SP600125

Coriolus versicolor‐derived protein‐bound polysaccharides trigger the caspase‐independent cell death pathway in amelanotic but not melanotic melanoma cells

Małgorzata Pawlikowska1 | Jakub Piotrowski1 | Tomasz Jędrzejewski1 |Wiesław Kozak1 | Andrzej T. Slominski2,3 | Anna A. Brożyna4

Abstract

We have investigated the potential cell death mechanism promoted by Coriolus versicolor fungus‐derived protein‐bound polysaccharides (PBPs) in melanoma cells. Knowing that melanogenesis has the potential to affect the tumor behavior and melanoma therapy outcome, the cytotoxic effects of PBPs were evaluated in human SKMel‐188 melanoma cell line, whose phenotype, amelanotic versus pigmented, depends on the concentration of melanin precursors in the culture medium. Our results showed that inhibitory effect of PBPs (100 and 200 μg/ml) towards melanoma cells is inversely associated with the pigmentation level. This cytotoxicity induced in nonpigmented melanoma cells by PBPs was caspase‐independent; however, it was accompanied by an increased intracellular reactive oxygen species (ROS) generation. The ROS production was controlled by c‐Jun N‐terminal kinase (JNK) because SP600125, a JNK inhibitor, significantly reduced ROS generation and protected cells against PBPs‐induced death. We also found that PBPs‐induced lactate dehydrogenase release in amelanotic melanoma cells was abolished by co‐treatment with receptor‐interacting serine/threonine‐protein kinase 1 inhibitor, implying engagement of this kinase in PBPs‐induced death pathway. The results suggest that PBPs induce an alternative programmed cell death, regulated by receptor‐interacting protein‐1 and ROS and that this process is modified by melanin content in melanoma cells. These findings are remarkable when considering the use of commercially available Coriolus versicolor by patients who suffer from melanoma cancer.

KEYWORDS
caspase‐independent cell death, melanin, melanoma cells, necroptosis, protein‐bound polysaccharides, ROS

1 | INTRODUCTION

Schadendorf, 2001; Miller et al., 2016). Despite improved early diagMelanoma, one of the most aggressive forms of skin cancer with high nosis and impressive advancement in new therapies, (Lo & Fisher, metastatic potential, is a tumor with the most rapidly increasing 2014; Rajkumar & Watson, 2016; Shah & Dronca, 2014) the high mortality rate among melanoma patients represents a serious clinical problem once the metastatic process started (Nikolaou & Stratigos, 2014; Rajkumar & Watson, 2016). Thus, search for new alternative methods of melanoma treatments is needed.
A number of bioactive molecules, including anticancer agents, have been identified in various mushroom species (Jiménez‐Medina et al., 2008). Among them, the protein‐bound polysaccharides (PBPs) are one of the best known and most potent bioactive molecules that exert strong antitumor, as well as immunomodulating actions. PBPs, obtained from Coriolus versicolor (CV), the Chinese fungus with medicinal use in both, traditional herbalism as well as modern clinical practice, were initially believed to promote health, strength and longevity. Currently, products using CV extracts are approved as adjunct therapy in China and Japan for cancer patients already receiving chemotherapy or radiotherapy (Saleh, Rashedi, & Keating, 2017; Zong, Cao, & Wang, 2012). Despite numerous in vitro and in vivo studies and some clinical trials, which confirmed the direct inhibitory effects of PBPs on cancer cells, the exact mechanism underlying these effects is still poorly understood.
Previously, we have shown that CV‐derived polysaccharide peptides, induce differential effects depending on target cells (Kowalczewska, Piotrowski, Jędrzejewski, & Kozak, 2016). Acting as biological response modifiers, PBPs exert strong immune‐enhancing effect on blood lymphocytes and show strong tumor inhibitory properties. We have demonstrated that PBPs induced cytotoxicity in breast cancer cells (MCF‐7) is TNF‐α dependent (Kowalczewska et al., 2016).
In this study, we have investigated the PBPs effects towards human melanoma cells and the possible mechanism of the induced cell death. Because melanogenesis, a process started by enzymatic oxidation of L‐tyrosine and L‐DOPA, has the potential to affect the tumor behavior and melanoma therapy outcome (Brozyna, Jozwicki, Carlson, & Slominski, 2013; Brozyna, Jozwicki, Roszkowski, Filipiak, & Slominski, 2016; Li, Slominski, & Slominski, 2009; Slominski, Paus, & Mihm, 1998; Sniegocka et al., 2018), we have performed our investigation on experimental model, human SKMel‐188 melanoma cell line, whose phenotype, amelanotic versus melanotic can be regulated by the concentration of melanin precursors in culture medium (Slominski, Ermak, & Wortsman, 1999; Slominski, Zbytek, & Slominski, 2009). The SKMel‐188 cell line represents a unique research model that allows to obtain the results independent of individual cell line properties resulting from differences at the DNA level. Thus, this cell line enables conducting comparative studies on the biology of both forms of melanoma. In the current study, we have shown that cytotoxicity induced by PBPs towards melanoma cells is inversely proportional to the activity of melanogenic pathway. Further molecular analysis revealed that PBPs extract induce a caspase‐independent programmed cell death, regulated by receptor‐interacting serine/threonine‐protein kinase 1 (RIPK1) and reactive oxygen species (ROS). To the best of our knowledge, this is the first study reporting that CV PBPs cause caspase‐independent cell death with characteristic features of necroptosis in amelanotic, but not melanotic melanoma cells.

