROS balance, the characteristics of this important organelle will now be further discussed.

There are two well-known mechanisms regulating cell apoptosis: the extrinsic and the intrinsic pathways. The extrinsic pathway begins with the activation of death receptors via death ligands, such as tumor necrosis factor [alpha] (TNF-[alpha]), the Fas ligand (FasL) and TNF-related apoptosis-inducing ligand (TRAIL; Wallach et al. 2008), while the intrinsic pathway begins with intracellular stress-mediated dysfunction of the mitochondria (Tait and Green 2010). Caspases in mammals can be broadly classified by their roles in apoptosis (caspases-3, -6, -7, -8 and -9) and inflammation (caspases-1, -4, -5, -11 and -12). With regards to apoptosis, caspases-8 and -9 are initiator caspases in the extrinsic and intrinsic pathways, respectively, while caspase-3, -6 and -7 are executioner caspases in both pathways (Me Ilwain et al. 2013; Philchenkov et al. 2004). In our study, treatment of HL60 cells with compound F resulted in enhanced caspases-3 and -9 activities, while caspase-8 was not affected, suggesting that apoptosis occurred mainly through the mitochondrial pathway.

The mitochondria generate adenosine triphosphate (ATP) as an energy source for cellular processes and have global regulation over calcium flux and the redox states. The mitochondrial properties of cancer cells, such as mitochondrial DNA, MMP and oxygen/glucose consumption, have been frequently recognized as targets for chemotherapeutic agents or as a means to overcome chemotherapeutic resistance (Don and Hogg 2004). A great many natural alkaloids have been identified through cancer research as affecting the cellular mitochondria, and inducing apoptosis and autophagy (Gong et al. 2012; Siedlakowski et al. 2008). The effects of such alkaloids on the mitochondria include alternations in morphology, biogenesis and changes in the metabolic pathways, as well as alternations in cross-talk between the endoplasmic reticulum and mitochondria (Urra et al. 2013). In the intrinsic mitochondrial-dependent apoptotic pathway, disruption of the mitochondrial outer membrane integrity is followed by a reduction of [DELTA][[PSI].sub.m] and the translocation of cyto c and other pro-apoptotic molecules from the mitochondria to the cytosol (Thangam et al. 2014). Once released, cyto c binds with apoptotic protease activating factor-1 (Apaf-1) and ATP, which then bind to procaspase-9 to create a protein complex known as an apoptosome. The apoptosome then activates caspases-3 and -9, leading to the cleavage of PARP (Green and Kroemer 2004). Thus, we further examined the above-mentioned apoptotic pathway in drug-treated HL60 cells through the MMP, the translocation of cyto c, and the cleavage of caspase-3 and PARP. PARP catalyzes the poly(ADP-ribosyl)ation of many nuclear proteins, while the cleavage of PARP is known to prevent the depletion of energy and the repair of DNA strand breaks, which are required for the occurrence of apoptosis (Boulares et al. 1999). As expected, after 24 h treatment with compound F (1-30 [micro]M), HL60 cells showed a dose-dependent loss of [DELTA][[PSI].sub.m], while translocation of cyto c, and the cleavage of caspase-3 and PARP were affected after 48 h.

The Bcl-2 family is a group of well-studied proteins controlling the MMP during cell apoptosis and they can be pro-apoptotic (Bax, Bak, etc.) or anti-apoptotic (Bcl-2, Bcl-xL, etc.). Bcl-2 and Bax have similar structures to multi-Bcl-2 homology (multi-BH) domain proteins, but with opposite effects in terms of apoptosis. Bcl-2 is over-expressed on the OMM of leukemia cells and acts on apoptotic signals to maintain the stability of the mitochondria (Ola et al. 2011). Bax, which is usually located in the cytosol, transfers to the OMM during apoptosis and oligomerizes to form pores facilitating the release of pro-apoptotic molecules (Reed 2006). Our data showed that compound F increased Bax expression and decreased Bcl-2 expression in a dose-dependent manner at both the transcriptional and translational levels. The resulting reduction of the Bcl-2/Bax ratio was in accordance with previous reports on cancer cell apoptosis induced by other natural alkaloids (Mantena et al. 2006; Zheng et al. 2013).


In conclusion, we isolated a new bisindole alkaloid from C. roseus, called cathachunine (compound F), which exerted an antitumor effect on human leukemia cells with much lower cytotoxicity toward normal human endothelial cells. Compared to other C. roseus alkaloids, the anti-proliferation and pro-apoptosis abilities of compound F were increased, and the induction of apoptosis in HL60 cells as a result of treatment was shown to occur through an ROS-dependent mitochondria-mediated intrinsic pathway regulated by the Bcl-2 protein family. In the future, efforts should be focused on this alkaloid's antitumor effect in animal models, and its signal transduction pathway should be investigated in more detail.


Article history:

Received 26 May 2015

Revised 1 March 2016

Accepted 4 March 2016

Abbreviations: ATCC, American type culture collection; DCFH-DA, 2', 7'-dichlorodihydrofluorescein diacetate; DEPC, diethylpyrocarbonate; HL60, human promyelocytic leukemia cells; LDH, lactate dehydrogenase; MMP, mitochondrial membrane potential; OMM, outer mitochondrial membrane; PARP, poly-ADP ribose polymerase; PI, propidium iodide; PVDF, polyvinylidene difluoride; ROS, reactive oxygen species; RP-HPLC, reverse phase high performance liquid chromatography.

