International Journal of Biological Macromolecules 58 (2013) 95–103 Contents lists available at SciVerse ScienceDirect International Journal of Biological Macromolecules journal homepage: www.elsevier.com/locate/ijbiomac Cytotoxic effect of Agaricus bisporus and Lactarius rufus ␤-d-glucans on HepG2 cells Amanda do Rocio Andrade Pires a , Andrea Caroline Ruthes a , Silvia Maria Suter Correia Cadena a , Alexandra Acco b , Philip Albert James Gorin a , Marcello Iacomini a,∗ a b Departamento de Bioquímica e Biologia Molecular, Universidade Federal do Paraná, CP 19046, CEP 81531-980, Curitiba, PR, Brazil Departamento de Farmacologia, Universidade Federal do Paraná, Curitiba, Paraná, Brazil a r t i c l e i n f o Article history: Received 30 January 2013 Accepted 16 March 2013 Available online 26 March 2013 Keywords: Agaricus bisporus Lactarius rufus (-d-Glucans Cytotoxicity HepG2 cells a b s t r a c t The cytotoxic activity of ␤-d-glucans isolated from Agaricus bisporus and Lactarius rufus fruiting bodies was evaluated on human hepatocellular carcinoma cells (HepG2). NMR and methylation analysis suggest that these ␤-d-glucans were composed of a linear (1→6)-linked and a branched (1→3), (1→6)-linked backbone, respectively. They both decreased cell viability at concentrations of up to 100 ␮g mL−1 , as shown by MTT assay. The amount of LDH released and the analysis of cell morphology corroborated these values and also showed that the ␤-d-glucan of L. rufus was more cytotoxic to HepG2 cells than that of A. bisporus. The treatment of HepG2 cells with L. rufus and A. bisporus ␤-d-glucans at a dose of 200 ␮g mL−1 for 24 h promoted an increase of cytochrome c release and a decrease of ATP content, suggesting that these polysaccharides could promote cell death by apoptosis. Both ␤-d-glucans were tested against murine primary hepatocytes at a dose of 200 ␮g mL−1 . The results suggest that the L. rufus ␤-d-glucan was as cytotoxic for hepatocytes as for HepG2 cells, whereas the A. bisporus ␤-d-glucan, under the same conditions, was cytotoxic only for HepG2 cells, suggesting cell selectivity. These results open new possibilities for use of mushroom ␤-d-glucans in cancer therapy. © 2013 Elsevier B.V. All rights reserved. 1. Introduction Hepatocellular carcinoma (HCC) has become the third most common cancer-related cause of death worldwide and the leading cause of death in patients with cirrhosis [1]. Many studies have been done to decrease the incidence of these deaths [2–5]. Human hepatocellular carcinoma cells (HepG2) are considered an excellent model for investigating a compound’s cytotoxicity [6]. However, many toxic compounds impair normal cellular functions, preventing their use. Therefore, assessing the effects of these compounds on normal cells, such as hepatocytes, can indicate selectivity. The medicinal health benefits and pharmacological potential of both edible and non-edible mushrooms have long Abbreviations: Ab, Agaricus bisporus linear (1→6)-linked ␤-d-glucan; ASA, acetylsalicylic acid; BSA, bovine serum albumin; DMSO, dimethylsulfoxide; HCC, hepatocellular carcinoma; HEPES, 4-(2-hydroxyethyl)-1-piperazine ethanesulfonic acid; LDH, lactate dehydrogenase; Lr, Lactarius rufus branched (1→3) (1→6)-linked ␤-d-glucan; MTT, 3-(4,5-dimethythiazol-2-yl)-2,5-diphenyltetrazolium bromide; TRIS, tris(hydroxymethyl)-aminomethane. ∗ Corresponding author. Tel.: +55 41 3361 1655; fax: +55 41 3266 2042. E-mail address: iacomini@ufpr.br (M. Iacomini). 0141-8130/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.ijbiomac.2013.03.040 been recognized, including their antitumor, immunostimulatory and anti-inflammatory activities. Because their constituents are believed to promote health and longevity, and also due to their taste, texture, high fiber, and low fat contents [7–10], mushrooms have long been consumed, especially in Asian countries. The biological importance of many mushrooms is primarily related to the great structural diversity of their polysaccharides. The mechanisms involved in the antitumor activity of polysaccharides are not fully known. Most mushroom polysaccharides seem to exert their antitumor activity via activation of multiple pathways in the host immune response, or by acting as biological response modifiers [11,12]. Promising antitumor activity has been attributed to several polysaccharides [12,13], and many of them are also used as adjuvants in chemotherapy treatment for different types of tumors. Among the best known are the commercial polysaccharides lentinan, derived from Lentinus edodes; PSK (krestin), extracted from Coriolus versicolor; and schizophyllan, produced by the fungus Schizophyllum commune [14]. Polysaccharides with antitumor action possess a specific chemical composition and molecular configuration and may be in the form of homo- or heteropolymers. Among the molecules with antitumor activity, most belong to the ␤-d-glucan group [11,15,16]. Their structure consists mainly of a ␤-(1→3)-linked main-chain 96 A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 with branches at O-6, which are believed to be necessary for their antitumor action [13]. Glucans with a high molecular weight seem to be more effective than those with a low molecular weight. The main polysaccharides found in mushrooms belong to the class of ␤-d-glucans [17–19], although heterogalactans [20–24] and xylomanans [25] can also be found. Considering