2 | MATERIAL AND METHODS

2.1 | Cell culture

Human SKMel‐188 melanoma cell line was a gift of Dr Chakraborty, Yale University. The cells were cultured in either Ham’s F10 or Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 5% fetal bovine serum and 1% antibiotics (penicillin/streptomycin/ amphotericin) as described previously (Slominski et al., 1999; Slominski et al., 2009). All components were provided from Sigma Aldrich (Darmstadt, Germany). The melanogenic phenotype of this cell line is dependent on the concentration of L‐tyrosine in the medium, with amelanotic phenotype in Ham’s F10 medium (10μM L‐tyrosine) and melanotic one in DMEM medium (400μM L‐tyrosine). The cells were cultured at 37°C in humidified atmosphere of 5% CO2, and the media were changed every second day following protocols described previously (Slominski et al., 1999).

2.2 | Preparation of CV PBPs extract

PBPs were extracted from commercially available Chinese fungus CV capsules (MycoMedica, Czech Republic) following protocols described previously (Kowalczewska et al., 2016; Pawlikowska, Jędrzejewski, Piotrowski, & Kozak, 2016). Briefly, phosphate‐buffered saline (PBS 1×, pH 7.4; Sigma Aldrich, Darmstadt, Germany) was used to dissolve CV powder in order to obtain the stock solution of 8 mg/ml and 4 mg/ml. After continuous agitation for 48 hr at room temperature, insoluble material was removed by centrifugation at 2,000 x g for 10 min. The soluble PBPs, which were found to be the main active compound of CV extract and the most prominent in antitumor and immunomodulatory efficacy (Ho et al., 2004), constitute 25% of the CV capsule as confirmed by certificate of analysis from manufacture. The soluble supernatant, containing 2 mg/ml or 1 mg/ml of PBPs, was than sterilized using 0.22‐μm filters and diluted with appropriate cell culture media to give the final concentrations indicated in experiments. The doses of PBPs used in the current study were established previously (Kowalczewska et al., 2016; Pawlikowska et al., 2016).

2.3 | MTT test

3‐(4,5‐Dimethyl‐2‐thiazolyl)‐2,5‐diphenyl‐2H‐tetrazolium bromide(MTT) tests were performed to evaluate melanoma cells viability after PBPs treatment. This colorimetric assay detects the reduction of MTT (Sigma Aldrich, Darmstadt, Germany) by mitochondrial dehydrogenase to blue formazan product. Melanoma cells (nonpigmented and pigmented) were seeded at density of 5,000 cells per well into 96‐well plates in appropriate medium. Following addition of PBPs extract (100 μg/ml or 200 μg/ml) for 2, 4, 6, 8, 12, and 24 hr, 10 μl of filtered (0.22 μm) 5 mg/ml MTT dissolved in PBS was added to each well, and the plates were incubated in a dark, at 37°C, humidified atmosphere containing 5% CO2 for 4 hr. After the incubation period, the supernatants were removed, and 100 μl of dimethyl sulfoxide was added. The plates were mixed horizontally for 15 min, and the optical density was measured at 570 nm (with reference wavelength of 630 nm) using Synergy HT Multi‐Mode Microplate Reader (BioTek; Winooski, VT, USA). The results were expressed as percentage of control cells; 570/630 nm ratio in control wells (incubated with PBS, a solvent of PBP) serves as 100%.

2.4 | Lactate dehydrogenase leakage assay

The leakage of lactate dehydrogenase (LDH) in melanoma cells was determined using a CytoTox 96 Non‐Radioactive Cytotoxicity Assay (Promega, Madison, WI, USA). Initially, cells were seeded at a density of 10,000 cells per well in a 96‐well plate at 37°C and 5% CO2 atmosphere in 100 μl media per well (Ham’s F10 or DMEM). After 24 hr, the medium was replaced with fresh medium containing PBPs extract (100 or 200 μg/ml). In control wells, PBS (solvent) was added with the corresponding media. Following incubation for 2, 4, 6, 8, 12, or 24 hr, 50 μl of aliquots of supernatants were transferred to new wells of 96well plate and mixed with equal amounts of freshly prepared assay CytoTox 96 Reagent supported by the assay kit. The microtiter plate was incubated for 30 min at room temperature in the dark as described by manufacturer. The absorbance was measured at 490 nm using the Synergy HT Multi‐Mode Microplate Reader. The PBP mediated cytotoxicity was calculated according to the formula:%cytotoxicity ¼ experimental LDH release ðOD490Þ=maximumLDH release ðOD490Þ;where the maximal release was obtained after lysis of the cells with a control solution provided by the manufacturer. Culture medium background was subtracted from all values.

2.5 | Bcl‐2 expression and caspase 3/7 activity assay

The Bcl‐2 protein expression was analyzed in amelanotic and melanotic melanoma cells after 24 hr PBPs stimulation using the human Bcl‐2 ELISA Kit (Abcam, UK). The results are expressed in nanogram per milligram of total protein (assayed by bicinchoninic acid method). Luminescent caspase‐Glo 3/7 assay was carried out on PBPs treated amelanotic and melanotic SKMel‐188 cells according to the manufacturer’s instructions (Promega, Madison, WI, USA). The cells were seeded in white 96‐well plates and treated with the PBPs extract (100 and 200 μg/ml) or 1μM staurosporine as a positive control for 2, 4, 6, 8, 12, and 24 hr. The cells were than incubated with caspase‐Glo 3/7 reagent for 1 hr at room temperature, which was followed by measuring the enzymatic activity of caspase through luminescent signal using Synergy HT Multi‐Mode Microplate Reader. The results are expressed in relative luminesce units.

