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Abstract

Pyropolyporus fomentarius, a fungus of the polyporaceae family, has been used in the treatment of various diseases, such as gastroenteric disorder, hepatocirrhosis, oral ulcer, inflammation, and several cancers. This study was conducted to investigate the compositions and cell growth inhibition effects of P. fomentarius chloroform (CHCl₃) fraction and to clarify the possible mechanisms. Gas chromatography–mass spectrometry analysis was performed to investigate the composition of the P. fomentarius CHCl₃ fraction. Cell viability was measured using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay. Cell membrane damage was evaluated with a scanning electron microscope and flow cytometry following propidium iodide and bis-(1,3-dibarbituric acid)-trimethine oxanol staining. Apoptosis was analyzed using annexin V-PE/7-amino-actinomycin D (7-AAD) staining. Generation of intracellular calcium ion (Ca²⁺), reactive oxygen species (ROS), and changes of mitochondrial membrane potential (Δψ _m) were detected by flow cytometry using fluo 3-acetoxymethyl ester, 2′,7′-dichlorofluorescin-diacetate, and rhodamine 123. Our obtained data indicate that P. fomentarius CHCl₃ fraction could inhibit proliferation of K562 cells depending on both the dosage and the incubation time, cause cell membrane damage, influence intracellular [Ca²⁺]_i variation, promote the yield of ROS, decrease the level of Δψ _m, and initiate the apoptotic response in K562 cells.

Keywords

gas chromatography–mass spectrometry antitumor K562 cells

Introduction

Pyropolyporus fomentarius (L. ex Fr.) Teng, also called Fomes. fomentarius, is a fungus of the Polyporaceae family, belonging to the genus of Fomes. This herb has a long history of medicinal use. For instance, it was applied as hemostatic dressings and bandages during the 18th and 19th centuries.¹ The same polypore together with the birch polypore (Piptoporus betulinus), which has had a variety of medicinal and other uses, was found with the body of the famous 5300-year old “Ice Man” in a glacier of Italian Alps in 1991.^2,3 In Japan, it is also used to make a popular drink, which is believed to be a tonic and to have preventive effects against wide-ranging diseases.⁴ As a traditional Chinese medicine, this mushroom has been applied in the treatments of various diseases, such as gastroenteric disorder, hepatocirrhosis, oral ulcer, inflammation, and several cancers.⁵

The traditional use of the polypore has been demonstrated by modern pharmacological studies. Chen et al.’s⁵ investigation illustrated that the ethanol extract of P. fomentarius had a direct antiproliferative effect on human gastric cancer cell lines SGC-7901 and MKN-45. Recently, a study described moderate cytotoxic effect of sterols isolated from ethanol extract of P. fomentarius.⁶ Besides, Lu et al.⁷ reported that chloroform (CHCl₃) extract of P. fomentarius strongly inhibited the proliferation of Hela cells. Our previous screening studies indicated that the CHCl₃ fraction, from the ethanol extract of the P. fomentarius, caused more significant cell damages in K562 cells than either the petroleum ether or n-butyl alcohol fractions did (data unpublished).

However, to the best of our knowledge, little research on anticancer mechanisms of the P. fomentarius CHCl₃ extract has been performed before. Therefore, this study was conducted to investigate the cell growth inhibition effects of the P. fomentarius CHCl₃ fraction and its underlying mechanisms.

Materials and methods

Plant materials

The sporophores of P. fomentarius was collected from Pinghe-liang, south of QinLing Mountains, Shaanxi province, China (latitude, 33°27′ N; longitude, 108°30′ E; altitude, 2305 m). It was identified by Prof. Yaping Xiao in the Ministry of Education, Key Laboratory for Medicinal Plant Resource and Natural Pharmaceutical Chemistry, Shaanxi Normal University, Xi’an, Shaanxi, People’s Republic of China.

Preparation of the CHCl₃ extract

Crushed plant material was soaked in 95% ethanol (EtOH, w/v) at 45°C and extracted three times (each for 30 min) using ultrasonic-assisted extraction. After filtration through filter paper, the filtrate was vacuum concentrated and freeze-dried. The ethanol extract was then homogenized in water, followed by extraction with petroleum ether, CHCl₃ and n-butyl alcohol, successively. Under reduced pressure, the various polarity fractions were obtained by evaporation. Then, the CHCl₃ fraction was dissolved in 95% EtOH to get a stock solution of 20 mg ml⁻¹ and was passed through 0.22 µm filter before use. Note that in this work, the CHCl₃ fraction from P. fomentarius was called PFCF for short.

