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Experimental Biology and Medicine 228:413-423 (2003)
© 2003 Society for Experimental Biology and Medicine

ORIGINAL RESEARCH ARTICLE

Humic Acid Induces Oxidative DNA Damage, Growth Retardation, and Apoptosis in Human Primary Fibroblasts

Mei-Ling Cheng*, Hung-Yao Ho*,{dagger}, Yi-Wen Huang*, Fung-Jou Lu§ and Daniel Tsun-Yee Chiu*,1

* Graduate Institute of Medical Biotechnology and School of Medical Technology, Chang Gung University, Kwei-san, Tao-yuan, Taiwan;
{dagger} Ronald O. Perelman Department of Dermatology, New York University School of Medicine, New York, New York 10016; and
§ Department of Biochemistry and Internal Medicine, College of Medicine, National Taiwan University, Taipei, Taiwan


   Abstract
Top
 Abstract
Introduction
 Methods
 Results
 Discussion
 References
 
Humic acid (HA) has been implicated as an etiological factor of Blackfoot disease endemic in the southwest coast of Taiwan. Dysfunction of endothelial cells and vasculopathy have been proposed to explain the onset of ulcerous changes at extremities. However, little is known about the effect of HA on activities of cells in these nonhealing wounds. In the present study, we demonstrate that HA adversely affects the growth properties of fibroblasts, one of the key players in wound repair. HA treatment caused growth arrest and apoptosis in human foreskin fibroblasts (HFF). This was accompanied by a significant increase in the level of 8-hydroxy-2'-deoxyguanosine (8-OHdG) in cellular DNA. The increased fluorescence in dichlorofluorescin (H2DCF)-stained and HA-treated cells suggests the involvement of reactive oxygen species (ROS) in HA-induced biological effects. Conversely, vitamin E pretreatment, which significantly reduced the 8-OHdG formation in HA-treated cells, alleviated the growth-inhibitory and apoptosis-inducing effects of HA. These results indicate that HA initiates oxidative damages to fibroblasts, and leads to their dwindling growth potential and survival. The present study suggests that HA-induced growth retardation and apoptosis of fibroblasts may play a role in the pathogenesis of Blackfoot disease.

Key Words: DNA damage • growth arrest • apoptosis • humic acid • 8-OHdG • ROS • vitamin E


   Introduction
Top
 Abstract
 Introduction
Methods
 Results
 Discussion
 References
 
Humic acid (HA) is a group of high-molecular-weight polymers that result from decomposition of organic matter, particularly dead plants. It exists abundantly in peat, soil, well water, and other sources (1). It consists of a mixture of closely related complex aromatic polymers, the exact composition of which varies with HA from different geographic locations. Chemical and infrared spectroscopic analyses revealed the presence of aromatic rings, phenolic hydroxyl, ketone carbonyl, quinone carbonyl, carboxyl, and alkoxyl groups in HA (2). A hypothetical "type" structure has been proposed: a polymer is composed of aromatic rings of the di- and trihydroxybenzene type that are bridged by ether, methylene, amine, imine, and other linkages; it also contains quinone groups and a variety of functional groups mentioned earlier (2–5).

HA was implicated as a causal factor of goiter (6–8), cancer (9), Blackfoot disease, an endemic peripheral vascular disease prevailing in the southwest coast of Taiwan (10–12), and Kashin-Beck disease, a chronic osteoarthritic disorder with necrosis of chondrocytes prevailing in mainland China (13). Epidemiologic and geochemical studies disclosed the presence of a high concentration of HA (approximately 200 ppm) in artesian well water from areas of endemic Blackfoot disease. The daily intake by an average resident in these regions was estimated to be as high as 400 mg (14, 15). Radioisotope tracing with iodinated HA in animals indicated that up to 60% of HA was retained in the body 24 hr after administration (16). It is believed that the intake of a huge amount of HA compromises the health of inhabitants, leading to the pathogenesis of Blackfoot disease.

Blackfoot disease begins with numbness or coldness, and can eventually proceed to black discoloration, ulceration, or gangrenous change in the extremities of patients. In pathology, Blackfoot disease is considered an arteriosclerotic disease (17, 18). Pathologic changes in blood vessels, such as mural thickening and accretion of subintimal plaque, account for an impaired peripheral circulation, ischemia, and the onset of ulceration in extremities. Thrombosis, embolism, superimposed infection and/or alterations in blood viscosity, and blood cell deformability may aggravate such pathologic changes (19, 20). Most studies have been focused on interactions between HA and endothelial cells, which normally protect the vessels from atherogenesis and aberrant thrombus formation (21, 22). However, this is not a complete picture for ulcer, which also reflects an inability to self-repair in an orderly manner. The normal course of wound repair involves a cascade of well-orchestrated events, starting with hemostasis, proceeding through a delimited inflammatory stage, granulation tissue formation, and re-epithelialization. followed by scar formation and remodeling (23). In the ulcer, the wound remains in the inflammatory stage, and the healing process does not set in.