Conflict of interest

The authors declare no competing financial interest. Acknowledgments

This work was supported by the Tianjin Applied Basic and Cutting-edge Technology Research Program (Grant numbers 13JCZDJC28600 and 14JCYBJC29000), the National Natural Science Foundation of China (Grant numbers 81403059, 81202396, 81503215), the Science Foundation of Shenzhen, China (Grant numbers JCYJ20150525092941057, JCYJ20130326112757843, KQCX20140522111508785, SFG(2013) 180), the Natural Science Foundation of Shenzhen University, (Grant number 201410), and the Fundamental Research Funds for the Central Universities (Grant number 11615305).


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Xiao-Dong Wang (a,c,d,1), Chen-Yang Li (a,c,d,1), Miao-Miao Jiang (b), Dong Li (b), Ping Wen (e), Xun Song (a,c,d), Jun-Da Chen (a), Li-Xuan Guo (a), Xiao-Peng Hu (a,c,d), Guo-Qiang Li (f), Jian Zhang (a,c,d), Chun-Hua Wang (b), *, Zhen-Dan He (a,c,d), **

(a) Department of Pharmacy, School of Medicine, Shenzhen University, Shenzhen 518060, Guangdong, PR China

(b) Tianjin Key Laboratory of Modern Chinese Medicine, Tianjin University of Traditional Chinese Medicine, Tianjin 300193, PR China

(c) Institute of Biotherapy, Shenzhen University, Shenzhen 518060, Guangdong, PR China

(d) Engineering Laboratory of Shenzhen Natural Micromolecule Innovative Drugs, Shenzhen University, Shenzhen 518060, Guangdong, PR China

(e) Business Technology Department, Shenzhen Institute for Drug Control, Shenzhen 518057, Guangdong, PR China Experiment and Technology Center, Jinan University, Guangzhou 510632, Guangdong, PR China

* Corresponding author. Tel./fax: +86 22 27386453.

** Corresponding author at: Department of Pharmacy, School of Medicine, Shenzhen University, Shenzhen 518060, Guangdong, PR China. Tel.: +86 755 86671916; fax: +86 755 86671906.

E-mail addresses: (C.-H. Wang), (Z.-D. He).

(1) These authors contributed equally to this work.

Table 1
Sequences of primers used in the paper.

Name      Sequence


Table 2
NMR data of compound F (in CD[Cl.sub.3] [sup.1]H NMR
for 400 MHz, [sup.13]C NMR for 100 MHz, J in Hz).

Position        H                     C

2               3.63 s                 81.8
3a              3.24 overlapped        50.3
3b              2.74 overlapped
5a              3.22 overlapped        50.2
5b              2.39 overlapped
6a              2.17 m                 44.9
6b              1.73 m
7               --                     53.7
8               --                    123.9
9               6.52s                 122.7
10              --                    119.7
11              --                    158.1
12              6.08 s                 93.6
13              --                    153.2
14              5.67 dd (9.6, 4.0)    121.9
15              5.46 d (9.6)          136.0
16              --                     77.3
17a             1.95 s                 38.0
18              0.89 t (7.6)            8.6
19a             1.31 m                 34.6
19b             1.12 m
20              --                     37.2
21              2.48 S                 66.9
COOC[H.sub.3]   --                    173.6
COOC[H.sub.3]   3.78 s                 52.2
N-C[H.sub.3]    2.71 s                 37.9
Ar-OCHj         3.78 S                 55.9
2'              --                    132.6
3'a             3.99 dd (10.4 1.2)     50.2
3'b             3.06 dd (10.4 2.0)
5'a             4.27 dd (13.6, 2.8)    43.2
5'b             2.99 t (13.6)
6'a             3.29 t (15.6)          25.9
6'b             3.15 dd (15.6 3.6)
7'              --                    114.9
8'              --                    129.1
9'              7.53 d (7.6)          118.3
10'             7.13 m                119.4
11'             7.16 m                123.0
12'             7.11 d (7.6)          110.6
13'             --                    134.8
14'             1.54 m                 35.7
15'a            2.84 m                 32.4
15'b            2.46 overlapped
16'             --                     55.6
17'a            2.42 dd (15.2, 4.8)    40.4
17'b            1.91 overlapped
20'             --                    179.0
COOC[H.sub.3]   --                    174.3
COOC[H.sub.3]   3.58 S                 52.6

Table 3
[IC.sub.50] values ([micro]M) of compounds on cell growth
inhibition after 48 h treatment. (95% confidence interval).

[IC.sub.50]        HL60                K562              EAhy926

A            64.3 [+ or -] 3.8  58.5 [+ or -] 13.3         >100
B            56.2 [+ or -] 6.2  57.8 [+ or -] 11.0         >100
C            72.8 [+ or -] 7.5  76.9 [+ or -] 5.3          >100
D            >100               >100                       >100
E            62.7 [+ or -] 7.4  69.9 [+ or -] 11.8         >100
F             9.1 [+ or -] 0.7   9.3 [+ or -] 1.8   84.5 [+ or -] 11.6
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Author:Wang, Xiao-Dong; Li, Chen-Yang; Jiang, Miao-Miao; Li, Dong; Wen, Ping; Song, Xun; Chen, Jun-Da; Guo,
Publication:Phytomedicine: International Journal of Phytotherapy & Phytopharmacology
Article Type:Report
Geographic Code:9CHIN
Date:Jun 1, 2016
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