Component analysis by GC/MS

To investigate the composition of the PFCF, gas chromatography–mass spectrometry (GC-MS) analysis was performed with a 7890A GC/5975C MS system (Agilent Technologies, Santa Clara, California, USA) and fitted with a fused silica capillary column (HP-5MS, 30 m × 25 mm internal diameter, 0.25 μm film thickness; Agilent J&W Scientific, Folsom, California, USA). One microliter of the sample was analyzed with a split ratio of 10:1 (v/v). The programs of temperature rise were followed by initial temperature of 50°C for 2 min, raised at 10°C min⁻¹ to 200°C, maintained for 2 min, and then raised to 250°C at 5°C min⁻¹, maintained for 6 min, and then it was raised to 280°C at 10°C min⁻¹ and maintained isothermally for 5 min. The injector temperature was set at 280°C, and the interface temperature was at 150°C. The MS source helium was used as the carrier gas at a flow rate of 1 ml min⁻¹. The ionization source temperature was 250°C. MS was determined by full-scan method with range from 50 to 600 (m/z). Metabolites were identified by comparison with the National Institute of Standards and Technology (NIST) Mass Spectral Search Program 2008 database (version 2.0, FairCom Co., Columbia, Missouri, USA).

Cell culture

K562 cells (human chronic myelogenous leukemia cell line) were obtained from the Chinese Academy of Medical Sciences (Beijing, China). Cells were grown in Roswell Park Memorial Institute (RPMI) 1640 medium, supplemented with 10% fetal bovine serum and 1% penicillin, streptomycin, l-glutamine, and incubated at 37°C in a humidified 5% carbon dioxide atmosphere. Cells in the exponential phase were used in our experiments.

Isolation of PBMC

Peripheral blood mononuclear cells (PBMC) cells were isolated with the lymphocyte separation kit (Applygen Technologies Inc., Beijing, China) from healthy donors in the hospital of Shaanxi Normal University, China. The whole procedures were performed according to the instructions provided by the manufacturer. The viability of the obtained PBMC cells was evaluated using trypan blue exclusion test and in experiment the viability was above 95%.

Assessment of cell viability after PFCF treatment

To detect the effect of PFCF on K562 cell proliferation, at a cell density of 1 × 10⁵, cells were incubated in 96-well plates. Different concentrations of PFCF (0, 10, 20, 30, 40, 50, 60, 70, and 80 μg ml⁻¹) were added into the cells and then incubated for 24, 48, and 72 h. Cell viability of each group was determined by using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (Sigma-Aldrich, St Louis, Missouri, USA) assay.⁸ The absorbance values at 570 nm were recorded using a microplate reader (Bio-Tek ELX800, Winooski, Vermont, USA). Cell viability of PFCF-treated group was then acquired by comparing with the control group. The group containing 95% EtOH was used to indicate the effects of EtOH on cells.

In this experiment, PBMC cells were used as normal cells to evaluate the cytotoxicity of PFCF. K562 cells and PBMC cells suspension (100 μl) were seeded at a density of 1 × 10⁵ cells ml⁻¹ in 96-well culture plates, respectively, and incubated with PFCF at different doses (0, 20, 40, 60, and 80 μg ml⁻¹) for 24 h. Two types of cells were then harvested and stained with trypan blue (0.4%). Using a hemocytometer, the stained and unstained cells were counted. Cell viability was determined according to the following equation: cell viability = number of unstained cells/number of cells in the control group × 100%. Each experiment was repeated for at least three times.

Membrane integrity testing

Cell membrane integrity can be monitored using propidium iodide (PI) staining. PI, a nucleic acid binding dye, cannot penetrate the membrane of viable cells but can enter cells through a damaged cell membrane. Therefore, when cell membrane integrity is poor, the fluorescence of PI will increase significantly.