Fibroblast is a critical player in the formation of granulation tissue. Apart from producing such extracellular matrix constituents as collagen, elastin, and glycosaminoglycans, fibroblasts also secrete a number of mitogenic and chemotactic cytokines for keratinocytes, endothelial cells, and themselves (23, 24). Additionally, they act as cellular stitch: fibroblasts can change into a less proliferative and more contractile form known as myofibroblasts (25); these cells express smooth muscle contractile proteins, such as {alpha}-smooth muscle actin, and can generate forces necessary for wound contraction (26, 27). Any dysfunction of fibroblasts may pose difficulty for a chronic ulcer to heal. It is plausible that HA may induce damage of fibroblasts, contributing to the persistence of ulcers in Blackfoot patients.

In this article, we show that HA caused growth arrest and apoptosis in human foreskin fibroblasts. The level of 8-hydroxy-2'-deoxyguanosine (8-OHdG), a de facto marker of oxidative damage to genomic DNA (28–30), was significantly elevated after HA treatment. Increased fluorescence was observed in dichlorofluorescin (H2DCF)-stained, HA-treated cells. Vitamin E, which greatly lessened the HA-induced damage to DNA, also protected the cells from growth inhibition and apoptosis-inducing effects of HA. These results demonstrate that HA acts via an oxidative mechanism. Taken together, the present study implies that HA-induced dysfunction of fibroblasts may play a role in the pathogenesis of Blackfoot disease.


   Methods
Top
 Abstract
 Introduction
 Methods
Results
 Discussion
 References
 
Chemicals.
Unless stated otherwise, all of the chemicals were obtained from Sigma (St. Louis, MO). Dulbecco’s modified Eagle’s medium (DMEM), fetal calf serum (FCS), penicillin, streptomycin, amphotericin, and trypan blue were purchased from GIBCO-Life Technology (Gaithersburg, MD). HA (sodium salt, technical grade) was obtained from Aldrich Chemical (Milwaukee, WI). Dichlorofluorescin diacetate (H2DCFDA) was available through Molecular Probes (Eugene, OR). Antibodies to Rb (14001A), p21Cip1 (C24420), and p27Kip1 (K25020) were obtained from BD PharMingen (San Diego, CA); the anti-actin antibody (SC1615) was obtained from Santa Cruz Biotechnologies (Santa Cruz, CA). 8-OHdG and 2'-deoxyguanosine (dG) were purchased from Sigma. Nuclease P1 and alkaline phosphatase were obtained from Roche (Mannheim, Germany).

Purification of Aldrich HA.
All of the experiments were performed with the same batch of HA purified from the one commercially available through Aldrich. Our purification method was similar to that described by Schnitzer (31) and is given below. The HA from Aldrich was first dissolved in 0.1 N NaOH solution, and any undissolved material was removed by centrifugation. The pH of supernatant was adjusted to 1.0 with 1 N HCl. Any precipitate formed was collected by centrifugation at 10,000g for 30 min, and was redissolved in 0.1 N NaOH. Such procedure of alkaline-acid treatment was repeated thrice. After the third round of acid precipitation, the precipitate was dissolved in 0.1 N NaOH, and pH of the resultant solution was adjusted to 7.2–7.4. It was then passed through the Sephadex G-25 column, and HA was eluted with distilled water. Elution profile was monitored by UV detector. Fractions corresponding to the first peak was collected, concentrated, and dried with a rotavapor. The HA was stored as dried powder, and was dissolved in phosphate-buffered saline (PBS) or water before experiments.

Cell Culture.
Primary human foreskin fibroblasts were isolated as described elsewhere (32, 33). They were cultured in DMEM supplemented with 10% FCS, 100 units/ml penicillin, 100 units/ml streptomycin, and 0.25 mg/ml amphotericin at 37°C in a humidified atmosphere of 5% CO2. The cells were fed every other day and were passaged at a ratio of 1:6 before the cultures were confluent. The cell number was determined by the trypan blue dye exclusion method.

Flow Cytometric Analysis of Cell Cycle.
Cells were rinsed with ice-cold PBS, trypsinized, and resuspended in 0.3 ml of PBS. They were then fixed and permeabilized by addition of 0.7 ml of ethanol. After a brief wash, the cells were gently resuspended in 1 ml of propidium iodide (PI) stain solution (40 µg/ml propidium iodide, 100 µg/ml RNase A, and 0.5% Triton X-100 in PBS), and were incubated at room temperature for 30 min before analysis on a FACSCAN flow cytometer (Becton Dickinson, Mountain View, CA). The sub-G1 fraction is considered as the apoptotic cells and taken as a measure of the extent of apoptosis (34). The sub-G1 phase was quantified using Modfit software (Becton Dickinson).