that some polysaccharides extracted from mushrooms have antitumor activity, the effect of two different ␤-d-glucans on HepG2 cells was evaluated with the intent of showing potential cytotoxic effects and also identifying the pathways involved. To evaluate the influence of polysaccharide structures, a linear and a branched ␤-d-glucan, extracted from Agaricus bisporus and Lactarius rufus fruiting bodies, respectively, were tested. A. bisporus, commonly known as the button mushroom, is the most widely consumed mushroom and contains many compounds with medicinal properties, including polysaccharides [22,26–28]. In contrast, L. rufus is generally not recommended for consumption [29,30] because, like other mushrooms belonging to the genus Lactarius, it releases a milky juice, lactate, when the fruiting bodies are injured [31]. However, in some regions, this mushroom is used as a condiment after special treatment [32]. In the majority of Lactarius species, different types of sesquiterpenes play an important biological role, mainly constituting a chemical defense system against predators [31,33], meanwhile the biological potential of their polysaccharides is not well known [22]. Consequently, comparisons of the effects of isolated ␤-dglucans from A. bisporus and L. rufus on the viability and morphology of tumor cells and primary hepatocytes were carried out, and a selective antitumor activity was demonstrated. 2. Materials and methods 2.1. Reagents d-Mannitol, HEPES, EDTA, BSA, digitonin, penicillin, streptomycin, Tris, bicarbonate, MTT, collagenase type IV and 1A, glucagon, insulin, epidermal growth factor and trypsin were obtained from Sigma–Aldrich. High glucose DMEM, fetal bovine serum and hepatozyme were purchased from Gibco, and DMSO, KCl, sodium phosphate and potassium phosphate, together with other reagents of high purity, were purchased from Merck. 2.2. Polysaccharides 2.2.1. Biological material Dried fruiting bodies of A. bisporus were obtained from Makoto Yamashita Company (Miriam Harumi Yamashita), São José dos Pinhais, State of Paraná, Brazil. L. rufus fruiting bodies were collected in the middle of May 2005 from the soil of a Pinus sp. reforestation project located in Mafra, State of Santa Catarina, Brazil, at latitude: 26◦ 13′ S; longitude: 49◦ 50′ W and altitude of 826 m above sea level. These samples were cleaned, vacuum dried, and then ground to a powder. 2.2.2. Extraction and purification of ˇ-d-glucans Dried, Wiley-milled powder fruiting bodies from A. bisporus (Ab; 100 g) and L. rufus (Lr; 42.2 g) were each submitted to successive cold and hot aqueous extraction, both for 6 h (6×; approximately 1000 mL each). The cold water extractions were performed to remove cold water-soluble compounds, which were not used in the present study. The extracted polysaccharides were recovered from aqueous extracts by addition of excess ethanol (3:1; v/v), followed by centrifugation at 8000 rpm at 5 ◦ C for 20 min. The sediments were dialyzed against tap water for 24 h (12–14 kDa cut-off), concentrated under reduced pressure, and freeze-dried. The A. bisporus and L. rufus extracts were named HW-Ab and HW-Lr, respectively. Both crude extracts were fractionated by a freeze-thawing process [34], where Hw-Ab and HW-Lr fractions were dissolved in water, and the solutions were subjected to freezing and slow thawing until complete separation of cold-water soluble (SHW) and insoluble polysaccharides (IHW). An A. bisporus insoluble fraction (IHW-Ab), obtained after centrifugation (8000 rpm at 5 ◦ C for 20 min), was treated with dimetylsulfoxide (50 mL) for 2 h at 60 ◦ C, dialyzed against tap water for 24 h, and then resubmitted to the freeze-thawing process, resulting in a soluble ␤-d-glucan fraction, named Ab [35]. Meanwhile, to obtain L. rufus ␤-d-glucan, its cold water soluble fraction was treated with Fehling solution [36]. The Fehling-soluble fraction (FSHW) was isolated with an insoluble Cu2+ complex, centrifuged under the same conditions as described above, neutralized with HOAc, dialyzed against tap water, deionized with mixed-ion exchange resins and then freeze dried. This process resulted in the isolation of a ␤-d-glucan, named Lr [37]. The isolated ␤-d-glucans were dissolved in phosphate-buffered solution (PBS) at 1 mg mL−1 and sterilized by heat at 100 ◦ C for 90 min for assays with cells. 2.3. Monosaccharide composition of polysaccharide fractions Each polysaccharide fraction (1 mg) was hydrolyzed with 2 M TFA at 100 ◦ C for 8 h, followed by evaporation. The resulting residue was reduced with NaBH4 (1 mg) and acetylated with Ac2 O–pyridine (1:1, v/v; 200 ␮L) at 100 ◦ C for 30 min following the method of Sassaki et al. [38]. The resulting alditol acetates were analyzed by gas chromatography–mass spectrometry (GC–MS), using a Varian model 3300 gas chromatograph linked to a Finnigan Ion-Trap, Model 810-R12 mass spectrometer. A DB-225 capillary column (30 m × 0.25 mm i.d.) held from 50 ◦ C during injection and then programmed to 220 ◦ C (constant temperature) at 40 ◦ C min−1 was used for qualitative and quantitative analysis of alditol acetates. The alditol acetates were identified by their typical retention times and electron impact profiles. 