Cells (1 × 10⁵ cells ml⁻¹) were incubated with the indicated concentrations of PFCF for 6 h, collected by centrifugation, washed two times with phosphate-buffered saline (PBS), then stained with 5 μg ml⁻¹ PI (Sigma-Aldrich, St Louis, Missouri, USA) for 5 min in the dark and analyzed by flow cytometry (Guava easyCyte 8HT; Millipore Corp., Bedford, Massachusetts, USA). Using Flow Cytometry Analysis (FCS) Express V3, histograms were analyzed.

The measurement of cell membrane potential

Bis-(1,3-dibarbituric acid)-trimethine oxanol (DiBAC₄(3)) is used as a dye for monitoring changes in the plasma membrane potential.⁹ The depolarized membrane potential could increase the distribution of DiBAC₄(3) in the cytosol and then the intracellular fluorescence intensity will rise subsequently. In contrast, the hyperpolarized membrane potential will decrease the intracellular fluorescence intensity.

After treated by PFCF at various concentrations for 4 h, K562 cells were harvested by centrifugation, washed with PBS, stained with 5 μmol l⁻¹ DiBAC₄(3) (D8189; Sigma-Aldrich, St Louis, Missouri, USA) at room temperature and then detected by flow cytometry. Histograms were analyzed with FCS Express V3.

SEM observation

After 24 h, following 40 μg ml⁻¹ PFCF treatment, cells of each group were fixed by 2.5% glutaraldehyde, washed with PBS, dehydrated by graded alcohol, displaced, dried at the critical point, evaporated with gold, and then observed under a SEM (S-3400 N, Hitachi, Tokyo, Japan).

Annexin V-PE/7-AAD staining experiment

Annexin V-PE/7-amino-actinomycin D (7-AAD) double staining was used to detect the rate of apoptosis. During the early stage in apoptosis, annexinV-PE detects the translocation of phosphatidylinositol in the outer cell membrane, while in the late-staged apoptotic and dead cells, 7-AAD can enter the cell.

Apoptotic cells were measured using Guava Nexin Reagent (4700–1140, Millipore, Billerica, Massachusetts, USA). The assay was performed according to the instructions provided by the manufacturer. Briefly, K562 cells (1 × 10⁵ cells ml⁻¹) were treated with 40 µg ml⁻¹ of PFCF for 24 h at 37°C. Cells were collected by centrifugation, resuspended with 100 µl binding buffer, and then stained with 100 µl of Guava Nexin Reagent for 20 min at room temperature in the dark. Finally, samples were detected by flow cytometry, and the apoptotic indexes were analyzed using FCS Express V3.

Measurement of [Ca²⁺]_i

Flu 3-acetoxymethyl ester (Fluo 3-AM; F312; Dojindo Laboratories, Kumamoto, Japan) was used to detect the intracellular calcium (Ca²⁺) ion concentration. Fluo 3-AM has long been used as the nonfluorescent acetoxymethyl ester, which is cleaved and turns to Fluo-3 in cells. Fluo-3, a Ca²⁺ indicator, can bind to Ca²⁺ ions and present strong fluorescence.

After treated with PFCF for 4 h and 8 h, both the control and treated cells were collected by centrifugation, resuspended with PBS, loaded with 2 μmol l⁻¹ Fluo 3-AM for 30 min at 37°C, washed with PBS, and then immediately analyzed by flow cytometry.

Detection of intracellular ROS generation

We use the sensitive fluorescent probe 2′,7′-dichlorofluorescein-diacetate (DCFH-DA; Invitrogen, Carlsbad, California, USA) to detect the changes of intracellular ROS. DCFH-DA could easily pass through cell membrane, hydrolyzed by intracellular esterases, and then turn to non-fluorescent DCFH. Since DCFH can be subsequently oxidized and form highly fluorescent oxidized DCFH (DCF) in the presence of reactive oxygen species (ROS), the generation of intracellular ROS will be proportional to the fluorescence intensity.

After incubated with PFCF at various concentrations (0, 20, 60 µg ml⁻¹) for 4 h, cells (1 × 10⁵ cells ml⁻¹) were harvested, stained with 10 μM DCFH-DA, and incubated at 37°C for 30 min in the dark. After washed with PBS, samples were immediately analyzed by flow cytometry. Excitation wavelength was 488 nm and emission wavelength was 530 nm. Histograms were analyzed using FCS Express V3.