Terminal dUTP Nick End (TUNEL) Labeling.
TUNEL was performed with in situ cell death detection kit (Roche) according to the instructions of the manufacturer. In brief, the cells were trypsinized, washed with PBS, and resuspended in PBS. The cells (about 1 x 106 cells in 100 µl) were then fixed at room temperature for 1 hr by adding an equal volume of fixative solution (4% paraformaldehyde in PBS). The cells were washed, and then resuspended in permeabilization solution (0.1% Triton X-100 in 0.1% sodium citrate) at 4°C for 2 min. The sample was washed twice with PBS, resuspended in 50 µl of TUNEL label solution, and incubated at 37°C for 1 hr. Finally, the cells were washed, resuspended in 500 µl of PBS, and analyzed by flow cytometry. The TUNEL-positive fraction was quantified by CellQuest software (Becton Dickinson) (33).

Detection of Reactive Oxygen Species (ROS) by H2DCF Staining.
Formation of ROS in cells was detected by monitoring the fluorescence of dichlorofluorescein (DCF), the oxidation product of H2DCF, or that of rhodamine, the oxidation product of dihydrorhodamine 123 (DHR 123) (35, 36). The cells were loaded with 10 µM dichlorofluorescin diacetate (H2DCFDA) or DHR 123 for 5 min at 37°C. After treatment, they were immediately refed with fresh medium and observed under fluorescence microscope.

Ratio of 8-OHdG to dG.
DNA was extracted from untreated or treated cells as described elsewhere (37). The DNA was dissolved in 200 µl of 20 mM sodium acetate (pH 5.2), digested to nucleotide level with 20 units of nuclease P1 at 37°C for 2 hr, and followed by treatment with 6 units of alkaline phosphatase in 20 mM Tris buffer (pH 8.5) at 37°C for 1.5 hr. The hydrolysate was filtered through microcon YM-10 (Millipore, Bedford, MA) before HPLC analysis to remove enzymes and other macromolecules. Nucleosides in the filtrate were separated by a reverse-phase high-performance liquid chromatography (HPLC) system (ESA, Inc., Chelmsford, MA) equipped with a C8 column (3 µm, 4.6 x 150 mm, YMC-BD), and eluted at a flow rate of 1.0 ml/min with the mobile phase of 5% methanol in 100 mM sodium acetate buffer (pH 5.2) (38). The 8-OHdG and dG were monitored with an ESA Coulochem II electrochemical detector; the high sensitivity analytical cell 5010 were set at 150 mV for electrode 1, 300 mV for electrode 2 (with a full range deflection of 100 nA), 700 mV for electrode 3, and 800 mV for electrode 4 (with a full range deflection of 100 µA), respectively. The amounts of 8-OHdG and dG were quantified with respect to standards. The 8-OHdG levels are expressed as the number of 8-OHdG molecules per 106 dG.

Western Blotting.
The cells were rinsed with cold PBS, scraped, and collected by centrifugation. They were immediately lysed in lysis buffer (20 mM Tris/HCl, pH 8, 1% Triton X-100, 137 mM NaCl, 1.5 mM MgCl2, 10% glycerol, 1 mM EGTA, 50 mM NaF, 1 mM Na3VO4, 10 mM ß-glycerophosphate, 1 mM phenylmethylsulfonyl fluoride [PMSF], 1 µg/ml leupeptin, and 1 µg/ml aprotinin). Protein concentration of the lysate was determined by the Bradford method. The sample was analyzed by SDS-PAGE and immunoblotting with antibodies to Rb, p21Cip1, p27Kip1, and actin according to manufacturers’ instructions.

Statistical Analysis.
Student’s t test was used to determine the statistical significance between treatment groups in each experiment. Differences were considered statistically significant at a value of P < 0.05.


   Results
Top
 Abstract
 Introduction
 Methods
 Results
Discussion
 References
 
Growth Inhibition by HA.
To determine the cellular effect of HA on fibroblasts, we treated human foreskin fibroblasts (HFF) with increasing HA concentrations, and examined their growth kinetics. The untreated control displayed an exponential growth pattern. On the contrary, the growth rates of HFF cells incubated with HA for only 36 hr were significantly reduced (Fig. 1AGo). Such effect was conspicuous at the HA concentration as low as 10 µg/ml, and became more prominent at higher doses of HA; the cell number in cultures treated with 100 µg/ml HA for 36 hr was about 50% lower than that of control (Fig. 1BGo). Prolonged HA treatment (for up to 72 hr) resulted in even greater growth suppressive effect (Fig. 1AGo).