2.4. Methylation analysis of ˇ-d-glucans Per-O-methylation of isolated ␤-d-glucans (Ab and Lr; 10 mg) was carried out using NaOH–Me2 SO–MeI as described by Ruthes et al. [39]. This process, after isolation of the products by neutralization (HOAc), dialysis, and evaporation, was repeated, and the methylation was then complete. The per-O-methylated derivatives were hydrolyzed with 45% formic acid (HCO2 H, 1 mL) for 15 h at 100 ◦ C [18], followed by NaBD4 reduction and acetylation as above (Section 2.3), to give a mixture of partially O-methylated alditol acetates, which was analyzed by GC–MS using a Varian model 4000 gas chromatograph, equipped with CP-Sil-43CB fused silica capillary columns. The injector temperature was maintained at 210 ◦ C, with the oven starting at 50 ◦ C (hold 2 min) to 90 ◦ C (20 ◦ C min−1 , then held for 1 min), 180 ◦ C (5 ◦ C min−1 , then held for 2 min) and to 210 ◦ C (3 ◦ C min−1 , then held for 5 min). Helium was used as the carrier gas at a flow rate of 1.0 mL min−1 . Partially O-methylated alditol acetates were identified from m/z by comparing their positive ions with standards, the results being expressed as a relative percentage of each component [40]. 2.5. Nuclear magnetic resonance (NMR) spectroscopy Monodimensional (13 C and DEPT) NMR spectra were prepared using a 400 MHz Bruker model DRX Avance spectrometer incorporating Fourier transform. Analyses were performed at 70 ◦ C on samples dissolved in D2 O or Me2 SO-d6 . Chemical shifts of water-soluble samples are expressed in ı (ppm) relative to A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 acetone at ı 30.20 (13 C), and at ı 39.70 (13 C) for those dissolved in Me2 SO-d6 . 2.6. Animals Male Wistar rats (180–200 g) were obtained from the Central Animal House of the Federal University of Parana (PR, Brazil). The animals were housed at 22 ± 1 ◦ C under a 12 h light–12 h dark cycle and had free access to standard laboratory food (Purina® ) and tap water. The experiments were conducted following the recommendation of Brazilian Law 6638, 05/11/1979 for the scientific management of animals, and the procedures were approved by the Institutional Animal Ethics Committee (number 548). 2.7. HepG2 cell culture and treatment HepG2 cells were acquired from the Rio de Janeiro Cell Banc, Brazil. They were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM)-high glucose (Gibco) supplemented with 10% fetal bovine serum, 100 U mL−1 penicillin G, 100 ␮g mL−1 streptomycin, 20 mmol L−1 HEPES, adjusted to pH 7.4 with 1 mol L−1 sodium bicarbonate. HepG2 cells were grown in poly-l-lysinecoated flasks in a humidified incubator in the presence of 5% CO2 at 37 ◦ C. Sub-culturing was performed at approximately 48 h intervals, and cell growth was monitored with an Olympus inverted microscope. Polysaccharide treatments occurred 24 h after the cells were seeded on the plates. The ␤-d-glucans were dissolved in culture medium at 50, 100, 200 or 400 ␮g mL−1 and incubated for 12, 24 or 48 h. 2.8. Isolation of primary murine hepatocyte, culture and treatment Hepatocytes were obtained under monovascular liver perfusion of Wistar rats, as described previously by Seglen [41], with modifications. Male rats were weighed and anesthetized intraperitoneally with a mixture of cetamine (60 mg kg−1 ) and xylazine (7.5 mg kg−1 ). Following laparotomy, 100 ␮L of sodium heparin (5000 U mL−1 ) were injected into the abdominal cava vein. The portal and thoracic cava veins were cannulated, and the liver was perfused with Krebs solution (NaCl 2.399 mol L−1 ; KCl 96 mmol L−1 ; KH2 PO4 24 mmol L−1 ; MgSO4 24 mmol L−1 ; NaHCO3 480 mmol L−1 and HEPES buffer 1 mol L−1 , pH 7.4) containing CaCl2 1.3 mol L−1 , collagenase (type IA and IV) and carbogen (O2 95%; CO2 5%). The liver was excised, and the cells were released by mechanical action, filtered through 50 ␮m nylon membranes and centrifuged at 400 rpm for 5 min at 4 ◦ C. Subsequently, the cells were centrifuged four times with Krebs solution supplied with 20% bovine serum albumin (BSA), also treated with carbogen. The cells were suspended in DMEM high glucose medium supplied with bovine fetal serum (10%), insulin (100 nmol L−1 ), glucagon (10 nmol L−1 ), epidermal growth factor (10 ng mL−1 ), dexamethasone (50 nmol L−1 ), penicillin (100 U mL−1 ) and streptomycin (100 ng mL−1 ). The viability was determined using the Trypan blue (0.4%, w/v) exclusion method, previously described by Philips [42]. Only cell suspensions with viabilities higher than 80% were plated (1 × 106 cells/plate on a 60 mm plate) and cultured for further experiments. Four hours after plating, the medium was replaced by Hepatozyme® , with or without polysaccharide, and incubated for 18 h prior to assays. 2.9. Viability assays HepG2 cells (1 × 104 cells/well) were seeded in 96-well plates and cultured in a humidified incubator overnight for adhesion. 97 The medium was replaced by medium with or without polysaccharides at 50–400 ␮g mL−1 for 24, 48 or 72 h. After incubation, the medium was discarded, and a tetrazolium dye (MTT) solution (0.75 mg mL−1 ) was added. For hepatocyte assays, cells were only treated with the polysaccharides for 24 h. The plates were allowed to stand at 37 ◦ C for 3 h. The MTT-formazan crystals that formed were dissolved in DMSO, and absorbance was measured at 540 nm using a microplate reader [3,43]. Cell viability is expressed as percent of control. Under the same conditions, the viability of HepG2 cells and hepatocytes was also determined by releasing LDH into the culture medium, using an LDH assay kit (Labtest) according to the manufacturer’s recommended protocol. After the polysaccharide treatment, the supernatant was collected and centrifuged at 1200 rpm for 10 min, and LDH activity was measured. The results are expressed as a percentage of the control. 