Examination of the Δψ _m

We use the fluorescent probe rhodamine 123 (Rh123, Sigma) to detect the changes of mitochondrial membrane potential (Δψ _m). Rh123 selectively enters mitochondria normally and can be retained in them.¹⁰ Once Δψ _m is decreased, the amount of Rh123 will subsequently decline.

Incubated with PFCF at various concentrations (0, 20, and 60 µg ml⁻¹) for 12 h, cells (1 × 10⁵ cells ml⁻¹) were collected and then stained for 20 min with 2 µg ml⁻¹ Rh123. After washed with PBS, cells were immediately analyzed by flow cytometry. Excitation wavelength was 488 nm and emission wavelength was 530 nm. Histograms were analyzed using FCS Express V3.

Statistical analysis

Data are expressed as mean ± standard deviation of at least three independent experiments. Statistical analysis was performed by one-way analysis of variance. Statistical significance was established at a value of p < 0.05.

Results

Chemical compounds in PFCF

Upon GC-MS analysis, chemical composition of the main constituents from PFCF was identified in Table 1. The main constituents were considered “identified,” when their mass spectral fit values were at the default value of 90% or above. A total of 13 constituents were identified in the extract and the chemical structures of the identified compounds are shown in Figure 1. For the first time, our study reports the chemical compositions of the CHCl₃ extract of P. fomentarius fruit body.

Figure 1.

The structures of 13 identified compounds in CHCl₃ fraction. CHCl₃: chloroform.

Table 1.

Identification of PFCE metabolites using GC-MS analysis.

Retention time (min)	Compound	Peak area (%)	Molecular formula
17.457	Tetradecanal	1.4	C₁₄H₂₈O
19.491	n-Hexadecanoic acid	6.63	C₁₆H₃₂O₂
19.591	1,2-Benzenedicarboxylic acid, butyl 2-methylpropyl ester	3.60	C₁₆H₂₂O₄
20.475	Hexadecanal	7.91	C₁₆H₃₂O
21.875	2-Nonadecanone	1.62	C₁₉H₃₈O
22.358	9,12-Octadecadienoic acid (Z, Z)-	5.51	C₁₈H₃₂O₂
22.45	9-Octadecenoic acid, (E)-	7.72	C₁₈H₃₄O₂
22.517	13-Octadecenoic acid, (E)-	2.87	C₁₈H₃₄O₂
23.909	Oxirane, heptadecyl-	8.73	C₁₉H₃₈O
27.293	(Z)-13-Docosen-1-ol	2.13	C₂₂H₄₄O
31.861	(9Z)-9,17-Octadecadienal	3.99	C₁₈H₃₂O
42.238	Ergosterol	1.45	C₂₈H₄₄O
42.496	Ergosta-7,22-dien-3-one	1.13	C₂₈H₄₄O

PFCE: chloroform fraction from P. fomentarius; GC-MS: gas chromatography–mass spectrometry.

Effect of PFCF on cell viability

K562 cells at a density of 1 × 10⁵ cells ml⁻¹ were treated with PFCF with specific doses for 24, 48, and 72 h. As shown in Figure 2(a), PFCF inhibited K562 cell proliferation in a dose- and time-dependent manner. Compared with the control, the extract (10 μg ml⁻¹) showed no cell damage (p > 0.05), while there were significant cell damages when PFCF doses were 20 μg ml⁻¹ or above, and the prolonged incubation enhanced the viability loss. Of note, EtOH didn’t show obvious effect on cancer cells proliferation under the maximum dose range.

Figure 2.

Cytotoxicity effect of CHCl₃ fraction on K562 cells and PBMC. (a) K562 cells were treated with 0, 10, 20, 30, 40, 50, 60, 70, and 80 μg ml⁻¹ of PFCF for 24, 48, and 72 h; (b) PBMC and K562 cells were incubated at the same experimental conditions for 24 h. Solvent ethanol effect was also suggested. Each value is expressed as a mean ± SD of three independent experiments. *p < 0.05 and **p < 0.01 versus the control. CHCl₃: chloroform; PBMC: peripheral blood mononuclear cell.

Compared with K562 cells, PBMC cells showed no significant cell death in the experimental groups except the 80 μg ml⁻¹ group (Figure 2(b)).

