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Carcinogenesis, Vol. 20, No. 11, 2101-2108, November 1999
© 1999 Oxford University Press

Molecular Epidemiology and Cancer Prevention

Tissue distribution of silibinin, the major active constituent of silymarin, in mice and its association with enhancement of phase II enzymes: implications in cancer chemoprevention

Jifu Zhao1 and Rajesh Agarwal1,2,3

1 Center for Cancer Causation and Prevention, AMC Cancer Research Center, Denver, CO 80214 and
2 University of Colorado Cancer Center, University of Colorado Health Sciences Center, Denver, CO 80262, USA


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 References
 
Polyphenolic antioxidants are being identified as cancer preventive agents. Recent studies in our laboratory have identified and defined the cancer preventive and anticarcinogenic potential of a polyphenolic flavonoid antioxidant, silymarin (isolated from milk thistle). More recent studies by us found that these effects of silymarin are due to the major active constituent, silibinin, present therein. Here, studies are done in mice to determine the distribution and conjugate formation of systemically administered silibinin in liver, lung, stomach, skin, prostate and pancreas. Additional studies were then performed to assess the effect of orally administered silibinin on phase II enzyme activity in liver, lung, stomach, skin and small bowel. For tissue distribution studies, SENCAR mice were starved for 24 h, orally fed with silibinin (50 mg/kg dose) and killed after 0.5, 1, 2, 3, 4 and 8 h. The desired tissues were collected, homogenized and parts of the homogenates were extracted with butanol:methanol followed by HPLC analysis. The column eluates were detected by UV followed by electrochemical detection. The remaining homogenates were digested with sulfatase and ß-glucuronidase followed by analysis and quantification. Peak levels of free silibinin were observed at 0.5 h after administration in liver, lung, stomach and pancreas, accounting for 8.8 ± 1.6, 4.3 ± 0.8, 123 ± 21 and 5.8 ± 1.1 (mean ± SD) µg silibinin/g tissue, respectively. In the case of skin and prostate, the peak levels of silibinin were 1.4 ± 0.5 and 2.5 ± 0.4, respectively, and were achieved 1 h after administration. With regard to sulfate and ß-glucuronidate conjugates of silibinin, other than lung and stomach showing peak levels at 0.5 h, all other tissues showed peak levels at 1 h after silibinin administration. The levels of both free and conjugated silibinin declined after 0.5 or 1 h in an exponential fashion with an elimination half-life (t1/2) of 57–127 min for free and 45–94 min for conjugated silibinin in different tissues. In the studies examining the effect of silibinin on phase II enzymes, oral feeding of silibinin at doses of 100 and 200 mg/kg/day showed a moderate to highly significant (P < 0.1–0.001, Student's t-test) increase in both glutathione S-transferase and quinone reductase activities in liver, lung, stomach, skin and small bowel in a dose- and time-dependent manner. Taken together, the results of the present study clearly demonstrate the bioavailability of and phase II enzyme induction by systemically administered silibinin in different tissues, including skin, where silymarin has been shown to be a strong cancer chemopreventive agent, and suggest further studies to assess the cancer preventive and anticarcinogenic effects of silibinin in different cancer models.

Abbreviations: CDNB, 1-chloro-2,4-dinitrobenzene; DCP-IP, 2,6-dichlorophenol-indophenol; EC, electrochemical; GST, glutathione S-transferase; OSA, 1-octanesulfonic acid; QR, quinone reductase; TEA, triethylamine; TPA, 12-O-tetradecanoylphorbol-13-acetate.


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 References
 
Silymarin, a polyphenolic flavonoid isolated from the seeds of milk thistle [Silybum marianum (L.) Gaertn] (1), is composed mainly of silibinin (or silybin, Figure 1Go) with small amounts of other silibinin stereoisomers, namely isosilybin, dihydrosilybin, silydianin and silychristin (2). Silymarin and silibinin are used clinically in Europe and Asia for the treatment of liver diseases (ref. 3 and references therein). In patients with liver disorders, treatment with silymarin or silibinin has been shown to improve liver function more rapidly than in those receiving placebo (4). Another multicenter trial showed that 420 mg daily administration of silymarin for several years resulted in a significant reduction in the mortality of patients suffering from alcoholic liver cirrhosis (5). The human population in Europe have used silymarin or silibinin as a liver tonic and current research indicates that it can be used in a whole range of liver and gall bladder conditions, including hepatitis and cirrhosis as well as dermatological conditions (6,7). In more recent years, silymarin has been marketed in the USA and Europe as a dietary supplement.



