Rolf Gebhardt*
Institute of Biochemistry, Medical Faculty, University of Leipzig, Johannisallee 30, 04103 Leipzig, Germany
ARTICLE INFO
Article history:
- Received 21 May 2008
- Received in revised form 11 December 2008
- Accepted 18 January 2009
- Available online 31 January 2009
Keywords:
- Antioxidants
- Apoptosis
- Cadmium
- Cytochrome C
- Hepatoprotection
- Plant tinctures
ABSTRACT
Cadmium is a heavy metal of considerable environmental concern that causes liver damage. This study examined the possible prevention of cadmium toxicity in human HepG2 cells and primary rat hepatocytes by Hepeel®, a combination preparation of tinctures from seven different plants. Hepeel® prevented cadmium chloride (CdCl₂)-induced cell death in both HepG2 cells and hepatocytes, and also reduced the loss of glutathione, lipid peroxidation, nuclear fragmentation, caspase activation and release of mitochondrial cytochrome C. To compare their relative efficacy, the seven constituent plant tinctures of Hepeel® were also separately tested. The tinctures China and Nux moschata, which exert solely anti‑oxidative effects, failed to reduce cytotoxicity, and only protected against loss of glutathione and lipid peroxidation. In contrast, the tinctures Carduus marianus and Chelidonium, demonstrated anti‑apoptotic effects, and protected HepG2 cells and primary hepatocytes against CdCl₂-induced cell death. These results demonstrate how the effectiveness of Hepeel® is determined by the synergistic features of its constituent tinctures. Furthermore, we conclude that cadmium toxicity in the liver is mainly due to stimulation of the intrinsic apoptotic pathway, but may be intensified by increased oxidative stress.
© 2009 Elsevier B.V. All rights reserved.
1. Introduction
Environmental exposure to fluctuating concentrations of heavy metals poses an enormous challenge for biological organisms. Toxic metals cause a vast array of adverse effects, including neurotoxicity, hepatotoxicity and carcinogenicity (Waalkes et al., 2000; Godt et al., 2006). Due to the global dispersion of heavy metals and their extensive use in modern society, some human exposure to toxic metals is inevitable. This ongoing prevalence of metal exposure necessitates protective measures at the environmental, social and individual level.
Cadmium is one of the most common toxic heavy metals, due to its primary accumulation in the liver and kidney (Godt et al., 2006). Cadmium causes hepatic, renal, skeletal, respiratory, and vascular disorders in humans (Nordberg, 1992; Waalkes et al., 2000), and it may also affect Leydig cells of the testes and hepatocytes and stellate cells of the liver (Koizumi et al., 1992; Dudley and Klaassen, 1984; Fariss, 1991; Souza et al., 2004a,b). Furthermore, cadmium is a potent carcinogen (Godt et al., 2006).
There is growing evidence that the oxidative stress (Sarkar et al., 1995) via reactive oxygen species (ROS) generation and mitochondrial damage are among the basic mechanisms of cadmium toxicity (Sarkar et al., 1995).
The combination preparation Hepeel® is frequently used to stimulate liver function and improve antioxidant function in acute and chronic diseases, such as cholangitis and cholecystitis (Gebhardt, 2003). Hepeel® also demonstrates several other protective features, such as induction of glutathione-S-transferase activity (Gebhardt, 2003). These findings prompted the present investigation of the hepatoprotective potential of Hepeel®, and its seven constituent plant tinctures, against cadmium-induced hepatocellular damage. To thoroughly examine this, and to provide comparative experimental data for two different cell types, we used the human hepatoblastoma cell line HepG2 and primary rat hepatocytes. Exposure to Hepeel® largely prevented cell death, and oxidative and apoptotic pathomechanisms were differentially affected by the constituent tinctures. The combined anti-oxidative and anti-apoptotic properties of Hepeel® and its constituent tinctures support its overall protective effect against cadmium-induced toxicity in liver cells.