2.6 | Cellular ROS determination by DCF‐DA method

Intracellular formation of reactive oxygen species was assessed using oxidation sensitive dye 2’,7’‐dichlorofluorescin diacetate (DCF‐DA; Sigma Aldrich, Darmstadt, Germany) as a substrate. Melanotic or amelanotic SKMel‐188 cells were seeded into 96‐well microtiter plates at 25,000 per well and preincubated at 37°C for 24 hr. Then, the cells were washed with PBS, loaded with 20μM DCF‐DA in Hank’s balanced salt solution and incubated at 37°C for 45 min in the dark. Following incubation, the DCF‐DA solution was removed, and cells were washed with PBS. Subsequently, the proper medium (Ham’s F10 or DMEM) was added to each well, and the cells were incubated with PBPs extract (100 and 200 μg/ml) for 2, 4, 6, 8, 12, and 24 hr. The fluorescence was measured in a Synergy HT Multi‐Mode Microplate Reader using excitation at 485 nm and emission at 528 nm.The results are expressed in relative fluorescence units.

2.7 | Monitoring amelanotic melanoma cell death induced by polysaccharidopeptides

The cell death of amelanotic melanoma cells induced by PBPs was monitored using Tomocube HT‐1S microscope (Inc., Daejeon, Korea). Briefly, 300,000 of the amelanotic SKMel‐188 cells were seeded on TomoDishes, special dishes for live cell imaging designed for reproducible results for live cell imaging. After 24 hr of preincubation, the medium was changed, and PBPs extract was added (100 μg/ml or 200 μg/ml). The control cells were treated with PBS. The twodimensional and three‐dimensional visualization of the cellular images, based on 3D RI distributions of the cells, were rendered with Tomostudio, Software (Korea) after 24 hr of treatment.

2.8 | c‐Jun N‐terminal kinase inhibition

To test whether c‐Jun N‐terminal kinase (JNK) activity controls the cytotoxicity and ROS generation in melanoma cells after PBPs treatment, we have performed additional MTT and DCF‐DA assays using JNK inhibitor (SP600125, Sigma Aldrich, Darmstadt, Germany). Briefly, after preparation of stock solution of SP600125 (10 mM, using 100% dimethyl sulfoxide), the amelanotic melanoma cells were pretreated with JNK inhibitor at the final concentration of 20 μM for 30 min. The cells were washed with PBS and treated with PBPs extract (100 or 200 μg/ml) for 12 and 24 hr, and the viability of cells and ROS generation were assessed according to the procedures described above. The results are expressed as percent of control, represented by the cells incubated with PBS (the control for PBPs only stimulated cells) or the cells incubated with PBS pretreated with SP600125 (the control for PBPs stimulated cells pretreated with JNK inhibitor).

2.9 | Inhibition of RIP1 kinase

To characterize the molecular mechanism of PBP‐induced cytotoxicity of human amelanotic melanoma cells, the necrostatin‐1 (Nec‐1; Sigma Aldrich, Darmstadt, Germany), specific receptor‐interacting protein‐1 (RIP1) kinase inhibitor was used. The amelanotic SKMel‐188 cells maintained in 96‐well plates were treated with PBPs extract (100 nd 200 μg/ml) alone or in combination with RIPK1 inhibitor, Nec‐1 (30 μM). After 2, 4, 6, 8, 12, and 24 hr of treatment, the LDH release was evaluated according to the procedure described above. The results were expressed as percent of the cytotoxicity compared with controls.