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  		Fig. 1. Chemical structure of silibinin.

 
In vivo pretreatment of experimental animals with silymarin or silibinin has been shown to protect against hepatotoxicity induced by a wide range of toxicants, including allylalcohol, carbon tetrachloride, galactosamine, phalloidin, thioacetamide and microcystin-LR (refs 138 and references therein). Other in vivo studies employing animal models and in vitro studies utilizing hepatocytes and liver microsomes have shown that both silymarin and silibinin afford significant protection against liver reduced glutathione depletion and lipid peroxidation induced by xenobiotic agents (9–12). Mechanistic studies in rodents and in cell culture have shown that silymarin is a strong antioxidant capable of scavenging free radicals (13–17). Besides, limited in vitro studies have shown that silymarin inhibits: (i) the formation of transformed rat tracheal epithelial cell colonies induced by exposure to benzo[a]pyrene (18); (ii) 12-O-tetradecanoylphorbol-13-acetate (TPA)-induced anchorage-independent growth of JB6 mouse epidermal cells (19); (iii) 7,12-dimethylbenz[a]anthracene-initiated and TPA-promoted mammary lesion formation in organ culture (20).

Based on the strong antioxidant activity of silymarin and the fact that silymarin and silibinin are already being used clinically as therapeutic agents, we started studies assessing the cancer chemopreventive and anticarcinogenic effects of silymarin in both long-term animal protocols and short-term cell culture models. Studies published in recent years from our laboratory showed that: (i) silymarin inhibits skin tumor promoter induction of ornithine decarboxylase activity and mRNA expression in mouse epidermis (21); (ii) topical application of silymarin protects significantly against UVB radiation-induced non-melanoma skin cancers in mice and that this effect of silymarin largely involves inhibition of UVB radiation-induced ornithine decarboxylase and cyclooxygenase activities, sunburn cell formation and skin edema (22); (iii) topical application of silymarin results in a highly significant to complete protection against skin tumor promoter-caused papilloma formation in mouse skin (23,24); (iv) silymarin inhibits the growth and proliferation of human cervical, breast and prostate carcinoma cells by inhibiting mitogenic signaling and inducing perturbations in cell cycle progression (25–27). More recently, studies from our laboratory have shown that silibinin, the major active constituent present in silymarin, has comparable (to silymarin) inhibitory effects towards human prostate, breast and cervical carcinoma cell growth, DNA synthesis and cell viability and is as strong an antioxidant as silymarin (N.Bhatia, J.Zhao and R.Agarwal, manuscript under review). Taken together, these studies suggested that both silymarin and silibinin have the potential to be developed as preventive and interventive agents against several human cancers. Despite a wide range of studies with both silymarin and silibinin and their clinical usage, the biodistribution and metabolism of silymarin and silibinin have not been studied in experimental animals. As a part of our systematic cancer chemopreventive and anticarcinogenic studies with silymarin and silibinin, in the present study we have assessed the tissue distribution and conjugate formation of systemically administered silibinin in mice. Additional studies were also performed to assess the effect of orally administered silibinin on the levels of phase II enzymes in different mouse tissues.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 References
 
Animal treatment with silibinin for tissue distribution studies
Six to seven-week-old male SENCAR mice were purchased from the National Cancer Institute (Frederick Cancer Research and Development Center, Frederick, MD). The animals were housed five per cage at 24 ± 2°C and 50 ± 10% relative humidity and subjected to a 12 h light/12 h dark cycle. They were acclimatized for 1 week before use in the present study and fed a Purina chow diet and water ad libitum. Prior to the study, the dorsal side of the skin was shaved using electric clippers to facilitate skin homogenization for silibinin extraction. The animals were starved for 24 h prior to dosing and were hydrated with 0.5 ml of tap water by intragastric intubation 1 h before administration of silibinin. Each time point had five mice. Silibinin, obtained commercially from Sigma Chemical Co. (St Louis, MO), was dissolved as 1.5 mg/ml silibinin in a water-based dosing solution containing 0.9% sodium chloride (w/v), 3% ethanol (v/v), 1% Tween 80 (v/v) and 6.6 mM sodium hydroxide. To make this solution, silibinin was first completely dissolved in the desired volume of ethanol and Tween 80 and then diluted with sodium chloride and sodium hydroxide solutions to achieve the final dosing solution detailed above. Silibinin (at a dose of 50 mg/kg body wt) was administered orally by feeding needle. A group of five mice was also similarly administered vehicle only and served as a negative control for tissue recovery studies. At different times after silibinin administration, mice were killed and liver, lung, stomach, skin, prostate and pancreas were removed. Each tissue sample was quickly snap frozen in liquid nitrogen and stored at –80°C until further analysis. Control mice were killed 1 h after the vehicle treatment and the desired tissues were removed and stored at –80°C.