2. Materials and methods
2.1 Materials
Hepeel® tinctures were prepared from seven different plants, according to procedures 3 and 4 of the German Homeopathic Pharmacopoeia (HAB, 2000), and were provided by the Biologische Heilmittel Heel GmbH (Baden-Baden, Germany). The following seven constituent tinctures were used: (1) Chelidonium majus, prepared from Chelidonium majus L. (Ch-B 007009, 10⁻² dilution), (2) Carduus marianus, prepared from Silybum marianum L. (Ch-B 007034, 10⁻² dilution), (3) Verratrum album L. (Ch-B 007050, 10⁻³ dilution), (4) Colocynthis, prepared from Citrullus colocynthis L. (Ch-B 007058, 10⁻³ dilution), (5) Lycopodium, prepared from Lycopodium clavatum L. (Ch-B 007001, 10⁻³ dilution), (6) Nux moschata, prepared from Myristica fragrans Houtt (Ch-B 007026, 10⁻³ dilution), and (7) China, prepared from Cinchona pubescens, Vahl (Ch-B 007018, 10⁻³ dilution). Hepeel® is a combination of all tinctures at the dilutions given above, with the addition of Phosphorus, a 10⁻³ dilution of yellow phosphorus. Hepeel® was supplied in sterile ampoules by Biologische Heilmittel Heel GmbH. The relative volume composition of 1:1 mL Hepeel® injection solution is: Chelidonium majus, 10⁻³ dilution) 1.1 μL; Carduus marianus (Silybum marianum, 10⁻³ dilution) 0.55 μL; Verratrum album, 10⁻² dilution) 2.2 μL; Colocynthis (Citrullus colocynthis, 10⁻³ dilution) 3.3 μL; Lycopodium clavatum, 10⁻² dilution) 1.1 μL; Nux moschata (Myristica fragrans, 10⁻³ dilution) 1.1 μL; China (Cinchona pubescens, 10⁻³ dilution) 1.1 μL and Phosphorus (white phosphorus 0515.41).
Dichlorodiphenyltrichloroethane (DDT) was purchased from Sigma (Daisenhofen, Germany). All other chemicals were from Roche Diagnostics (Mannheim, Germany), Merck (Darmstadt, Germany), Roth (Karlsruhe, Germany) or Sigma (Daisenhofen, Germany). Cell culture plates with tissue culture filter inserts were from Techno Plastic Products AG (Trasadingen, Switzerland).
2.2 Culture of HepG2 cells
HepG2 hepatoblastoma cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco, Eggenstein, Germany) supplemented with 2 mM glutamine, 10% fetal calf serum, 40 U/mL streptomycin and 50 U/mL penicillin, as previously described (Gebhardt, 1997). Cells were passed weekly, when confluent. Cell stocks (passage ≤31 till 40) were kept frozen in liquid nitrogen. Frozen cells were thawed, cultured for one week, and passed at least once before use. Confluent HepG2 cell cultures were used for all experiments.
2.3 Preparation and culture of rat hepatocytes
Sprague-Dawley rats were bred and maintained at the Medizinisch-Experimentelles Zentrum at the University of Leipzig, according to local ethical rules for animal care. They were kept on normal maintenance diet V1534 (Sniff, Soest, Germany) and tap water, ad libitum. Primary hepatocyte cultures were prepared by the livers of male rats (200–310 g) with collagenase perfusion, as previously described (Gebhardt, 1997). Cells were cultivated in Williams medium E (Lonza, Verviers, Belgium) on collagen-coated plastic plates, at a uniform cell density of 125,000 cells/cm². During the first 2 h, culture medium was supplemented with 10% fetal calf serum, and culture medium was used thereafter. The medium volume was maintained at 100 μL/cm² of plating area. Additional details of cell culture have been reported elsewhere (Gebhardt, 1997; Gebhardt et al., 1994). For toxicity experiments, incubation in various agents usually started 2 h after plating.