3 | STATISTICAL ANALYSIS

All values are reported as means ± standard error of the means (SEM) and were analyzed using analysis of variance followed by Bonferroni multiple comparisons test with the level of significance set at p < .05. Statistical analyses were performed with GraphPad Prism 7.0 (La Jolla, CA, USA). 4 | RESULTS 4.1 | PBPs‐induced decrease in viability of melanoma cells depends on the pigmentary phenotype Using an unique in vitro human melanoma SKMel‐188 cell line, in which the level of melanin pigmentation can be controlled by the concentration of melanin precursors in the culture medium (Slominski et al., 1999; Slominski et al., 2009), we have demonstrated that the cells sensitivity to PBPs extract is inversely related to melanin presence (Figure 1a,b). The PBPs decreased the amount of metabolically active nonpigmented (cultured in Ham's F10) human melanoma cells. The effect was time‐dependent with a gradual decrease of viability observed at 2–8 hr of treatment. After 12 hr, the viability of cells increased; however, it was still significantly lower (p < .001) as compared with control cells incubated with PBS (75.8% ± 2.49% or 68.4% ± 2.87% of control cells for 100 or 200 μg/ml of PBPs, respectively). Twenty‐four hours of treatment resulted in further decline of cells viability (65.4% ± 1.74% and 62.6% ± 2.94% of control cells for 100 and 200 μg/ml of PBPs, respectively). The effect of PBPs on a viability of pigmented melanoma cells was different from that mediated towards amelanotic ones. As shown in Figure 1b, after initial decrease (2 or 4 hr) in number of viable cells, there was an increase in cell viability up to control values seen at 12 and 24 hr, for PBPs at 100 and 200 μg/ml, respectively. Ham's F10—amelanotic phenotype or Dulbecco's Modified Eagle's Medium—melanotic phenotype and treated with different concentrations of PBPs (100 and 200 μg/ml) for 2, 4, 6, 8, 12, and 24 hr. Cells viability (a and b) is expressed as the mean percent 3‐(4,5‐dimethylthiazol‐2‐yl)‐2,5diphenyltetrazolium bromide absorbance (ratio of absorbance in wells with PBPs‐treated cells to that with untreated counterparts, control cells×100%) ±SEM of three independent experiments with six wells for each experiment. Cytotoxicity was assessed by LDH release in (c) nonpigmented and (d) melanotic SKMel‐188 cells. The released LDH was measured using Promega's CytoTox 96 Non‐Radioactive Cytotoxicity Assay. Total cytotoxicity was calculated by comparing the levels of released LDH in the experimental samples with total levels of cellular LDH obtained by lysing 1 × 104 corresponding cells with Promega cell lysis buffer. Asterisk indicates significant differences compared with the control cells (*p < .05, **p < .01, ***p < .001). LDH, lactate dehydrogenase; PBPs, protein‐bound polysaccharides In order to confirm that the differences observed between the effects of PBPs on SKMel‐188 cells are due to the presence of melanin, we performed the additional experiment with melanogenesis inhibitor. The melanotic melanoma cells were treated with a tyrosinase inhibitor, kojic acid (5‐hydroxy‐2‐hydroxymethyl‐1,4‐pyrone). After 3 days of culturing cells in DMEM medium with melanogenesis inhibitor, the cells were treated with PBPs at a concentration of 200 μg/ml. The results revealed that kojic acid at concentration of 6 μg/ml is potent to inhibit melanin synthesis and sensitized melanoma cells to PBPs (Data S1). These results confirmed our findings that inhibitory effects of PBPs towards melanoma cells depends on their pigmentation status. 4.2 | Cytotoxic effect of PBPs is inversely correlated to melanin content To test the toxicity of PBPs, the release of cytosolic LDH into the culture medium by melanoma cells was measured. Figure 1c shows that the PBPs‐stimulated nonpigmented SKMel‐188 cell response was significantly higher in comparison with control cells. The LDH release was the highest after 8, 12, and 24 hr of treatment. The significant differences in LDH leakage in response to 100 or 200 μg/ml of PBPs were observed after 8 and 24 hr of treatment. These data are in agreement with MTT test (presented in Figure 1a). The high melanin content attenuated the cytotoxic effect of PBPs on melanoma cells (Figure 1d). Thus, the LDH release in pigmented melanoma cells did not change after treatment with PBPs as compared with control. We also observed that spontaneous LDH leakage from untreated (control) pigmented cells was higher than that from nonpigmented controls (Figure 1c,d). Moreover, total LDH release after 12 hr of incubation was significantly higher in wells with cells cultured in DMEM medium only as compared with those stimulated with PBPs(Figure 1d). 4.3 | PBPs neither alter Bcl2 expression nor activate caspase 3/7 in melanoma cells In many systems, members of the Bcl‐2 family modulate apoptosis. The Bcl‐2 family of proteins controls a critical step in commitment to apoptosis by regulating the permeabilization of the mitochondrial outer membrane (MOMP). Anti‐apoptotic Bcl‐2 proteins reside on the mitochondrial outer membrane (MOM) and prevent apoptosis by inhibiting the activation of the pro‐apoptotic family members Bax and Bak (Lindsay, Esposti, & Gilmore, 2011). Bcl‐2, encoding the 26kD protein that blocks programmed cell death, is frequently overexpressed in many types of tumors (Chan & Yu, 2004). Therefore, in our study, we have evaluated the effect of PBPs extract on Bcl‐2 expression. Regarding the protein level, our results demonstrated that there was no alteration in the relative Bcl‐2 expression in PBPsstimulated cells compared with control cells. After 24 hr treatment, we did not detect any differences, neither in amelanotic nor melanotic melanoma cells (Figure 2a,b). Although there is a feedback loop between bcl‐2 and caspases (especially caspase‐3), bcl‐2 