Extraction of free and conjugated silibinin from different tissues
Each tissue sample obtained from different silibinin treatment time points was assessed for both free and conjugated forms of silibinin. Briefly, tissues (50–150 mg) were suspended in 3 vol of 50 mM Tris–HCl, pH 7.4, and homogenized thoroughly at room temperature using a Polytron PT-10 homogenizer (VWR Scientific, Plainfield, NJ). Each homogenate was divided into two parts. One part (80% of the total homogenate) was added to an equal volume of butanol:methanol (95:5 v/v) and mixed with the homogenizer for 5 s. The samples were centrifuged at 1500 r.p.m. at 4°C for 5 min and the organic layer collected. The butanol:methanol extraction was repeated twice more and the three butanol extracts were combined and stored at –80°C until HPLC analysis. The remaining tissue homogenate (20% of the total homogenate) was added to 100 µl of enzyme solution containing 500 U of ß-glucuronidase and 40 U of sulfatase (both from Sigma Chemical Co.) in 50 mM Tris–HCl, pH 7.4. The solution was incubated at 37°C for 1.5 h and then extracted with the same volume of butanol:methanol (95:5 v/v) three times. The three butanol extracts were combined and stored at –80°C until HPLC analysis.

HPLC analysis
The HPLC system (ESA, Bedford, MA) consisted of two ESA 580 model pumps, a UV detector, an electrochemical (EC) detector and ESA 5600 model HPLC control and analysis software. The UV detection wavelength was set at 270 nm and the potential for EC detection was 500 mV. An ESA C18 reversed phase analytical column (3 µm, 4.6x250 mm) was employed in all the HPLC analyses. The HPLC mobile phase contained solvent A [7.5% methanol in 100 mM acetate buffer containing 50 mM triethylamine (TEA) and 1 mM 1-octanesulfonic acid (OSA), pH 4.8] and solvent B (80% methanol in 100 mM acetate buffer containing 50 mM TEA and 1 mM OSA, pH 4.8). The linear gradient system employed at room temperature was: 0–5 min, 75% solvent A and 25% solvent B; 5–15 min, 75% solvent A and 25% solvent B to 50% of both solvents A and B; 15–20 min, 50% of solvents A and B to 30% solvent A and 70% solvent B; 20–25 min, isocratic 30% solvent A and 70% solvent B; 25 min, end of run. The solvent flow rate throughout the HPLC run was 0.6 ml/min and the column eluate was monitored by UV absorbance at 270 nm followed by EC detection.

Before the analysis and quantification of free and conjugated silibinin extracted from tissue samples, different concentrations of standard silibinin were analyzed by HPLC to find the quantitative linear range for both UV and EC detection. As needed, every tissue sample extract was adjusted to make sure that the amount of silibinin was within the linear range of detection. In each case, a 20 µl tissue extract was initially injected into the HPLC column and the silibinin peak was detected by the UV detector (270 nm) and further confirmed by the EC detector (500 mV). Silibinin quantification was based on peak area under the curve analysis and comparison with standard silibinin. The recovery of silibinin following extraction from different tissue homogenates was checked by adding a known amount of silibinin to each tissue homogenate and, following vigorous mixing, its extraction and quantification by HPLC as detailed above. In order to quantify the levels of conjugated silibinin in different tissue samples, the total silibinin levels obtained from enzyme-digested tissue homogenates were corrected for the levels of free silibinin. In all cases, therefore, the data shown for conjugated silibinin are after subtraction of free silibinin concentrations.