2.10. Morphological analysis The morphological changes in hepatocytes and HepG2 cells were examined with an Axiovert 40CSFL inverted microscope with 20× magnification. 2.11. Measurement of ATP content The intracellular ATP content was determined using an ATP Bioluminescent assay kit from Sigma-Aldrich (FLAA) according to the manufacturer’s protocol. The ATP measurement is based on the oxidation of d-luciferin catalyzed by firefly luciferase enzyme, which consumes ATP with concomitant light emission. Bioluminescence was measured with a TECAN Infinite reader, and the ATP concentration (nmol L−1 ) was calculated using a standard curve (ATP ranging from 0 to 100 ␮mol L−1 ). The results are expressed as a percentage of the control. 2.12. Measurement of cytochrome c release Cytochrome c release was monitored spectrophotometrically by absorbance variation at 414 nm according to Appaix et al. [44]. HepG2 cells (1 × 106 /plate) were seeded in a 60 mm plate and incubated with polysaccharides for 24 h. After incubation, cells were harvested using trypsin solution and centrifuged at 1200 rpm for 10 min at 4 ◦ C. The pellet was suspended in a medium containing sucrose (50 mmol L−1 ), Tris–HCl (10 mmol L−1 ) and EGTA (3 mmol L−1 ). Cells were permeabilized with digitonin at 10 ␮mol mL−1 for 30 min at 4 ◦ C. The cell suspension was centrifuged at 10,000 × g for 10 min. The supernatant was filtered through a 22 ␮m individual filter, and optical density was measured at 414 nm, considering ε = 100 mM−1 cm−1 . The results are expressed as a percentage of the control. 2.13. Protein determination The protein concentration in samples was determined using the Bradford method [45]. 2.14. Statistical analysis The data were statistically analyzed using variance analysis and Tukey’s test for average comparison using GraphPad Prism software. Mean values ± S.D. were used; values were considered significant at P < 0.05. 98 A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 Fig. 1. 13 C NMR spectra of glucans Ab (A) and Lr (B) with inserts of DEPT–CH2 inversion (A′ and B′ , respectively). ␤-d-Glucans were prepared in Me2 SO-d6 (A) and D2 O (B) at 70 ◦ C (chemical shifts are expressed in ı ppm). 3. Results and discussion 3.1. ˇ-d-Glucans structural characterization The ␤-d-glucans obtained from A. bisporus (Ab; 8.0 g, 8%) and L. rufus (Lr; 0.6 g, 1.4%) were shown to be homogeneous on HPSEC, with molecular weights of 2.9 × 104 g mol−1 and 11.3 × 104 g mol−1 , respectively. Each purified Ab and Lr fraction contained only glucose as their monosaccharide component (GC–MS). GC–MS analysis of their partially O-methylated alditol acetates suggested that the Ab fraction was composed of a linear (1→6)-linked ␤-d-glucan due to the presence of only 2,3,4-Me3 Glc (100%). In contrast, the Lr fraction was composed of a branched (1→3), (1→6)-linked ␤-d-glucan due to the presence of 2,3,4,6Me4 -Glc (32.0%), 2,4,6-Me3 -Glc (36.4%), and 2,4-Me2 -Glc (31.6%), O-methyl alditol acetates. 13 C NMR spectroscopy of fraction Ab showed a (1→6)-linked ␤-glucan by the presence of only six signals (Fig. 1A). All carbon frequencies were assigned according to literature data [46], and the ␤-configuration was confirmed by the high frequency of its anomeric carbon at ı 103.3 [47]. The other carbon frequencies were assigned as C-2 (ı 73.5), C-3 (ı 76.7), C-4 (ı 70.2), C-5 (ı 75.5), and C-6 (ı 68.6) [35], as shown in Fig. 1A. The O-substituted −CH2 (C-6) signal was confirmed by its inversion in a DEPT-13 C NMR spectrum (Fig. 1A′ ). On the other hand, NMR spectroscopy of fraction Lr confirmed the presence of a branched (1→3), (1→6)-linked ␤-d-glucan (Fig. 1B). Its 13 C NMR spectrum showed three distinct signals in the anomeric region at ı 103.0, 102.9 and 102.6, corresponding to the nonreducing end-, 3-O- and 3,6-di-O-substituted residues, respectively. The ␤-configuration was also confirmed by high-frequency C-1 signals (ı 103.1, 102.9 and 102.6) (Fig. 1B) [47]. The glycosidic linkages of fraction Lr were shown by the presence of 3-O-substituted signals at ı 85.5, 85.1, 84.9 and 84.7 (Fig. 1B), while the O-6 substitution was confirmed from the respective inverted peak in its DEPT spectrum (Fig. 1B). After a controlled Smith degradation, the backbone structure of Lr glucan was shown by formation of its linear main-chain of a (1→3)-linked ␤-d-glucan, due to the presence of typical 13 C signals at ı 102.9 (C-1); 86.2 (C3); 76.5 (C-5); 72.8 (C-2); 68.5 (C-4), and 61.0 (C-6) [37,48]. In the degraded fraction, low intensity signals from O-6 substitution were still present, indicating that this glucan is mostly substituted at O-6 by single units of ␤-d-glucopyranose with a minor proportion by side chains of 3-O-substituted ␤-d-Glcp groups [37]. 