Plasma membrane damage measurement

Firstly, combined with flow cytometry, PI staining was used to evaluate cell membrane integrity after treated by PFCF. Images in Figure 3 show that when PFCF concentrations are 20 and 60 μg ml⁻¹, the percentage of cells with higher PI fluorescence obviously increases to 86.13% and 85.4% at 6 h, while in the control group, such data is 5.4%.

Figure 3.

Effects of CHCl₃ fraction on the cell membrane integrity of K562 cells at 6 h after treatment. (a) cells were treated with CHCl₃ fraction (a) control; (b) 20 μg ml⁻¹; (c) 60 μg ml⁻¹, the percentage of fluorescent cells in each group was shown. (b) The results represent the mean ± SD of three independent experiments. **p < 0.01: statistically significant difference with control. CHCl₃: chloroform; PBMC: peripheral blood mononuclear cell.

Secondly, we measured the membrane potential of K562 cells after PFCF treatment. As shown in Figure 4, compared with the control, cells in the treated groups showed greatly dose-dependent cell membrane damage (p < 0.01). The percentage of cells with D_iBAC₄(3) fluorescence were 2.57%, 42.9%, and 79.04% when PFCF concentrations were 0, 20, and 60 μg ml⁻¹, respectively. The result indicated a depolarization in the membrane potential after treated with PFCF.

Figure 4.

Membrane potential changes in K562 cells. Cells were treated with CHCl₃ fraction at 0, 20, and 60 μg ml⁻¹ for 4 h, labeled with DiBAC and the fluorescence intensity in individual cells was detected by flow cytometry. Data are mean ± SD of three independent experiments. **p < 0.01: statistically significant difference with control. CHCl₃: chloroform; DiBAC: bis-(1,3-dibarbituric acid)-trimethine oxanol.

Finally, membrane damage effect was morphologically observed under SEM (Figure 5). In the control group, cells appeared round in shape with abundant microvilli, while cells treated with 40 μg ml⁻¹ PFCF showed a significant decrease in the number of microvilli and shrunk obviously with relatively smooth surfaces. Moreover, some small convex structures were observed in PFCF group.

Figure 5.

SEM images of K562 cells at 24 h with 0 and 40 μg ml⁻¹ CHCl₃ fraction treatments. Magnification of the images was ×5000. Scale bars: 10 mm. SEM: scanning electron microscopy; CHCl₃: chloroform.

Apoptosis assessment

Using annexin V-PE/7-AAD double staining, we analyzed the exposure of phosphatidylserine as a marker of apoptosis. In Figure 6, after 24 h treatment, the viable cells were 96.4% in the control group; the cell population in the early stage of apoptosis (lower right) was 1.4%; the cell population in the late stage of apoptosis (upper right) was 1.9%. However, compared with the control, in the treated group the percentages of cells with annexin V-positive staining increased gradually (p < 0.01) from 3.3% to 36.8%, suggesting that PFCF could induce apoptosis in K562 cells.

Figure 6.

Dot plots of annexin V and 7-AAD uptake after different treatments in K562 cells. (a) Cells were analyzed at 24 h post-treatment by flow cytometry (a′) control and (b′) 40 μg ml⁻¹ CHCl₃ fraction. (b) The ratio of annexin V-positive and -negative cells are represented as means ± SD of three independent experiments. 7-AAD: 7-amino-actinomycin D; CHCl₃: chloroform.

Changes in intracellular [Ca²⁺]_i

As shown in Figure 7, at 4 or 8 h after treatment, cells exposed to 20 μg ml⁻¹ PFCF did not show obvious changes in [Ca²⁺]_i. However, cells after treatment with 60 μg ml⁻¹ PFCF showed increased [Ca²⁺]_i concentration compared with the control (p < 0.01).

Figure 7.

Time course of changes in [Ca²⁺]_i after CHCl₃ fraction treatment. Cells were treated with CHCl₃ fraction (a) 4 h, (a′) control; (b′) 20 μg ml⁻¹; (c′) 60 μg ml⁻¹ and (b) 8 h, (a′) control; (b′) 20 μg ml⁻¹; (c′) 60 μg ml⁻¹, both the control and treated cells were loaded with Fluo 3-AM and analyzed by flow cytometry, and percentage of fluorescent cells in each group was shown. (c) Data are mean ± SD of three independent experiments. **p < 0.01: statistically significant difference with control at each time point. [Ca²⁺]_i: calcium ion concentration; CHCl₃: chloroform; Fluo 3-AM: fluo 3-acetoxymethyl ester.