Animal treatment with silibinin for phase II enzyme studies
Six to seven-week-old female SENCAR mice were used in the study and housed in the animal facility as detailed above. The animals were acclimatized for 1 week before use in the present study and fed a Purina chow diet and water ad libitum. Several different studies in the past have used different doses, modes of administration and solutions/suspensions of silymarin or silibinin in rodent studies (refs 1, 3, 8–11 and references therein). These include doses from as low as 8 mg/kg/day for 10 days to as high as 1500 mg/kg once; administered i.p., intragastrically, p.o., i.v., etc., and dissolved in Tween 80/water, suspended in water, suspended in carboxymethyl cellulose, dissolved in saline or 0.1 N sodium hydroxide, etc. (1,3,8–11). Based on all these studies, for the phase II enzyme studies reported here we selected two doses of silibinin, 100 and 200 mg/kg body wt, and administered them by oral intubation dissolved in 0.2 ml of cottonseed oil. The animals in control groups were administered an equal amount of cottonseed oil only by oral intubation. Each time point and treatment group had five mice. The animal treatments with the above-mentioned doses of silibinin or cottonseed oil alone were done once in the morning every day and 24 h after 3, 7 and 15 days mice in each group were killed and liver, lung, stomach, skin and small bowel were removed and immediately placed in ice-cold 0.1 M phosphate buffer, pH 7.4. Tissues were cleaned properly, minced and homogenized in the same buffer and 100 000 g supernatant fractions were prepared as described earlier (28). Glutathione S-transferase (GST) activity was determined according to Habig et al. (29) using 1-chloro-2,4-dinitrobenzene (CDNB) as substrate. Quinone reductase (QR) activity was determined as described by Benson et al. (30) using 2,6-dichlorophenol-indophenol (DCP-IP) as electron acceptor. The statistical significance of differences in enzyme activities between cottonseed oil-treated controls and two doses of silibinin-treated experimental groups was analyzed using Student's t-test. Throughout the feeding protocol of silibinin in cottonseed oil or vehicle alone, animals were watched for any apparent signs of toxicity and food and water consumption. No evident change was observed in these mice throughout the experimental protocols (results not shown).


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 References
 
HPLC standardization and linear range of silibinin detection
First we standardized the HPLC conditions and both UV and EC detection sensitivity in the linear range. For these, different amounts of commercially obtained silibinin were dissolved in HPLC grade n-butanol and subjected to HPLC separation under different solvent gradient conditions. As shown in Figure 2AGo, under the HPLC method and solvent gradient system detailed in Materials and methods we were able to detect commercially obtained silibinin as a single peak by both 270 nm UV absorbance and by EC detection. In both cases, silibinin characteristically showed a retention time of ~13.5 min (Figure 2AGo). These HPLC profiles of silibinin also show its purity as 100%. Based on these HPLC profiles of silibinin, a linear detection range of silibinin under these conditions was also sought. As shown in Figure 2BGo, the standard curve for silibinin was linear in the range 10–100 ng in the case of UV detection (r = 0.9999) and 4–120 ng in the case of EC detection (r = 0.9999). Whereas the sensitivity of EC detection was much higher than that of UV detection, EC detection showed a high level of variation in detection limits, possibly because of a small change in room temperature or a change of solvent (data not shown), although OSA and TEA were used in the solvents to improve EC detection sensitivity and stability. Because of this, those tissue extract samples which were found to have a high concentration of silibinin were quantified using UV detection, whereas for samples with a low concentration of silibinin EC quantification was used.



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  		Fig. 2. HPLC profile of silibinin by UV and EC detection (A) and standard curves for silibinin following UV and EC detection (B). Different concentrations of silibinin dissolved in HPLC grade n-butanol (20 µl) were subjected to reverse phase analytic HPLC as detailed in Materials and methods. The column eluate was monitored at 270 nm UV absorbance followed by EC detection as described in Materials and methods. Based on the HPLC profiles of different concentrations of silibinin detected by both UV and EC, standard curves were derived using peak area as described in Materials and methods. In each case, a.u. stands for arbitrary units. The data points shown in (B) are means ± SD of four independent HPLC runs.

 
Tissue distribution of silibinin
After establishing the above HPLC conditions and linear detection ranges, we next analyzed tissue extract samples using appropriate volumes so that the silibinin levels fell in the linear range of detection. As shown in Figure 3Go, in representative HPLC profiles of tissue extract samples silibinin separated as a single sharp peak with an identical retention time to that of standard silibinin. In addition, several other endogenous substances also separated in these tissue extract samples, although they did not interfere with silibinin separation and quantification, suggesting that silibinin can be easily separated from other physiological constituents in all tissue samples (Figure 3Go). The recovery of silibinin extracted from six different tissues reported here, using butanol:methanol extraction, was 92 ± 5%.