3.2. Cellular viability Cell viability was evaluated by two methods, MTT and LDH release assays. The results of the MTT assays for HepG2 cells are shown in Fig. 2A–C. The ␤-d-glucan from L. rufus (Lr) was cytotoxic after 24 h of incubation at concentrations above 100 ␮g mL−1 , decreasing viability up to ∼30% at the highest concentration (400 ␮g mL−1 ; Fig. 2A). Meanwhile, A. bisporus ␤-d-glucan (Ab) was more toxic than Lr, promoting cell death at 50 ␮g mL−1 . At the highest concentration (400 ␮g mL−1 ), the polysaccharide decreased cell viability by ∼70%. These effects on HepG2 cell viability were more pronounced when the incubation with the polymers A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 99 Fig. 2. Cell viability assay by MTT. Conditions for this assay are described in Section 2. In vitro cytotoxicity of Lr and Ab against HepG2 cells after incubation of 24 h (A), 48 h (B) and 72 h (C) and primary hepatocyte after 18 h incubation (D). The results represent the mean ± S.D. of four different experiments. Asterisks indicate significant difference (***P ≤ 0.0001) relative to untreated control cells. was carried out for 48 h and 72 h (Fig. 2B and C). However, at these times, the toxicity of Lr and Ab at the highest concentration tested (400 ␮g mL−1 ) were equivalent, promoting almost complete cell death. These results demonstrate that both ␤-d-glucans tested were toxic to HepG2 cells and that this cytotoxicity is dependent on concentration and time of incubation. Considering the high cytotoxicity of the polymers after the longest periods of treatment (48 and 72 h), for the next experiments, the concentration of 200 ␮g mL−1 was used with an incubation time of 24 h. The viability of hepatocytes evaluated by the MTT method is shown in Fig. 2D. In these assays, the time of treatment was limited to 24 h, and the concentration of polysaccharides was limited to 200 ␮g mL−1 . In Fig. 2D, only Lr was cytotoxic for primary hepatocytes, decreasing cell viability by ∼75%. These results corroborate the effects of polysaccharides observed in HepG2 cells, where Lr was more cytotoxic on these cells under the same conditions (200 ␮g mL−1 and 24 h of incubation). Interestingly, while Ab (200 ␮g mL−1 and 24 h of incubation) was able to decrease HepG2 cell viability by ∼10% (Fig. 2A), it did not affect hepatocyte viability (Fig. 2D). The LDH assay (Fig. 3) also showed that Lr and Ab were differentially cytotoxic to HepG2 cells (Fig. 2). Treatment with ␤-d-glucans promoted an increase of LDH release that reflects a decrease in cell viability. In this assay, after 24 h incubation (Fig. 3A), Lr promoted an increase of LDH release of ∼29% at the lowest concentration (50 ␮g mL−1 ) and reached a release of ∼56% at 400 ␮g mL−1 , in comparison with the control. After 48 h (Fig. 3B) and 72 h (Fig. 3C), the values of LDH release are similar to those of the 24 h incubation (Fig. 3A), suggesting that this effect is independent of the incubation time. A similar profile was seen for Ab treatment. At 24 h, Ab promoted a ∼30% increase of LDH release at 50 ␮g mL−1 , reaching ∼35% at 200 ␮g mL−1 (Fig. 3A). These values remained unchanged after 48 h incubation (Fig. 3B) with Ab; however, a decrease after 72 h was observed, with LDH release near to 25% at 100 and 200 ␮g mL−1 . At the highest concentration (400 ␮g mL−1 ) Ab increased LDH release by ∼17% and ∼24% after 24 h and 48 h of incubation, respectively. Interestingly, after 72 h incubation (Fig. 3C), 50 and 400 ␮g mL−1 Ab was not significantly different from the control. Compared with the results of the MTT assay, the viability values measured by LDH leakage were less. The nature and difference of sensitivity between these methods may explain these differences. The MTT assay is based on enzymatic conversion of tetrazolium salt into formazan by mitochondrial dehydrogenases [49], whereas the LDH leakage assay is based on the release of the enzyme into the culture medium after cell membrane ruptures during apoptosis [50]. Fotakis and Timbrell [51] tested four different methods to evaluate the viability of HepG2 cells against cadmium chloride: MTT, LDH, neutral red and protein quantification. All methods showed results with different intensities, and the authors suggest that the MTT is more sensitive than the LDH assay. These data are in accordance with the results obtained in the present work. LDH release was also measured with primary hepatocytes, and the results corroborate the results of the MTT assay. The LDH release results are shown in Fig. 3D, where a great increase of LDH release in cells treated with Lr (∼58%) can be seen, while the incubation with Ab did not produce a significant effect. These data suggest that Ab may act selectively, being more aggressive to tumors (HepG2) than to normal cells (hepatocytes), whereas Lr is toxic to both. 100 A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 Fig. 3. Cell viability assay by LDH release. Conditions for LDH release measurements are described in Section 2. LDH release of HepG2 cells treated with Lr and Ab after incubation for 24 h (A), 48 h (B) and 72 h (C) and primary hepatocyte after incubation for 18 h (D). Note that 100% corresponds to 6.491 ± 0.101 ␮mol L−1 of NADH consumed per minute. The results represent the mean ± S.D. of four different experiments. Asterisks indicate significant difference (***P ≤ 0.0001) relative to untreated control cells. 