Detection of intracellular ROS generation and Δψ _m loss

Intracellular ROS was analyzed using the flow cytometer with DCFH-DA staining. Figure 8 shows that compared with the control, more obvious spectral shift of DCF fluorescence curve form left to the right in a dose-dependent manner after PFCF treatment, demonstrating that PFCF could markedly increase the level of intracellular ROS.

Figure 8.

CHCl₃ fraction stimulated intracellular ROS generation in K562 cells. (a) Cells were treated with CHCl₃ fraction for 4 h at (a′) control; (b′) 20 µg ml⁻¹; (c′) 60 µg ml⁻¹, and labeled with DCFH-DA. The fluorescence intensity of the oxidized product DCF in individual cells was detected by flow cytometry, the percentage of fluorescent cells in each group was shown. (b) Each value is expressed as a mean ± SD of three independent experiments. *p <0.05; **p <0.01: versus the control. CHCl₃: chloroform; ROS: reactive oxygen species; DCFH: 2′,7′-dichlorofluorescin; DA: diacetate

After incubation for 12 h, a significant Δψ _m loss could be detected in the treated group with the increasing dose of PFCF (Figure 9). The mean fluorescent intensity of Rh123 in the control group was 94.13, which was statistically much higher than that in the PFCF group (72.46 and 59.72; p < 0.01). The obtained results imply that the elevated ROS and decreased Δψ _m may both act as indicators in cell apoptosis.

Figure 9.

Effect of CHCl₃ fraction on the mitochondrial membrane potential of K562 cells. (a) Cells were treated with CHCl₃ fraction for 12 h at (a′) control; (b′) 20 µg ml⁻¹; (c′) 60 µg ml⁻¹, labeled with Rh123, and analyzed by flow cytometry. Histograms show number of cell channel (vertical axis) versus Rh123 fluorescence (horizontal axis). (b) The results represent the mean ± SD of three independent experiments. **p < 0.01: statistically significant difference with untreated cells. CHCl₃: chloroform; Rh123: rhodomine 123.

Discussion

Numerous cytotoxic polypore metabolites with lower molecular weight are related to the cytotoxic, antitumor, and immunomodulatory abilities of polypore extracts.^11,12 For example, hispidin, isolated as a naturally occurring styrylpyrone from Inonotus hispidus, is cytotoxic to cancerous cells while is little toxic in normal cells.¹³ Studies have shown that zhankuic acid A exhibited its cytotoxic activity against P-388 murine leukemia.¹⁴ Thus, the major objective of this work is to determine the chemical compositions of PFCF by GC-MS to investigate the in vitro antitumor effects and explore its underlying molecular mechanisms.

GC-MS analyses of the CHCl₃ extract have shown the presence of alkanes, terpenoid, ketones, fatty acid, aldehyde, and phytosterol (Figure 1). Because the chemical properties of the identified compounds varied widely, we couldn’t tell which individual compound was the major cause of the observed antitumor effects or whether such results were mainly due to the mixture of certain compounds. According to the results of gas chromatography, the nonvolatile compounds are not detected, which might have contributed to this capacity, so further investigations are required to identify the active ingredients which may be responsible for the antitumor activity of PFCF.

It has been proven that the P. fomentarius extract could inhibit the proliferation of tumor cells.⁵ In our article, data indicate that PFCF inhibits the proliferation of K562 cells depending on both the dosage and the incubation time. Meanwhile, K562 cells turned out to be much more sensitive to PFCF than PBMC cells (Figure 2(b)). This is partly in agreement with Beattie et al.’s report;¹⁵ their results indicated that 15 Australian fungi species showed low or no cytotoxic activity against human healthy cells and some had significant (Omphalotus nidiformis, Cordyceps Cranstounii, and Cordyceps gunnii) and selective (Xerula mundroola, Leratiomyces ceres, and Hypholoma fasciculare) activity against cancer cells. The data suggest that the P. fomentarius might be of great potential in cancer treatment.