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  		Fig. 3. HPLC analysis of skin and stomach tissue extracts and standard silibinin. Skin and stomach tissues were homogenized and extracted with butanol:methanol as described in Materials and methods. Butanol extracts (20 µl) and standard silibinin solution in butanol (20 µl) were subjected to reverse phase analytic HPLC and the column eluate was monitored at 270 nm UV absorbance as described in Materials and methods. In each case only representative HPLC profiles are shown.

 
The tissue distribution time kinetics of both free and conjugated silibinin clearly demonstrate that silibinin is quickly absorbed after oral administration and has a good bioavailability, at least in the tissues examined in the present study (Figures 4 and 5GoGo). The concentrations of both free and conjugated silibinin in the tissues examined reached their maximum levels within 1 h after silibinin administration. In the time–response curves the peak levels of free silibinin were evident at 30 min in the case of liver, lung, stomach and pancreas and at 60 min for skin and prostate. Other than liver, where free silibinin concentration decreased sharply after a peak at 30 min, in all other tissues it was retained for a much longer time, after which silibinin was eliminated from these tissues exponentially (Figure 4Go), with the elimination half-time (t1/2) values ranging from 57 to 127 min (Table IGo). As shown by the data in Table IGo, the maximum concentrations of free silibinin in these tissues were 8.8 ± 1.6, 4.3 ± 0.8, 123 ± 21, 1.4 ± 0.5, 2.5 ± 0.4 and 5.8 ± 1.1 (mean ± SD) µg silibinin/g tissue for liver, lung, stomach, skin, prostate and pancreas, respectively. When the time–response curves were analyzed for sulfate and ß-glucuronidate conjugates of silibinin, their maximum levels were evident at 30 min in the case of lung and stomach and 60 min in liver, skin, prostate and pancreas (Figure 5Go). After their peak levels, these conjugates of silibinin were eliminated from these tissues in an exponential manner (Figure 5Go), with the t1/2 values ranging from 45 to 94 min (Table IGo). The maximum tissue levels of sulfate and ß-glucuronidate conjugates of silibinin were found to be 5.7 ± 0.7, 2.8 ± 0.7, 270 ± 49, 4.3 ± 0.4, 6.1 ± 1.7 and 10.6 ± 1.5 µg/g tissue for liver, lung, stomach, skin, prostate and pancreas, respectively (Table IGo). The data summarized in Table IGo also show that in the case of liver and lung the levels of free silibinin were higher than conjugated silibinin and that the t1/2 values of free silibinin in these two tissues were shorter than that for conjugated silibinin. Conversely, the levels of conjugated silibinin were higher than free silibinin in stomach, skin, prostate and pancreas, but the t1/2 values of conjugated silibinin in these four tissues were shorter than that for free silibinin (Table IGo).



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  		Fig. 4. Tissue levels of free silibinin following its systemic administration to SENCAR mice. At the indicated time points after silibinin treatment at 50 mg/kg dose, animals were killed and the desired tissues were removed. Tissue homogenates were prepared, free silibinin was extracted using butanol:methanol and the tissue extracts were subjected to reversed phase HPLC as detailed in Materials and methods. The peak area for silibinin was analyzed for quantification. In each case, the data points shown are means ± SD of five independent mice; each sample was analyzed by HPLC twice.

 


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  		Fig. 5. Tissue levels of sulfate- and ß-glucuronidate-conjugated silibinin following its systemic administration to SENCAR mice. At the indicated time points after silibinin treatment at 50 mg/kg dose, animals were killed and the desired tissues were removed. Tissue homogenates were prepared, digested with sulfatase and ß-glucuronidase, silibinin was extracted using butanol:methanol and the extracts were subjected to reversed phase HPLC as described in Materials and methods. The peak area for silibinin was analyzed for quantification. In each case, the data points shown are means ± SD of five independent mice; each sample was analyzed by HPLC twice.