3.3. Cell morphology Fig. 4A–C shows the morphology of HepG2 cells in the absence (Fig. 4A-control) or presence of Lr (Fig. 4B) and Ab (Fig. 4C) at a dose of 200 ␮g mL−1 incubated for 24 h. Fig. 4A (control) shows a high contact and confluent culture of elongated HepG2 cells. Treatment with Lr (Fig. 4B) and Ab (Fig. 4C) fractions promoted a reduction in cell adhesion and increased the number of cells with a round morphology (indicated by the arrows), which is suggestive of cell distress. These effects were more pronounced for Lr-treated cells (Fig. 4C) and are in agreement with the effects of polysaccharides on HepG2 viability (Figs. 2A and 3A), in which Lr was the most cytotoxic (200 ␮g mL−1 and 24 h of incubation). Assays with primary hepatocytes (Fig. 4D–F) are also in agreement with the viability results (Figs. 2D and 3D). Fig. 4D shows a highly confluent culture with hepatocytes with a typical cubic cell shape and often two nuclei. The treatment with Lr promoted significant alterations in cell morphology (Fig. 4E), with a predominance of round cells and dark features, suggesting cell distress. These results coincide with the viability decrease (∼75%; Fig. 2D). The treatment with Ab (Fig. 4F) did not promote significant changes in hepatocyte morphology, when compared with the control assay (Fig. 4D), and corroborates the viability results (Fig. 2D). consumes energy, thus ATP levels are expected to decrease in comparison with necrotic cells. The ATP content of HepG2 cells treated with ␤-d-glucans is shown in Fig. 5. In this assay, the polysaccharides were tested at a dose of 200 ␮g mL−1 and incubated for 24 h before harvesting. Acetylsalicylic acid (ASA) was used as positive control because it is able to induce cell death of HepG2 by apoptosis [53]. It was used at a concentration of 10 mmol L−1 and incubated for 24 h. In this condition, ASA treatment promoted a reduction of ATP content by ∼45% in comparison with the control (absence of treatment). In turn, Lr and Ab treatment decreased the levels of ATP by ∼14% and ∼27%, respectively, and the statistical data show that, in comparison, this effect is different for the two polysaccharides (data not shown). These results are consistent with the effects of Lr on viability (Fig. 2A) and morphology (Fig. 4B); however, Ab is less effective than Lr in modulating viability and morphology of HepG2 cells under the same experimental conditions (Figs. 2A and 4C, respectively). Here, it is important to consider that the methodologies used are different and, therefore, do not have the same sensitivity. At first glance, these results are unexpected because Lr was more toxic to HepG2 cells than Ab. However, Lr may be able to induce cell death, but not by apoptosis pathway. Thus, Ab may be impairing the ATP pathway, therefore decreasing the ATP content. 3.4. ATP content 3.5. Cytochrome c release During the process of cell death, energetic and metabolic statuses are modified according to the related pathway, particularly necrosis or apoptosis [52]. Apoptosis is an organized process that Cytochrome c is released from mitochondria in response to cell injury and triggers apoptosis [54]. High levels of cytochrome c in the cytoplasm indicate that the cells are undergoing apoptosis. In our A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 101 Fig. 4. Cell morphology assay. Conditions for morphology assay are described in Section 2. HepG2 cells were incubated with Lr and Ab for 24 h, and primary hepatocytes were incubated with Lr and Ab for 18 h. In sequence images were obtained using inverted microscopy. The images represent (A) untreated HepG2 cells; (B) Lr treated HepG2 cells; (C) Ab treated HepG2 cells; (D) untreated primary hepatocytes; (E) Lr treated primary hepatocyte; (F) Ab treated primary hepatocytes. Arrows indicate round cells. assay, ASA was used as a positive control, promoting an increase in cytochrome c release of ∼90% (Fig. 6). Treatment with Lr and Ab also increased cytochrome c release at ∼66% and ∼20%, respectively. Therefore, in this assay, Lr was also more effective than Ab at increasing cytochrome c release, which corroborates the viability and morphology results (Figs. 2A and 4B and C). These results reinforce the suggestion that Lr is able to induce HepG2 cell death by apoptosis. 4. Discussion The present research shows, for the first time, the cytotoxic effects of ␤-d-glucans from L. rufus and A. bisporus on human Fig. 5. Measurement of ATP content of HepG2 cell treated with Lr and Ab glucans. Conditions for ATP measurement are described in Section 2. HepG2 cells were incubated with ASA (10 mmol L−1 ), Lr (200 ␮g mL−1 ) or Ab (200 ␮g mL−1 ) for 24 h before ATP measurement. Untreated HepG2 cells were used as a control, and 100% corresponds to 1.612 ± 0.132 nmol of ATP per mg protein. The results represent the mean ± S.D. of three different experiments. Asterisks indicate significant difference (*P ≤ 0.05 and ***P ≤ 0.0001) relative to control cells. liver hepatocellular carcinoma cells (HepG2). Both polymers were cytotoxic, which may occur by induction of cell death by apoptosis, as shown by the ATP and cytochrome c assays. More specific assays are necessary to confirm these results; however, other polysaccharides are known to decrease viability of tumor cells by induction of apoptosis. An example is a protein-bound polysaccharide (PSK) that is derived from the fungus C. versicolor [55]. Treatment with 100 ␮g mL−1 of PSK promoted decreased cell viability of many cells line: B16F10, B9 murine MCA-induced fibrosarcoma, Ando-2 human melanoma, AGS human gastric cancer, A-549 human lung cancer, Hela and Jurkat T lymphoma leukemia. An assay with annexin V/7ADD double staining and caspases-3 expression showed that PSK promoted apoptosis. The authors suggested that PSK has in vitro cytotoxic activity on tumor Fig. 6. Measurement of cytochrome c release of HepG2 cell treated with Lr or Ab glucans. Conditions for cytochrome c measurement are described in Section 2. HepG2 cells were incubated with ASA (10 mmol L−1 ), Lr (200 ␮g mL−1 ) or Ab (200 ␮g mL−1 ) for 24 h before cytochrome c measure. Untreated HepG2 cells were used as a control, and 100% corresponds to 0.1357 ± 0.0383 nmol of cytochrome c per mg protein. The results represent the mean ± S.D. of three different experiments. Asterisks indicate a significant difference (***P ≤ 0.0001) relative to control cells. 