cannot always inhibit apoptosis. Recent evidence suggests that caspase family of enzymes may be able to inactivate the bcl‐2 anti‐apoptotic function and further enhance cell death, even when apoptosis is triggered via a non‐Bax/Bcl‐2 dependent pathway (Yang et al., 2002). Thus, in our study, we performed an analysis of caspase 3/7 activity in PBPs treated melanoma cells for 2, 4, 6, 8, 12, and 24 hr. No significant increases in caspase 3/7 activity were observed after treatment with PBPs of amelanotic melanoma cells (Figure 2c). The additional assessment of caspase‐8 expression also did not reveal changes between treated and control cells (data not shown). Melanotic melanoma cells exhibited higher endogenous activity of caspase 3/7 in comparison with the amelanotic cells. Especially after 24 hr of incubation, the level of this activity in nontreated melanotic cells was comparable with that induced by 1μM staurosporine (positive control; Figure 2d). The examined PBPs extract, however, had no effect on the caspase activity in melanotic cells in comparison with controls. The significant activity of caspase 3/7 in melanotic melanoma cells results from the inherent cytotoxicity of melanin precursors. The intermediates of melanogenic process affect strongly the cells growth; however, the cell culture is still viable (Data S2). 4.4 | ROS are stimulated in amelanotic melanoma cells by PBPs treatment Once we determined that cell death pathway of nonpigmented SKMel‐188 melanoma might be Bcl‐2 and caspase‐independent, we investigated whether PBPs would affect intracellular ROS generation. The generation of cellular ROS in melanoma cells was observed using a DCF‐DA assay. Figure 2e shows the induced ROS generation in the amelanotic melanoma cells. The density of DCF‐positive cells significantly increased in a time‐dependent manner, implying that nonpigmented cells cultured with PBPs generated a higher amount of intracellular ROS as compared with the control. After 4, 8, and 24 hr of stimulation, 200 μg/ml of PBPs increased the cytosolic ROS generation by 2.1, 3.5 and 6.5 fold, respectively, compared with control cells (PBPs in concentration 100 μg/ml: 1.3, 2.4, and 6.4 fold after 4, 8, and 24 hr of treatment, respectively). We did not, however, observed any significant alteration in cellular ROS generation in PBPs‐stimulated melanotic cells when compared with control (Figure 2f). This result is in agreement with the cytotoxicity assays shown in Figure 1d. 4.5 | PBPs induce morphological changes in amelanotic melanoma cells In order to closely examine the features of amelanotic melanoma cells death upon PBPs treatment, we performed analysis of PBP effects at an individual cell level. By monitoring the cell death using Tomocube HT‐1S microscope, we captured the changes in amelanotic SKMel188 PBPs‐treated cells (Figure 3b,c,e,f) morphology such as plasma membrane rupture and perforation as well as membrane protrusion compared with control, nontreated cells (Figure 3a,d). 4.6 | JNK inhibitor abolishes the PBPs‐induced ROS generation and restores the viability of nonpigmented melanoma cells Because we have shown that amelanotic melanoma cell death induced by PBPs is caspase‐independent, but can be dependent on ROS production, we have examined the potential role of JNK in this process. JNK plays an important role in programmed necrosis, it is activated by ROS, and also controls ROS generation (Fortes et al., 2012). As shown in Figure 4a,b, treatment with a selective JNK inhibitor abrogated the cell death induced by PBPs after 12 and 24 hr. Moreover, inhibition of JNK caused a significant reduction in intracellular ROS generation induced by the PBPs extract (Figure 4c,d). Our experiments demonstrated that JNK kinase regulates mitochondrial ROS production under the conditions studies here. By blocking the phosphorylation of JNK by its inhibitor, SP600125, we have shown that JNK is required for ROS production and cell death of amelanotic melanoma cells triggered by PBPs extract. These results suggest that multiple mechanisms including JNK activation and ROS generation contribute to PBPs‐induced cytotoxicity. 4.7 | PBPs‐induced cytotoxicity depends on RIP1 kinase Our results indicate that PBPs‐induced cell death of amelanotic melanoma cells is mediated through caspase‐independent mechanism in which intracellular ROS are involved. To further characterize the cell death pathway in nonpigmented melanoma cells after PBPs treatment, we examined the role of RIP1 kinase. We tested the effect of Nec‐1 (30 μM), a selective inhibitor of RIP1 kinase, on PBPs‐induced melanoma cell death. The treatment with Nec‐1 in the presence of PBPs, at both concentrations, 100 μg/ml as well as 200 μg/ml, abolished the PBPs‐induced cytotoxicity. The LDH release was significantly reduced after co‐treatment of cells with Nec‐1 in comparison with the PBPs only—treated cells (Figure 4e). In order to check whether the cytotoxic activity of PBPs is not only SKMel‐188 cell type specific, we performed similar analysis on other human amelanotic melanoma cell line, C32. The results of evaluated cells viability after PBPs stimulation alone, or in combination with Nec‐1, confirmed the previous findings showing that amelanotic melanoma cells are sensitive to cytotoxic activity of PBP and that the cell death is mediated by pathway involving RIP1 (Data S3). 