2.4 Induced toxicity with cadmium chloride
The nominal concentration of CdCl₂-induced cytotoxic effects was different for each cell type. For HepG2 cells, culture medium was supplemented with concentrations ranging from 3 to 8 μM. For primary rat hepatocytes, concentrations ranged from 2 to 6 μM. The highest CdCl₂ concentrations caused the greatest cell death in each cell type. In HepG2 cells, 8 μM CdCl₂ caused about 52% cell death, within 30 h of incubation, in hepatocytes, 6 μM CdCl₂ caused 72% cell death within 24 h of cultivation.
2.5 Preparation of Hepeel® tinctures
To prepare a working dilution of each tested compound, one part Hepeel® or tincture was mixed with 5 parts v/v of serum-free Williams Medium E, and gently shaken for 20 min at room temperature. This working solution of effective 0.1 dilution was used for further dilutions with Williams Medium E as specified in figure legends. Appropriate controls replaced each tincture or Hepeel® with equal volumes of ethanol.
2.6 Determination of cytotoxicity
Cytotoxicity of the tested compounds was determined using the colorimetric MTT-assay (MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide), as previously described (Gebhardt, 1997).
2.7 Determination of lipid peroxidation and ROS production
Malondialdehyde (MDA) measurements were used to quantify lipid peroxidation (Gebhardt, 1997). HepG2 cells or rat hepatocytes seeded on 60 mm petri dishes were incubated with or without CdCl₂ (3 or 4 μM) for 60 min after 30 h and 24 h of cultivation, respectively. In order to enhance oxidative stress, some plates were simultaneously exposed to t‑butyl hydroperoxide (t‑BHP; final concentration 1.5 mM). Thereafter, cells were washed with 0.9% NaCl, resuspended, and scraped into 1 mL of 50 mM potassium phosphate buffer (pH 7.4), then homogenised by sonication for 10 s (15 μs maximal sonpower, Sonipuls HD 2200, Bandelin electronic, Berlin, Germany). MDA was determined by thiobarbituric acid (TBA) assay (Esterbauer and Cheeseman, 1990; Gebhardt, 1997). The protein content of homogenates was measured following the procedure of Lowry et al. (1951).
Measurement of intracellular ROS was accomplished by using the DCFH assay (Wang and Joseph, 1999). HepG2 cells or rat hepatocytes cultivated on collagen-coated 95-well black flat bottom plates were washed 3 times with Krebs‑Ringer‑HEPES (KRH) solution pH 7.2 (Pavlica and Gebhardt, 2005). Cells were preloaded with 0.1 mM DCFH in either DMEM (HepG2 cells) or Williams Medium E (rat hepatocytes) for 30 min, then washed 3 times with KRH buffer. Cells were then treated simultaneously with CdCl₂ (3 μM) and the test compound diluted 1:10 with different starting dilutions (indicated in Table 1) for an additional 30 min. Fluorescence (485/520 nm, excitation 485/emission 520 nm, microplate reader, TECAN) was recorded for up to 30 min, while temperature was maintained at 37 °C. Percentage increase in fluorescence units/well was calculated by the formula: F₃₀/F₀ × 100, where F₃₀ = fluorescence at time 30 min, and F₀ = fluorescence at time 0 min (Pavlica and Gebhardt, 2005).
2.8 Determination of cellular glutathione content
To measure cellular glutathione (GSH) content, cells were cultured in 6-well plates for 30 h (HepG2 cells) or 24 h (primary rat hepatocytes). Test compounds were added 2 h after plating, along with the first change of medium. At the end of the incubation period, cells were washed and scraped into HEPES buffered isotonic medium as previously described (Pavlica and Gebhardt, 2005). Determination of GSH content was performed according to the method of Gebhardt and Faustel (1997).