 

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  		Table I. Tissue distribution of silibinin (50 mg/kg dose) orally administered to SENCAR mice
 
Enhancement of phase II enzyme activities by silibinin
The phase II enzymes GST and QR play an important role in detoxification of carcinogens and their metabolites and, therefore, are important components of the tumorigenesis process (31,32). Several studies have shown that cancer chemopreventive agents enhance phase II enzyme activities as one of their mechanisms of action (33–36). Based on the cancer chemopreventive and anticarcinogenic effects of silymarin reported by us (21–27) and the results of the present study showing the tissue distribution of both free and conjugated silibinin following its systemic administration in mice, studies were also performed to assess the effect of orally administered silibinin on both GST and QR enzyme activities. As shown in Table IIGo, silibinin administration resulted in a moderate to highly significant increase in both GST and QR activities in all the tissues examined in both a dose- and time-dependent manner. In the case of GST activity, compared with vehicle-treated controls, treatment with 100 and 200 mg/kg doses of silibinin for 3, 7 and 15 days resulted in 8–110 and 10–140% increases (P < 0.1–0.001, Student's t-test) in enzyme activity. The observed increase in GST activity was maximum in small bowel at both the doses employed in the study and accounted for the 110 and 140% increases (P < 0.001, Student's t-test) over control for the 100 and 200 mg/kg doses, respectively, after 15 days of treatment (Table IIGo). In liver, lung, stomach and skin the increases in GST activity were comparable in a dose-dependent manner, specifically after 15 days of treatment (Table IIGo). Unlike GST activity, compared with vehicle-treated controls, the maximum increases in QR activity (58 and 74%, P < 0.001, Student's t-test) was observed in skin at both the 100 and 200 mg/kg dose of silibinin after 15 days treatment (Table IIGo). In general, the increases in QR activity in all the tissues examined, liver, lung, stomach, small bowel and skin, were not as profound as GST activity (Table IIGo). Furthermore, whereas the observed effects of silibinin on the enhancement of QR activity were both dose- and time-dependent, only a moderate increase in enzyme activity was evident following 3 and 7 days of silibinin treatment at both the 100 and 200 mg/kg doses (Table IIGo).


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  		Table II. Dose- and time-dependent increase in phase II enzyme activity in different tissues by oral administration of silibinin to SENCAR micea
 

   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
References
 
The major findings in the present study are: (i) standardization of the HPLC method and development of an HPLC system to extract, analyze and quantify the cancer chemopreventive agent silibinin in different biological samples; (ii) that systemic administration of silibinin in mice results in a rapid tissue distribution of both free and conjugated silibinin in the different tissues examined; (iii) that orally administered silibinin in mice results in a significant induction of the phase II enzymes GST and QR in different tissues. The tissue levels of and increase in phase II enzyme activity induced by silibinin were clearly evident in skin and prostate, where silymarin has been reported by us to be a strong cancer preventive and anticarcinogenic agent (22–24,27). These results convincingly suggest the presence of silibinin in the target organs to exert its cancer inhibitory effect and form the basis for more detailed studies in the future to evaluate the cancer preventive and interventive effects of silibinin, the major active constituent in silymarin, in experimental models of carcinogenesis representing different organ sites.

The use of reversed phase HPLC has been extensively employed in the separation of polyphenolic antioxidants similar to silibinin (37–39). Using this system, earlier studies from our laboratory and by others have separated and quantified different polyphenols present in green and black tea (37,38). More recently, reversed phase HPLC equipped with an EC detector has been used to identify and quantify tea catechins in human blood, urine, saliva and other tissues (39). Corroborative of these studies, in the present study, using an analytical HPLC system equipped with both UV and EC detectors, we were able to detect silibinin and quantify it in small (ng range) amounts in a linear range. The detection limit for silibinin by the HPLC system used here was comparable with that reported recently for silibinin quantification from human plasma (40). The observed tissue levels of both free and conjugated silibinin in liver, lung, stomach, skin, prostate and pancreas as early as 30 min following intragastric administration suggest that silibinin is rapidly absorbed from the stomach and shows a rapid tissue distribution. The rapid absorption and biodistribution of free and conjugated silibinin in different tissues in mice, as reported in this study, is consistent with human studies reported in recent years with silibinin (41,42). For example, the plasma concentrations of free and conjugated silibinin after the intake of a single oral dose of a lipophilic silibinin–phosphatidylcholine complex (silipide, 80 or 120 mg silibinin equivalent) were reported in both healthy human subjects and in patients with cholestasis (41,42). HPLC analysis of the plasma showed that absorption of silibinin from the gastrointestinal tract occurred rapidly, with the peak concentration of free silibinin in plasma at 2.4 h after dosing, followed by a decline with a t1/2 of ~2 h (41,42). In these studies, the peak concentrations of conjugated silibinin were greater than free silibinin and occurred at a later time of ~3.8 h. Comparing the results summarized in Table IGo with these published studies, it can be suggested that a similar trend exists for the concentrations of both free and conjugated silibinin in these tissues in mice as reported in human plasma studies. More studies, however, are needed in rodents to assess the absorption, tissue distribution, metabolism and excretion of systemically administered silibinin.