5 | DISCUSSION Plant‐ and fungus‐derived compounds have become of significant interest in medicine as related to development of new anticancer treatment strategies (Pfisterer, Wolber, Efferth, Rollinger, & Stuppner, 2010). In this study, we have demonstrated that PBPs isolated from commercially available CV Chinese fungus capsules exert cytotoxic effect towards melanoma cells. Interestingly, this effect was inversely associated with the degree of cell pigmentation. The synthesis of melanin is a biochemical pathway characteristic for melanocytes, in which L‐tyrosine is transformed to heterogeneous melanin polymer through series of oxidation‐reduction reactions, which also affects cellular metabolism (Li et al., 2009). The synthesis of melanin not only serves as a diagnostic tool, but it also affects the behavior of normal as well as malignant melanocytes (Brozyna, VanMiddlesworth, & Slominski, 2008; Brozyna et al., 2016; Slominski et al., 1998). The animal‐based and experimental cell culture models have shown that melanogenesis has the potential to affect tumor behavior and the outcome of melanoma therapy (Brozyna et al., 2008; Cichorek, 2011; Cichorek, Kozłowska, & Bryl, 2007; Slominski et al., 1998; Slominski et al., 2009; Sniegocka et al., 2018). Previous reports have shown that melanotic and amelanotic melanomas differ in their sensitivity to ionizing and ultraviolet radiation (Brozyna et al., 2008; Chalmers, Lavin, Atisoontornkul, Mansbridge, & Kidson, 1976; Hopwood, Swartz, & Pajak, 1985; Sniegocka et al., 2018). The results of in vitro study conducted on two selected human melanoma cell lines with different melanin content have shown that the amount of intracellular melanin is inversely related to the radiosensitivity of melanoma cells (Kinnaert et al., 2000). In vitro studies on Bomirski melanoma cell lines have also revealed that camptothecin, a plant alkaloid produced by Camptotheca acuminata, caused death of amelanotic melanoma cells through the apoptotic way, whereas melanotic melanoma cells were insensitive to the activity of this agent (Cichorek et al., 2007). Furthermore, melanin has attenuated cytotoxicity of cyclophosphamide and lymphotoxicity against melanoma cells (Slominski et al., 2009). Our results showing that fungus PBPs act cytotoxic only on amelanotic melanoma cells but not pigmented ones are in agreement with the above mentioned studies. The observed decrease in the viability of nonpigmented cells, assayed by MTT test, was accompanied by a release of the cytoplasmatic LDH, which is the common characteristic feature of cell‐death processes, necrosis, and apoptosis. In parallel with the above mentioned data, the melanotic melanoma cells differed in their sensitivity to PBPs extract. Although the amount of metabolically active cells diminished during the first 8 hr of treatment, there were no cell membrane damages as demonstrated by lack of changes in LDH release. These results suggest that the growth inhibitory effects of PBPs observed during the first hours of treatment could be mediated rather by a cell cycle arrest than cell‐induced death. Longer treatment with PBPs extract (12 and 24 hr) did not affect the cells viability compared with control, supporting the cell cycle arrest hypothesis. The cytotoxic effect of PBPs towards amelanotic but not melanotic melanoma cells, observed in the current study, confirms the hypothesis that melanotic melanomas are indeed more resistant to therapy than amelanotic ones (Slominski et al., 1998). Because most of currently used anticancer agents are effective via induction of the apoptotic machinery, we investigated the potential cell death mechanism promoted by PBPs extract. Our results have shown that PBPs‐induced amelanotic melanoma cell death (confirmed by holotomographic assessment) is Bcl‐2‐ and caspase‐independent. The assessed expression of initiator caspase‐8, responsible among others (e.g., caspase‐2, ‐9, and ‐10) for initiating a downstream apoptotic cascade, was undetected in melanoma cells treated with PBPs. Furthermore, the activity of the effector caspases 3 and 7, accountable for the actual dismantling of the cells by cleaving cellular substrates, were also unchanged, providing an evidence for caspaseindependent mechanism of PBPs‐induced cell death. We also assessed caspase 3/7 in pigmented melanoma cells. Although PBPs extract did not affect their activity, we have noticed that these caspases in nontreated, control melanotic melanoma cells, were upregulated as compared with amelanotic ones. During melanization, oxidation products of tyrosine are generated, which are toxic to the cells (Pawelek, 1976). The observed cytotoxicity of the melanogenic intermediates DOPA (l‐3,4‐dihydroxyphenylalanine), DHI (5,6‐dihydroxyindole), and DHICA (DHI‐2‐carboxylic acid) increase with the time of incubation (Urabe et al., 1994); therefore after 24 hr of cells culturing, we detected the elevated activity of caspase 3/7 in melanotic melanoma cells. These results are consistent with other reports showing the cytotoxic effects of intermediates of melanogenesis (Brenner & Hearing, 2008). Although caspase‐dependent apoptosis is often considered as a molecular mechanism by which cancer therapies exert their antitumor effects, there is an increasing evidence of agents inducing cancer cell death that is independent of caspase activation (Kong, Lv, Yan, Chang, & Wang, 2018; van der Walt, Zakeri, & Cronjé, 2016). More recently, it was observed that some anticancer agents can induce other forms of cell death, such as programmed necrosis (Cho & Park, 2014; Mattia et al., 2018). Such caspase‐independent cell death pathways are important defense mechanisms particularly in case of cancers, where apoptotic pathway is evaded (Bröker, Kruyt, & Giaccone, 2005). In the present work, we have shown that protein‐bound polysaccharides induced cytotoxicity in nonpigmented melanoma cells is caspaseindependent, and it is mediated by generation of ROS. Reactive oxygen species are thought to act as oxidizing in addition to other cellular proteins, mitogen‐activated protein kinase phosphatases, whose normal function is to downregulate JNK signaling pathway. This mitogen‐activated protein‐kinase‐oxidizing activity of ROS results in prolonged JNK activation leading