2.9 Detection of apoptotic nuclei with DAPI
The blue nuclear dye DAPI (4′,6-Diamidino-2-phenylindole) was dissolved in methanol at 5 μg/mL and stored as stock solution. Cells were washed twice in potassium phosphate buffer (PBS) and fixed with ice-cold methanol. Thereafter, a working solution of DAPI (1 μg/mL) in methanol was added, and cell nuclei were stained for 15 min at 37 °C. Destaining was achieved by replacing methanol with pure methanol, followed by two rounds of washing with PBS.
2.10 Determination of caspase activity
Measurement of caspase-3 activity was based on the cleavage of a colorimetric substrate determined by the increase in absorbance at 405 nm. The assay was performed according to the instructions of the manufacturer (caspase-3 activity assay kit; Oncogene, Bad Soden, Germany) and adapted for HepG2 cells as described by Ochiai et al. (2004). Recombinant caspase-3 was used for assay calibration.
2.11 Preparation of cellular fractions and Western blot analysis
To measure cytochrome C release, cellular extracts were prepared by lysing cells in 10 mM Tris-buffer (pH 7.4) containing 2 mM EDTA, 1 μM pepstatin, 1 mM PMSF, leupeptin, 100 μM PMSF (phenylmethylsulfonyl fluoride), and 250 mM sucrose. Cells were homogenized by repeated passage through a 26-gauge needle, and were centrifuged at 14,000 × g for 10 min at 4 °C. Cytosolic supernatants and pellets containing mitochondria were collected and analyzed for spectral concentrations of mitochondrial protein, then used for Western blot analysis as previously described (Haupt et al., 2000). Cytochrome C was detected using a cytochrome C (Ab-8) antibody (sc-13156, Santa Cruz Biotechnology Inc., Heidelberg, Germany) followed by alkaline phosphatase-conjugated secondary antibody.
2.12 Statistical evaluation
Data were analysed for significance with a Student’s t-test for comparisons between two groups. Data are presented as mean ± standard deviation (SD) of three to four measures, except when stated otherwise.
3. Results
3.1 Cytotoxicity of cadmium chloride on hepatocellular populations
The cytotoxic effect of CdCl₂ on HepG2 cells was concentration- and time-dependent. Within the first 24 h of exposure, HepG2 cells tolerated up to 5 μM CdCl₂, but quickly died at higher concentrations (Fig. 1). At 7 μM CdCl₂, almost all cells were dead or had detached from the substrate. At 5 μM CdCl₂ or below, no visible alterations in cell morphology and nuclei were detectable after 24 h (data not shown). However, deterioration was seen at 5 μM CdCl₂ when cultivation was continued for another 6 h (Fig. 1). At that time, cadmium-induced cytotoxicity was already apparent at lower concentrations. The first signs of cytotoxic influence were detected above 2 μM, and almost all cells died at a concentration of 5 μM, as determined by MTT reduction to less than 10% of controls. The EC₅₀-value for CdCl₂-induced cytotoxicity in HepG2 cells was determined to be 5.9 μM after 24 h, and 2.8 μM after 30 h of cultivation.
Rat hepatocytes were even more sensitive to cadmium, and cytotoxicity was more prominent than in HepG2 cells, at all culture times. At 24 h after addition of CdCl₂, MTT reduction was already decreased in a concentration-dependent manner, above 2 μM doses (Fig. 2). At 6 μM, absorbance was reduced by approximately 70%. The EC₅₀-value for CdCl₂-induced toxicity was 3.7 μM. After 30 h, cell detachment in the MTT assay had further dropped, and were lower than those of HepG2 cells at all concentrations of cadmium (data not shown). Therefore, all subsequent measurements of cell viability were performed in HepG2 cells at 30 h of cultivation, and in rat hepatocytes at 24 h of cultivation.