In this study, we observed that significant plasma membrane damages are caused by PFCF in K562 cells (Figures 3 and 4). It was reported that cell membrane could just maintain homeostasis and regulate itself within a certain extent. Once the stimulus is beyond the threshold, cell membrane will be injured and physicochemical properties may be changed, which then drive cells toward apoptotic or necrotic death.¹⁶ Moreover, the PFCF-induced cell membrane damages were further evidenced by SEM observation (Figure 5). Changes on the surface of cell membrane affect cellular functions directly and may eventually lead to cell death. Thus, the results of membrane evaluation were consistent with the above-presented cell viability trends.

In terms of many types of cancers, defects in apoptosis are the common characters and are also essential in the resistance to different cancer therapies.^17
–19 Thus, apoptotic pathways are relevant targets of cancer therapies.¹⁸ Over the past two decades, bioactive compounds in a growing number have been extracted from various traditional Chinese medicines, while some of them are related to apoptotic induction.²⁰ In this study, we found that PFCF could induce apoptosis in K562 cells (Figure 6), implying that such extract has great potential in screening anticancer drugs.

Nowadays, studies on apoptosis pathways that were induced by compounds extracted from plants mainly focus on mitochondria/intrinsic pathway. Both ROS- and Ca²⁺-mediated mechanisms have been reported to play vital roles in targeting cancer cells.^21

–25 Besides, mitochondria were also involved in cell apoptosis triggered by various stimuli.^26,27 Therefore, we examined the changes of intracellular [Ca²⁺]_i, ROS formation, and Δψ _m after PFCF treatment in K562 cells. As can be seen from Figure 7, after treatment, PFCF at 60 µg ml⁻¹ could obviously enhance [Ca²⁺]_i. It is well established that cellular [Ca²⁺]_i overload may cause cytotoxicity and result in cell death.²⁸ Additionally, compared with the control group, the generation of intracellular ROS dramatically increased in a concentration-dependent manner (Figure 8). It has been reported that high levels of oxidative stress are toxic in cancer cells.²⁹ Such data suggest that ROS- and Ca²⁺-mediated mechanisms might be both involved in our system when treated with PFCF in K562 cells. Δψ _m, as detected by flow cytometry with RH123, gradually decreased with the increasing PFCF doses (Figure 9), indicating that mitochondria were seriously damaged after PFCF treatment. Thus, we assume that PFCF-induced cell apoptosis might be ROS- and Ca²⁺-mediated mitochondria dependent.

In summary, our preliminary results, for the first time, evaluated the cytotoxicity of the P. fomentarius CHCl₃ fraction in K562 cells, demonstrated that PFCF could cause cell membrane damage, influence intracellular [Ca²⁺]_i, promote the yield of ROS, decrease the level of Δψ _m, and then initiate apoptosis.

Footnotes

Conflict of interest

The authors declared no conflicts of interest.

Funding

This work was supported by National “Twelfth Five-Year” Plan for Science and Technology Support (2011BAI06B05), National Natural Science Foundation for Young Scholars of China (81202957) and Natural Science Foundation of Shaanxi Province, China (2014JM4137).

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Chemical compositions and antiproliferation activities of the chloroform fraction from Pyropolyporus fomentarius in K562 cells

Abstract

Keywords

Introduction

Materials and methods

Plant materials

Preparation of the CHCl3 extract

Component analysis by GC/MS

Cell culture

Isolation of PBMC

Assessment of cell viability after PFCF treatment

Membrane integrity testing

The measurement of cell membrane potential

SEM observation

Annexin V-PE/7-AAD staining experiment

Measurement of [Ca2+]i

Detection of intracellular ROS generation

Examination of the Δψ m

Statistical analysis

Results

Chemical compounds in PFCF

Effect of PFCF on cell viability

Plasma membrane damage measurement

Apoptosis assessment

Changes in intracellular [Ca2+]i

Detection of intracellular ROS generation and Δψ m loss

Discussion

Footnotes

Conflict of interest

Funding

References

Preparation of the CHCl₃ extract

Measurement of [Ca²⁺]_i

Examination of the Δψ _m

Changes in intracellular [Ca²⁺]_i

Detection of intracellular ROS generation and Δψ _m loss