102 A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 cell lines and that this effect is independent of PSK’s previously described immunomodulatory activity. They also affirmed that the antitumor activity is due to cytotoxic and immunomodulatory effects. Similarly, an aqueous extract of Phellinus gilvus inhibited proliferation of murine melanoma cells (B16F10) in vitro [56]. A dose-dependent effect of this polysaccharide extract was significant in doses up to 50 ␮g mL−1 and reached ∼80% inhibition at 150 ␮g mL−1 . Immunostained and TUNEL assays showed that a P. gilvus extract decreased proliferation through apoptosis induction. It was suggested that its anti-tumor action is due the induction of tumor cell apoptosis and that P. gilvus is much less toxic when compared with chemical anti-tumor drugs. The mechanism of anti-tumor action of polysaccharides has not yet been elucidated, but some investigations [55,56] have shown that the anti-tumor effect of polysaccharides may be due a combination of immunomodulatory and cytotoxic activities, although other pathways may be involved. Recently it was shown that a polysaccharide, MEP-II, isolated from the fermentation broth of Morchella esculenta, was cytotoxic to HepG2 cells. In that work, MEP-II was tested in a range of 150–1200 ␮g mL−1 against HepG2, AS49 and HeLa cells and was found to be more toxic to HepG2 cells. Annexin V-FITC/PI double stain, morphological analysis and mitochondrial potential results suggests that MEP-II promotes apoptosis in HepG2. The measurement of reactive oxygen species (ROS) formation using a DCFH-DA probe showed an increase in the accumulation of ROS in a concentration-dependent manner. Pre-incubation with N-acetyl-l-cysteine, an antioxidant, prevented ROS formation and decreased the externalization of phosphatidylserine. The authors suggest that ROS were most likely the crucial upstream signaling molecule for MEP-II-induced apoptosis [4]. The mechanisms of Ab and Lr antitumor activity are not yet understood, and the antitumor activity of other polysaccharides, such as those from mushrooms, can be due to a combination of effects. Another important aspect is the possible selective effect of these polymers. The results indicate that the most effective polymer (Lr) was also cytotoxic for primary hepatocytes. However, ␤-d-glucan Ab showed a cytotoxic effect on HepG2 cells without affecting the viability of hepatocytes. This difference in the intensity of cytotoxic action may be related to their structure because Lr has a branched structure, whereas Ab is a linear ␤-d-glucan. These branches may favor interaction with cell receptors and may be necessary for the induction of cell death. This interaction may be more effective in tumor cells than normal cells. Compounds that have toxic effects on tumors and not on normal cells are very promising and promote an increased interest in the application of ␤-d-glucans in cancer therapy. 5. Conclusion The ␤-d-glucans from L. rufus and A. bisporus were toxic against human liver hepatocellular carcinoma cells (HepG2), and this may occur by the induction of cell death by apoptosis. The L. rufus ␤-d-glucan was the most cytotoxic to HepG2 cells and was also cytotoxic to primary hepatocytes. However, A. bisporus ␤-d-glucan showed a cytotoxic effect on HepG2 cells without affecting hepatocyte viability. These results may open new biological applications for mushroom ␤-d-glucans. Acknowledgements The authors thank the Brazilian funding agencies CAPES (Coordenação de Aperfeiçoamento de Pessoal de Nível Superior) and CNPq (Conselho Nacional de Desenvolvimento Científico e Tecnológico) for financial support. References [1] F. Lodato, G. Mazzella, D. Festi, F. Azzaroli, A. Colecchia, E. Roda, World Journal of Gastroenterology 12 (2006) 71239–71249. [2] V. Balasubramaniyan, R. Shukla, G. Murugaiyan, R.R. Bhonde, N. Nalini, Biochimica et Biophysica Acta 1770 (2007) 1136–1144. [3] C.S. Cheung, K.K. Chung, J.C. Liu, C. Lau, P. Hon, J.Y. Chan, K. Fung, S.W. Au, Cancer Letters 253 (2007) 224–235. [4] M. Hu, Y. Chen, C. Wang, H. Cui, P. Duan, T. Zhai, Y. Yang, S. Li, Biotechnology Letters 35 (2012) 1–10. [5] Y. Hu, Y. Yang, Q. You, W. Liu, H. Gu, L. Zhao, K. Zhang, W. Wang, X. Wang, Q. Guo, Biochemical and Biophysical Research Communications 351 (2006) 521–527. [6] M. Pinti, L. Troiano, M. Nasi, R. Ferraresi, J. Dobrucki, A. Cossarizza, Journal of Biological Regulators and Homeostatic Agents 17 (2003) 166–171. [7] S. Chang, in: Royse (Ed.), Equality and Mutual Benefit, Mushroom Biology and Mushroom Products, 1996, pp. 1–10. [8] P.C.K. Cheung, Journal of Agricultural and Food Chemistry 44 (1996) 468–471. [9] J. Liu, T. Huang, M. Hsu, C. Chen, W. Lin, F. Lu, W. Chang, Toxicology and Applied Pharmacology 201 (2004) 186–193. [10] F.S. Reis, L. Barros, A. Martins, I.C.F.R. Ferreira, Food