to cell death (Christofferson & Yuan, 2010). In the current study, we have shown that ROS generation in PBPs‐treated cells was controlled by JNK kinase activity because inhibition of JNK with SP600125 protected amelanotic melanoma cells against PBPs‐induced cell death and significantly attenuated the intracellular ROS generation. Because ROS production appears to be a main executioner of necroptosis (Christofferson & Yuan, 2010; Song et al., 2011), we have also evaluated the activity of upstream component of this caspase‐independent necrotic cell death, a RIP1. Its serine/threonine kinase activity, which is essential for this necrotic death pathway, can be inhibited by Nec1, a small molecule acting as inhibitor of necroptosis. Correspondingly, we found that PBPs‐induced LDH release in nonpigmented melanoma cells was abolished by treating cells with Nec‐1. Because preventing necroptosis through RIPK1 inhibition serves as a useful means of differentiating between necroptosis and fortuitous forms of necrosis (Kroemer et al., 2009), we concluded that PBPs increase cellular ROS and induce cell necroptosis, but not apoptosis, because Bcl‐2 and caspases expression have been absent (Cho & Park, 2014; Holler et al., 2000). 6 | CONCLUSION In conclusion, the current study for the first time demonstrates that PBPs from commercially available CV capsules cause caspaseindependent cell death with characteristics of necroptosis in amelanotic but not melanotic melanoma cells. The cytotoxic effects of PBPs seem to require RIPK1, ROS, and JNK activity. Because melanoma cells are expected to be sensitive to therapeutic strategies inducing cancer cell death through ROS production, the use of PBPs, the natural compounds that interfere in redox regulation, might be a promising adjunctive therapy to selectively target amelanotic melanoma tumors. However, further studies elucidating the molecular death pathway are needed. These findings, after confirmation in in vivo studies, may introduce a new mechanistic perspective in the melanoma therapy and could provide new targets for therapeutic development. REFERENCES Brenner, M., & Hearing, V. J. (2008). The protective role of melanin against UV damage in human skin. Photochem Photobiol, 84(3), 539–549. https://doi.org/10.1111/j.1751‐1097.2007.00226.x Bröker, L. E., Kruyt, F. A., & Giaccone, G. (2005). Cell death independent of caspases: A review. Clin Cancer Res, 11(9), 3155–3162. https://doi.org/ 10.1158/1078‐0432.CCR‐04‐2223 Brozyna, A. A., Jozwicki, W., Carlson, J. A., & Slominski, A. T. (2013). Melanogenesis affects overall and disease‐free survival in patients with stage III and IV melanoma. Hum Pathol, 44(10), 2071–2074. https:// doi.org/10.1016/j.humpath.2013.02.022 Brozyna, A. A., Jozwicki, W., Roszkowski, K., Filipiak, J., & Slominski, A. T. (2016). Melanin content in melanoma metastases affects the outcome of radiotherapy. Oncotarget, 7(14), 17844–17853. Brozyna, A. A., VanMiddlesworth, L., & Slominski, A. T. (2008). Inhibition of melanogenesis as a radiation sensitizer for melanoma therapy. Int J Cancer, 123(6), 1448–1456. https://doi.org/10.1002/ijc.23664 Chalmers, A. H., Lavin, M., Atisoontornkul, S., Mansbridge, J., & Kidson, C. (1976). Resistance of human melanoma cells to ultraviolet radiation.Cancer Res, 36(6), 1930–1934. Chan, S. L., & Yu, V. C. (2004). Proteins of the bcl‐2 family in apoptosis signalling: From mechanistic insights to therapeutic opportunities. Clin Exp Pharmacol Physiol, 31, 119–128. https://doi.org/10.1111/j.14401681.2004.03975.x Cho, Y. S., & Park, S. Y. (2014). Harnessing of programmed necrosis for fighting against cancers. Biomol Ther (Seoul), 22(3), 167–175. https:// doi.org/10.4062/biomolther.2014.046 Christofferson, D. E., & Yuan, J. (2010). Necroptosis as an alternative form of programmed cell death. Curr Opin Cell Biol, 22(2), 263–268. https:// doi.org/10.1016/j.ceb.2009.12.003 Cichorek, M. (2011). Camptothecin‐induced death of amelanotic and melanotic melanoma cells in different phases of cell cycle. Neoplasma, 58(3), 227–234. https://doi.org/10.4149/neo_2011_03_227 Cichorek, M., Kozłowska, K., & Bryl, E. (2007). The activity of caspases in spontaneous and camptothecin‐induced death of melanotic and amelanotic melanoma cell. Cancer Biol Ther, 6(3), 346–353. https:// doi.org/10.4161/cbt.6.3.3701 Fortes, G. B., Alves, L. S., de Oliveira, R., Dutra, F. F., Rodrigues, D., Fernandez, P. L., … Bozza, M. T. (2012). Heme induces programmed necrosis on macrophages through autocrine TNF and ROS production. Blood, 119(10), 2368–2375. https://doi.org/10.1182/blood‐2011‐08375303 Helmbach, H., Rossmann, E., Kern, M. A., & Schadendorf, D. (2001). Drugresistance in human melanoma. Int J Cancer, 93(5), 617–622. https:// doi.org/10.1002/ijc.1378 Ho, C. Y., Lau, C. B., Kim, C. F., Leung, K. N., Fung, K. P., Tse, T. F., … Chow, M. S. S. (2004). Differential effect of Coriolus versicolor (Yunzhi) extract on cytokine production by murine lymphocytes in SP600125 vitro. Int Immunopharmacol, 4(12), 1549–1557. https://doi.org/10.1016/j. intimp.2004.07.021
Holler, N., Zaru, R., Micheau, O., Thome, M., Attinger, A., Valitutti, S., … Tschopp, J. (2000). Fas triggers an alternative, caspase‐8‐independent cell death pathway using the kinase RIP as effector molecule. Nat Immunol, 1(6), 489–495. https://doi.org/10.1038/82732
Hopwood, L. E., Swartz, H. M., & Pajak, S. (1985). Effect of melanin on radiation response of CHO cells. Int J Radiat Biol Relat Stud Phys Chem Med, 47(5), 531–537. https://doi.org/10.1080/09553008514550761
Jiménez‐Medina, E., Berruguilla, E., Romero, I., Algarra, I., Collado, A., Garrido, F., & Garcia‐Lora, A. (2008). The immunomodulator PSK induces in vitro cytotoxic activity in tumour cell lines via arrest of cell cycle and induction of apoptosis. BMC Cancer, 8, 78. 1‐10