3.2 Protection against cadmium cytotoxicity by Hepeel® and constituent tinctures
In the presence of Hepeel®, cadmium cytotoxicity was reduced in both cell types. In HepG2 cells at 30 h of culture, Hepeel® application resulted in the gradual increase of viability from 32% (control) to 53%, as dilutions changed from 10⁻³ to 10⁻¹ (Fig. 3A). At the 10⁻² dilution, there was significant enhancement of viability (P < 0.01).
Among the constituent tinctures, only Carduus marianus and Chelidonium, were effective in reducing cadmium cytotoxicity (Table 1). Within the range of 10⁻⁵ to 10⁻³ dilutions, Carduus marianus caused increased cell viability in a concentration-dependent manner, to values between 60 and 70% (Fig. 3B). Chelidonium application resulted in maximal values slightly above 60% (Fig. 3C). Also, the cell sensitivity was slightly higher with Carduus marianus, and significant differences were seen starting at the 2.5 × 10⁻⁴ dilution, whereas with Chelidonium significant differences were not apparent until the more concentrated dilutions of 10⁻⁴ and lower.
Similar results were obtained with rat hepatocytes after 24 h of cultivation. The 10⁻³ dilution of Hepeel® increased viability from 68% to almost 88%. At the same 10⁻³ dilution, Carduus marianus reached 95% and Chelidonium increased 87% viability (Table 1). As for HepG2 cells, the other constituents of Hepeel® did not reduce cytotoxicity (Table 1).
3.3 Cadmium-induced lipid peroxidation and ROS production
Exposure of HepG2 cells to CdCl₂ for 24 h did not change the rate of lipid peroxidation, as evidenced by the unchanged cellular production of malondialdehyde (MDA) compared to control measures (Table 2). However, when challenged with 1.5 mM t‑BHP, HepG2 cells exposed to 3 μM CdCl₂ responded with a 2.1-fold increase, and those exposed to 4 μM responded with a 2.3-fold increase of MDA, compared to control cells not exposed to cadmium.
Likewise, ROS production of HepG2 cells detected by DCFH fluorescence was stimulated by CdCl₂ only in the presence of t‑BHP (Table 2). The relative increase in ROS production was comparable to that for lipid peroxidation.
As shown in Table 3, Hepeel® significantly reduced t‑BHP-induced MDA production in both untreated HepG2 (control) cells and HepG2 cells exposed to CdCl₂ for 24 h. Among all tinctures, Carduus marianus was the most effective (Table 3). Dilutions were almost as broad.
Likewise, ROS production of HepG2 cells was reduced in a pattern similar to that of MDA measures: Hepeel® (31%), Carduus marianus (36%), China (18%), and Nux moschata (16%). All other tinctures were ineffective at reducing the CdCl₂-induced MDA production (data not shown).
The results for rat hepatocytes were different. In these cells, CdCl₂ led to an increase of MDA production of 55%, and an increase in ROS production of 32%, compared to control hepatocytes exposed to cadmium. However, as in HepG2 cells, the sensitivity to t-BHP in the presence of cadmium was also increased approximately 2-fold, from 155% to 302% for MDA, and from 132% to 273% for ROS. The following agents significantly counteracted the impact of CdCl₂, as apparent via the following reduction in MDA measures: Hepeel® (31%), Carduus marianus (36%), China (18%), and Nux moschata (16%). All other tinctures were ineffective at reducing the CdCl₂-induced MDA production (data not shown).
3.4 Cadmium-induced loss of GSH
A moderate drop in cellular GSH (19 ± 5%) was observed in HepG2 cells in response to exposure to CdCl₂ at a concentration of 3 μM (Table 4). This value is in accordance with an EC₅₀-value of approximately 4.5 μM. This loss was considerably enhanced (55 ± 4%) when cells were additionally exposed to t‑BHP. Only Hepeel® and the tinctures Carduus marianus, China and Nux moschata were able to significantly counteract the influence of CdCl₂, with or without additional t‑BHP (Table 4). When used alone, Hepeel® and Carduus marianus were able to completely restore cellular GSH content.