During the tumor initiation stage of multistage chemical carcinogenesis, chemical carcinogens are metabolized by the cytochrome P450 and epoxide hydrolase system to the ultimate carcinogenic metabolite that binds to DNA in the target cells, causing genetic alterations followed by tumor initiation (43,44). The collective action of phase II enzymes such as GST and QR is to afford protection against the adverse effects of reactive metabolites of procarcinogens (43,44). However, a decrease in phase II and antioxidant enzymes, following exposure to carcinogens and/or tumor promoters, reduces the protective ability of these enzymes against cell damage by carcinogens and/or their metabolites (43–46). Several cancer chemoprevention studies have shown that following administration of chemopreventive agents, the levels of phase II enzymes are elevated in various organs of the test animals (refs 33–36 and references therein). Consistent with these reports, in the present study we observed a moderate to statistically significant increase in the activity of the phase II enzymes GST and QR in liver, lung, stomach, skin and small bowel of mice orally administered silibinin. The dose- and time-dependent response of silibinin was also clearly evident in these studies. Together, these data suggest that in addition to several other cellular, biochemical and molecular mechanisms of action of silymarin reported by us and others which contribute to its cancer preventive and anticarcinogenic effects (21–27), an enhancement in phase II enzyme activity may also be associated with its cancer inhibitory potential. This suggestion is especially important for the study in which we found that topical application of silymarin for 7 days before tumor initiation with 7,12-dimethylbenz[a]anthracene results in a dose-dependent protection against tumor initiation in the SENCAR mouse skin initiation–promotion protocol (R.Agarwal et al., unpublished observation).

The multistage mouse skin carcinogenesis model of tumor initiation, promotion and progression has been extensively exploited over the last 40 years to define different stages of carcinogenesis and the mechanisms involved in them and to assess and identify cancer preventive agents (32). Utilizing this model, classical biochemical and DNA adduct formation studies have established the role of: cytochrome P450, epoxide hydrolase, phase II enzymes and DNA adduct formation in tumor initiation; induction of edema, hyperplasia, inflammation, cyclooxygenase, lipoxygenase, ornithine decarboxylase, a strong oxidative stress condition, etc. in tumor promotion; several additional genetic and epigenetic alterations and oxidative stress in tumor progression (32,43–46). However, studies in more recent years have focused on genetic alterations, signaling pathways, alterations in cell cycle regulatory molecules, tumor suppressor genes, transcription factors, etc. as the more defined molecular mechanisms associated with the tumor initiation, promotion and progression stages of carcinogenesis (32,47–49). Accordingly, cancer prevention studies identifying and defining the mechanistic aspects of preventive activity have included classical cellular and biochemical markers, as well as more detailed molecular events; a few examples are naturally occurring antioxidant green tea polyphenolics, genistein, curcumin, quercetin and apigenin (50–57). With regard to silymarin, studies over the last 6 years in our laboratory have focused on identifying its preventive and interventive effects in different models and defining the cellular, biochemical and molecular mechanisms of such effects. Based on our several published studies and ongoing work, it can be suggested that silymarin and silibinin have multifactorial mechanisms, including strong antioxidant activity, responsible for their cancer preventive and anticarcinogenic activities.


   Acknowledgments
 
This work was supported by USPHS grant CA64514 (to R.A.) from the National Cancer Institute, NIH.


   Notes
 
3 To whom correspondence should be addressed at: AMC Cancer Research Center, 1600 Pierce Street, Denver, CO 80214, USA Email: agarwalr{at}amc.org Back


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received March 30, 1999; revised May 24, 1999; accepted July 9, 1999.


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