and Chemical Toxicology 50 (2012) 191–197. [11] A.T. Borchers, J.S. Stern, R.M. Hackman, C.L. Keen, M.E. Gershwin, Proceedings of the Society for Experimental Biology and Medicine 221 (1999) 281–293. [12] H.X. Wang, W.K. Liu, T.B. Ng, V.E.C. Ooi, S.T. Chang, Life Sciences 57 (1995) 269–281. [13] S.V. Reshetnokov, S.P. Wasser, K.K. Tan, International Journal of Medicinal Mushrooms 3 (2001) 361–394. [14] A.T. Borchers, C.L. Keen, M.E. Gershwin, Experimental Biology and Medicine 229 (2004) 393–406. [15] T. Mizuno, T. Hagiwara, T. Nakamura, H. Ito, K. Shimura, T. Sumiya, A. Asakura, Agricultural Biology and Chemistry 54 (1990) 2889–2896. [16] T. Sasaki, N. Takasura, Carbohydrate Research 47 (1976) 99–104. [17] E.R. Carbonero, A.H.P. Gracher, F.R. Smiderle, F.R. Rosado, G.L. Sassaki, P.A.J. Gorin, M. Iacomini, Carbohydrate Polymers 66 (2006) 252–257. [18] E.R. Carbonero, A.C. Ruthes, C.S. Freitas, P. Utrilla, J. Gálvez, E.V. Silva, G.L. Sassaki, P.A.J. Gorin, M. Iacomini, Carbohydrate Polymers 90 (2012) 814–819. [19] P. Manzi, A. Aguzzi, L. Pizzoferrato, Food Chemistry 73 (2001) 321–325. [20] E.R. Carbonero, A.H.P. Gracher, D.L. Komura, R. Marcon, C.S. Freitas, C.H. Baggio, A.R.S. Santos, G. Torri, P.A.J. Gorin, M. Iacomini, Food Chemistry 111 (2008) 531–537. [21] D.L. Komura, E.R. Carbonero, A.H.P. Gracher, C.H. Baggio, C.S. Freitas, R. Marcon, A.R.S. Santos, P.A.J. Gorin, M. Iacomini, Bioresource Technology 101 (2010) 6192–6199. [22] A.C. Ruthes, Y.D. Rattmann, E.R. Carbonero, P.A.J. Gorin, M. Iacomini, Carbohydrate Polymers 87 (2012) 1620–1627. [23] F.R. Smiderle, L.M. Olsen, E.R. Carbonero, R. Marcon, C.H. Baggio, C.S. Freitas, A.R.S. Santos, G. Torri, P.A.J. Gorin, M. Iacomini, Phytochemistry 69 (2008) 2731–2736. [24] S.P. Wasser, Applied Microbiology and Biotechnology 60 (2002) 258–274. [25] F.R. Smiderle, E.R. Carbonero, C.G. Mellinger, G.L. Sassaki, P.A.J. Gorin, M. Iacomini, Phytochemistry 67 (2006) 2189–2196. [26] S.C. Jeong, Y.T. Jeong, B.K. Yang, R. Islam, S.R. Koyyalamudi, G. Pang, K.Y. Cho, C.H. Song, Nutrition Research 30 (2010) 49–56. [27] D. Kent, C.M. Sheridan, H.A. Tomkinson, S.J. White, P. Hiscott, L. Yu, I. Grierson, Wound Repair and Regeneration 11 (2003) 285–291. [28] L. Yu, D.G. Fernig, J.A. Smith, J.D. Milton, J.M. Rhodes, Cancer Research 153 (1993) 4627–4632. [29] D. Arora, Mushrooms Demystified: A Comprehensive Guide to the Fleshy Fungi, Ten Speed Press, Berkley, CA, 1986. [30] W.C. Roody, Mushrooms of West Virginia and the Central Appalachians, University Press of Kentucky, Lexington, KY, 2003102. [31] D. Luo, F. Wang, X. Bian, J. Liu, The Journal of Antibiotics 58 (2005) 456–459. [32] R. Phillips, Mushrooms, Pan MacMillan, London, 2006. [33] O. Sterner, R. Bergman, J. Kihlberg, B. Wickberg, Journal of Natural Products 48 (1985) 279–288. [34] P.A.J. Gorin, M. Iacomini, Carbohydrate Research 128 (1984) 119–131. [35] F.R. Smiderle, G. Alquini, M.Z. Tadra-Sfeir, M. Iacomini, H.J. Wichers, L.J.L.D. Van Griensven, Carbohydrate Polymers (2012), http://dx.doi.org/10.1016/j.carbpol. 2012.12.073. [36] J.K.N. Jones, R.J. Stoodley, Methods in Carbohydrate Chemistry 5 (1965) 36–38. [37] A.C. Ruthes, E.R. Carbonero, M.M. Córdova, C.H. Baggio, A.R.S. Santos, G.L. Sassaki, T.R. Cipriani, P.A.J. Gorin, M. Iacomini, Carbohydrate Polymers (2013), http://dx.doi.org/10.1016/j.carbpol.2013.01.026. [38] G.L. Sassaki, L.M. Souza, R.V. Serrato, T.R. Cipriani, P.A.J. Gorin, M. Iacomini, Journal of Chromatography A 1208 (2008) 215–222. [39] A.C. Ruthes, D.L. Komura, E.R. Carbonero, G.L. Sassaki, P.A.J. Gorin, M. Iacomini, Carbohydrate Polymers 81 (2010) 29–34. [40] G.L. Sassaki, P.A.J. Gorin, L.M. Souza, P.A. Czelusniak, M. Iacomini, Carbohydrate Research 340 (2005) 731–739. [41] P.O. Seglen, in: D.M. Prescott (Ed.), Methods in Cell Biology, Academic Press, New York, 1976. [42] H.J. Philips, in: P.F. Kruse Jr., M.K. Patterson Jr. (Eds.), Tissue Culture Methods and Applications, Academic Press, New York, 1973, pp. 406–408. [43] T.P. Reilly, F.H. Bellevue, P.M. Woster, C.K. Svensson, Biochemical Pharmacology 55 (1998) 803–810. A.d.R.A. Pires et al. / International Journal of Biological Macromolecules 58 (2013) 95–103 [44] F. Appaix, M. Minatchy, C. Riva-Lavieille, J. Olivares, B. Antonsson, V.A. Saks, Biochimica et Biophysica Acta 1456 (2000) 75–181. [45] M. Bradford, Analytical Biochemistry 72 (1976) 248–254. [46] E. Barreto-Bergter, P.A.J. Gorin, Advances in Carbohydrate Chemistry and Biochemistry 41 (1983) 67–103. [47] L.D. Hall, L.F. Johnson, Journal of the Chemical Society, Chemical Communications (1969) 509–510. [48] P.A.J. Gorin, Advances in Carbohydrate Chemistry and Biochemistry 38 (1981) 13–104. [49] T. Mosmann, Journal of Immunological Methods 65 (1983) 55–63. [50] T. Decker, M.L. Lohmann-Matthes, Journal of Immunological Methods 115 (1988) 55–63. 103 [51] G. Fotakis, J.A. Timbrell, Toxicology Letters 160 (2006) 171–177. [52] M. Leist, B. Single, A.F. Castoldi, S. Kühnle, P. Nicotera, Journal of Experimental Medicine 185 (1997) 1481–1486. [53] H. Raza, A. John, S. Benedict, European Journal of Pharmacology 668 (2011) 15–24. [54] S. Orrenius, P. Nicotera, B. Zhivotovsky, Toxicological Sciences 218 (2011) p4352. [55] E. Jiménez-Medina, E. Berruguilla, I. Romero, I. Algarra, A. Collado, F. Garrido, A. Garcia-Lora, BMC Cancer 8 (2008) 78–88. [56] J.S. Bae, K.H. Jang, H. Yim, H.K. Jin, Cancer Letters 218 (2005) 43–52.