3.5 Cadmium-induced apoptosis
Cadmium toxicity via apoptosis was measured by DAPI-staining in two different ways; counting of fragmented nuclei and monitoring of cell death. Within 30 h of 3 or 4 μM CdCl₂ exposure, apoptotic fragmentation in HepG2 cell nuclei was apparent after DAPI staining, and total cell numbers were decreased (Fig. 4). Specifically, the percentage of apoptotic nuclei increased from less than 0.1% (controls) to about 8% in the presence of 3 μM CdCl₂ (Table 5). At earlier time points, such as 24 h, the proportion of fragmented nuclei was lower than at 30 h.
Addition of Hepeel® to the culture medium considerably reduced the apoptotic response at all concentrations of CdCl₂ in HepG2 cells and hepatocytes (Table 5). This influence was particularly pronounced in hepatocytes exposed to 4 μM CdCl₂, wherein the proportion of apoptotic nuclei was diminished from 42% to 4% (Table 5). A similar but less pronounced effect of Hepeel® could be observed in the presence of 5 μM CdCl₂ (cf. Fig. 5D).
Similar to the results seen in the MTT assays, the co-application of either Carduus marianus or Chelidonium with CdCl₂ effectively reduced the number of apoptotic nuclei, and enhanced cell survival (Fig. 4C and D). In the presence of 4 μM CdCl₂ and 10⁻⁴ final tincture dilutions, the proportion of fragmented nuclei in hepatocytes was 7% for Carduus marianus and 11% for Chelidonium (Table 5).
3.6 Cadmium-induced activation of caspases
Results for caspase-3 activity measurements were similar to those for apoptosis. In HepG2 cells, 3 μM CdCl₂ induced a significant increase in caspase-3 activity within 24 h (Table 6), with a 1.8-fold increase in caspase-3 and a 2.5-fold increase of caspase activity as measured by caspase-3/7 assay. Simultaneous addition of Carduus marianus at a 10⁻⁴ dilution to the culture medium resulted in a decrease of caspase-3 activity to about 1.3-fold, and the 10⁻³ dilution decreased caspase-3 activity to about 1.2-fold, relative to the vehicle-treated controls. Chelidonium was slightly less effective, but still reduced caspase-3 activity significantly in both assays (Table 6). A similar result was obtained for the Hepeel® 10⁻⁴ dilution, which reduced CdCl₂-induced caspase activity in both assays by approximately 40% (Table 6). Aside from Carduus marianus and Chelidonium, none of the other constituent tinctures was effective (not shown), since many HepG2 cells detached or decomposed completely within 30 h of CdCl₂ exposure.
3.7 Cadmium-induced release of cytochrome C
The release of cytochrome C from mitochondria of HepG2 cells was significantly higher in the presence of 3 μM CdCl₂ than in unexposed cells, which showed almost no release (Fig. 6). Densitometric analysis revealed a 27-fold increase in cytochrome C in CdCl₂-treated versus vehicle control cells. Hepeel® (10⁻¹) reduced the release of cytochrome C by about 5-fold, and Carduus marianus (10⁻³) reduced it by 7-fold (Fig. 6). Chelidonium was almost as effective as Carduus marianus, while treatment with China showed no effect (data not shown).
4. Discussion
Our results demonstrate a strong protective effect of the combination preparation Hepeel® and several of its constituent plant tinctures against cadmium-induced hepatocellular damage in both human hepatoblastoma cell line HepG2 and primary rat hepatocytes. We showed that cadmium-induced hepatocellular damage is effectively counteracted by these agents, thus gaining insight into potential mechanisms of this protective effect, which focus on two aspects: oxidative stress, and occurrence of apoptosis.