Kinnaert, E., Morandini, R., Simon, S., Hill, H. Z., Ghanem, G., & Van Houtte, P. (2000). The degree of pigmentation modulates the radiosensitivity of human melanoma cells. Radiat Res, 154, 497–502. https://doi.org/ 10.1667/0033‐7587(2000)154[0497:TDOPMT]2.0.CO;2
Kong, Q., Lv, J., Yan, S., Chang, K. J., & Wang, G. (2018). A novel Naphthyridine derivative, 3u, induces necroptosis at low concentrations and apoptosis at high concentrations in human melanoma A375 cells. Int J Mol Sci, 19(10). https://doi.org/10.3390/ijms19102975 Kowalczewska, M., Piotrowski, J., Jędrzejewski, T., & Kozak, W. (2016). Polysaccharide peptides from Coriolus versicolor exert differential immunomodulatory effects on blood lymphocytes and breast cancer cell line MCF‐7 in vitro. Immunol Lett, 174, 37–44. https://doi.org/ 10.1016/j.imlet.2016.04.010
Kroemer, G., Galluzzi, L., Vandenabeele, P., Abrams, J., Alnemri, E. S., Baehrecke, E. H., … Melino, G. (2009). Classification of cell death: Recommendations of the Nomenclature Committee on Cell Death 2009. Cell Death Differ, 16(1), 3–11. https://doi.org/10.1038/cdd.2008.150
Li, W., Slominski, R., & Slominski, A. T. (2009). High‐resolution magic angle spinning nuclear magnetic resonance analysis of metabolic changes in melanoma cells after induction of melanogenesis. Anal Biochem, 386(2), 282–284. https://doi.org/10.1016/j.ab.2008.12.017
Lindsay, J., Esposti, M. D., & Gilmore, A. P. (2011). Bcl‐2 proteins and mitochondria—Specificity in membrane targeting for death. Biochim Biophys Acta, 1813(4), 532–539. https://doi.org/10.1016/j.bbamcr.2010.10.017
Lo, J. A., & Fisher, D. E. (2014). The melanoma revolution: From UV carcinogenesis to a new era in therapeutics. Science, 346(6212), 945–949.https://doi.org/10.1126/science.1253735
Mattia, G., Puglisi, R., Ascione, B., Malorni, W., Carè, A., & Matarrese, P. (2018). Cell death‐based treatments of melanoma: Conventional treatments and new therapeutic strategies. Cell Death Dis, 9, 112. 1‐14
Miller, K. D., Siegel, R. L., Lin, C. C., Mariotto, A. B., Kramer, J. L., Rowland, H. L., … Jemal, A. (2016). Cancer treatment and survivorship statistics.CA Cancer J Clin, 66(4), 271–289. https://doi.org/10.3322/caac.21349
Nikolaou, V., & Stratigos, A. J. (2014). Emerging trends in the epidemiology of melanoma. Br J Dermatol, 170(1), 11–19. https://doi.org/10.1111/ bjd.12492
Pawelek, J. M. (1976). Factors regulating growth and pigmentation of melanoma cells. J Invest Dermatol, 66(4), 201–209. https://doi.org/ 10.1111/1523‐1747.ep12482134
Pawlikowska, M., Jędrzejewski, T., Piotrowski, J., & Kozak, W. (2016). Fever‐range hyperthermia inhibits cells immune response to proteinbound polysaccharides derived from Coriolus versicolor extract. Mol Immunol, 80, 50–57. https://doi.org/10.1016/j.molimm.2016.10.013
Pfisterer, P. H., Wolber, G., Efferth, T., Rollinger, J. M., & Stuppner, H. (2010). Natural products in structure‐assisted design of molecular cancer therapeutics. Curr Pharm Des, 16(15), 1718–1741. https://doi.org/ 10.2174/138161210791164027
Rajkumar, S., & Watson, I. R. (2016). Molecular characterisation of cutaneous melanoma: Creating a framework for targeted and immune therapies. Br J Cancer, 115(2), 145–155. https://doi.org/10.1038/ bjc.2016.195
Saleh, M. H., Rashedi, I., & Keating, A. (2017). Immunomodulatory properties of Coriolus versicolor: The Role of Polysaccharopeptide. Front Immunol, 8, 1087. 1‐12
Shah, D. J., & Dronca, R. S. (2014). Latest advances in chemotherapeutic, targeted, and immune approaches in the treatment of metastatic melanoma. Mayo Clin Proc, 89(4), 504–519. https://doi.org/10.1016/j.mayocp.2014.02.002
Slominski, A., Ermak, G., & Wortsman, J. (1999). Modification of melanogenesis in cultured human melanoma cells. In Vitro Cell Dev Biol Anim, 35, 564–565. https://doi.org/10.1007/s11626‐999‐0093‐6
Slominski, A., Paus, R., & Mihm, M. C. (1998). Inhibition of melanogenesis as an adjuvant strategy in the treatment of melanotic melanomas:selective review and hypothesis. Anticancer Res, 18, 3709–3715.
Slominski, A., Zbytek, B., & Slominski, R. (2009). Inhibitors of melanogenesis increase toxicity of cyclophosphamide and lymphocytes against melanoma cells. Int. J. Cancer, 124(6), 1470–1477. https://doi.org/ 10.1002/ijc.24005
Sniegocka, M., Podgorska, E., Plonka, P. M., Elas, M., Romanowska‐Dixon, B., Szczygiel, M., … Urbańska, K. (2018). Transplantable melanomas in hamsters and gerbils as models for human melanoma. Sensitization in melanoma radiotherapy—From animal models to clinical trials. Int J Mol Sci, 19(4). https://doi.org/10.3390/ijms19041048
Song, K. J., Jang, Y. S., Lee, Y. A., Kim, K. A., Lee, S. K., & Shin, M. H. (2011). Reactive oxygen species‐dependent necroptosis in Jurkat T cells induced by pathogenic free‐living Naegleria fowleri. Parasite Immunol, 33(7), 390–400. https://doi.org/10.1111/j.1365‐3024.2011.01297.x
Urabe, K., Aroca, P., Tsukamoto, K., Mascagna, D., Palumbo, A., Prota, G., & Hearing, V. J. (1994). The inherent cytotoxicity of melanin precursors: A revision. Biochim Biophys Acta, 1221(3), 272–278. https://doi.org/10.1016/0167‐4889(94)90250‐X van der Walt, N. B., Zakeri, Z., & Cronjé, M. J. (2016). The induction of apoptosis in A375 malignant melanoma cells by Sutherlandia frutescens. Evid Based Complement Alternat Med, 2016, 4921067:1‐14. https:// doi.org/10.1155/2016/4921067
Yang, B., Johnson, T. S., Thomas, G. L., Watson, P. F., Wagner, B., Furness, P. N., & El Nahas, A. M. (2002). A shift in the Bax/Bcl‐2 balance may activate caspase‐3 and modulate apoptosis in experimental glomerulonephritis. Kidney Int, 62(4), 1301–1313. https://doi.org/10.1111/ j.1523‐1755.2002.kid587.x
Zong, A., Cao, H., & Wang, F. (2012). Anticancer polysaccharides from natural resources: A review of recent research. Carbohyd Polym, 90(4),1395–1410. https://doi.org/10.1016/j.carbpol.2012.07.026