There are conflicting reports in the literature about oxidative stress during cadmium cytotoxicity. While some authors report that cadmium toxicity is due to, or at least associated with, increased oxidative stress and lipid peroxidation (Dudley and Klaassen, 1984; Fariss, 1991; RIkans and Yamano, 2000; Souza et al., 2004a,b; Koizumi et al., 2006), other authors could not detect enhanced lipid peroxidation in response to cadmium exposure in vivo and in vitro (Harvey and Klaassen, 1983; Aydin et al., 2003).
Concerning the occurrence of apoptosis in response to cadmium exposure our results are consistent with findings in mouse and rat liver (Habeebu et al., 1998; Pourahmad et al., 2001; Li and Lim, 2007) as well as human hepatocytes (Lasfer et al., 2008), and corroborates similar conclusions based on the observation of DNA laddering and other markers of apoptosis in response to cadmium exposure in HepG2 cells (Aydin et al., 2003; Oh and Lim, 2006).
Our results with DAPI staining also showed that treatment with Hepeel® and the single plant tinctures, which protected against cadmium toxicity, reduced the number of apoptotic nuclei. Furthermore, these agents also inhibited the activation of pre-apoptotic caspases and the release of mitochondrial cytochrome C. Therefore, these results strongly suggest that the most effective single tinctures, Carduus marianus and Chelidonium, are able to counteract intracellular processes other than oxidative stress, such as events leading to caspase activation and subsequent apoptosis, in response to cadmium. Silybin is an active substance in the Carduus marianus tincture, and is known to exert an anti-apoptotic influence in other systems (Singh and Agarwal, 2004; Pock et al., 2006). Thus, silybin may contribute to the protective effects of the tincture. However, direct anti-apoptotic properties of Chelidonium have not yet been described. Interestingly, alkaloids derived from Chelidonium such as chelerythrine and sanguinarine interact with the cytoskeleton (Slaninova et al., 2001), and of these, the alkaloid chelerythrine is an inhibitor of protein kinase C (Herbert et al., 1999). In addition, chelerythrine recently described as an inhibitor of BclXL function, which may help explain the pro-apoptotic effect observed with Chelidonium (Chan et al., 2003). In fact, detailed studies on the molecular interactions of chelerythrine revealed binding sites distinct for the Bcl3 (Bcl-2 homology 3) binding cleft (Zhang et al., 2006). This finding raises the possibility of alternate mechanisms favouring interactions of pro-survival members of the Bcl-2 family. In light of these findings, the concentrations of chelerythrine in our experiments is much lower than the EC₅₀ value for its pro-apoptotic effect (Chan et al., 2003; Maliková et al., 2006). Thus, at low concentrations anti-apoptotic influences of chelerythrine and sanguinarine seem to predominate.
Therefore, our results strongly suggest that the protective function of Hepeel® against cadmium-induced cytotoxicity results from the synergistic actions of its composite tinctures. The decisive anti-apoptotic influence of Hepeel® may be supported by its antioxidative features that help stabilize cellular GSH content, and consequently the sulfhydryl status of cellular proteins. Further studies are needed to discern whether this protective effect is specific to cadmium toxicity in hepatocytes, or can be generalised to other toxins and cell populations.
In conclusion, Hepeel® efficiently antagonised cytotoxic and apoptotic effects of the heavy metal cadmium in hepatocyte cell populations. This protective function is likely based on anti-apoptotic influence distinct from anti-oxidative function, but may be rendered more efficient by the synergistic effects of both. These observations add to the list of beneficial effects recently reported with this preparation (Gebhardt, 2003), and support the possible therapeutic use of Hepeel®, particularly for cases of heavy metal poisoning.
Conflict of interest
None.
Acknowledgement
This work was supported in part by the University of Leipzig (KST 764100100); and Biologische Heilmittel Heel GmbH, Baden-Baden, Germany (97000-050). The author would like to thank Mrs. D. Keller, Mr. F. Struck and Mrs. B. Woite for excellent technical assistance and Dr. A. Gerasimova for valuable comments and editing.
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