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  		Figure 1. Effect of HA on growth kinetics of HFF. (A) About 2 x 105 cells were seeded overnight, and were then treated with 0, 10, 25, 50, or 100 µg/ml HA. Cell number was determined 36, 72, and 120 hr after addition of HA. (B) Growth inhibitory effect of HA on HFF cells showed dose dependence. About 2 x 105 cells were treated with the indicated concentrations of HA. The cell number was determined 36, 72, and 120 hr afterward, and is expressed relative to that of untreated control. Results are presented as mean ± SD of five determinations. (C) Expression of cell cycle-related proteins, Rb, p21Cip1 and p27Kip1, in HA-treated HFF cells. Cells were treated with indicated concentrations of HA, and harvested 36 hr later. Cell lysates were separated by SDS-PAGE and analyzed with indicated antibodies by Western blotting. It should be noted that the anti-Rb antibody recognizes both the hypo- (pRb) and hyperphosphorylated (ppRb) forms. The results shown here are representative of at least three experiments.

 
The HA-induced growth stagnation in HFF cells was characterized by the cell cycle arrest at G1 phase. After 36 hr of treatment with HA (from 10 to 100 µg/ml), the cell cycle distribution showed an accumulation of G1 cells in a dose-dependent manner (Table IGo); at the highest concentration used, HA caused a notable increase in G1 cells (86.31% ± 0.41% for the 100 µg/ml group vs 54.63% ± 2.56% for the untreated control), and the corresponding decreases in S phase cells (5.96% ± 0.83% for the 100 µg/ml group vs 28.18% ± 0.63% for the untreated control) and G2/M phase cells (7.73% ± 0.42% for the 100 µg/ml group vs 17.22% ± 1.93% for the untreated control). This was well correlated with the dwindling proliferation rate of HA-treated HFF cells.


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  		Table I. Effect of HA and Vitamin E on Cell Cycle of HFF Cells
 
G1 progression is controlled through activity of retinoblastoma (Rb) protein, which exists in hypo- and hyperphosphorylated forms. The active, hypophosphorylated, but not the hyperphosphorylated, form binds to and inhibits E2F transcription factors necessary for G1 to S transition (39, 40). Upon HA treatment, the level of hyperphosphorylated Rb decreased in dose-dependent manner (Fig. 1CGo). Rb is phosphorylated by cyclin-dependent kinase activity, which is in turn regulated in part by Cdk inhibitors such as p21Cip1 and p27Kip1 (41–45). Expression of both proteins was elevated in HA-treated HFF cells when compared with the untreated control (Fig. 1CGo). Such results are consistent with the growth kinetics of HA-treated cells.

Prolonged HA Treatment-Induced Apoptosis.
Intriguingly, growth curves revealed significant decreases in cell number when the HA treatment was extended from 72 to 120 hr (Fig. 1AGo). Such reduction in cell number showed dose dependence, and was due to an increased level of cell death. The dying cells exhibited convoluted outline and membrane blebbings, both of which are typical of apoptotic cells (data not shown).

The extent of apoptosis was quantified by the following approaches. TUNEL labeling of the free DNA ends formed by fragmentation of genomic DNA, followed by flow cytometric analysis, gives a reliable means for quantification of apoptosis (46, 47). Prolonged HA treatment caused an increase in mean relative fluorescence, indicating incorporation of fluorescein-dUTP onto ends of nuclear DNA fragments in apoptotic cells (Fig. 2AGo). Apoptosis was barely detectable in cells treated for 36 hr at all the HA concentrations; it became significant in the experimental groups receiving 72 hr of treatment with 10 µg/ml HA (Fig. 2BGo). Additional increases in incubation time and/or HA concentration drastically elevated the percentage of TUNEL-positive cells: 120 hr of treatment caused significant level of apoptosis at HA concentrations ranging from 10 to 100 µg/ml.




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  		Figure 2. Quantification of the extent of apoptosis in HFF cells by TUNEL assay. (A) HFF cells were treated with varying concentrations of HA, and the level of apoptosis was determined 120 hr later by TUNEL assay and flow cytometric analysis. Representative histograms of HFF cells treated with 0 and 50 µg/ml of HA are shown here. (B) The extent of apoptosis with varying HA concentrations for HFF cells is shown here. Results are expressed as mean ± SD from three experiments. #P < 0.05; *P < 0.005 vs. the untreated control.

 
The appearance of cells with a low DNA stainability (sub-G1 fraction) in cell population has been considered as a marker of apoptosis (48). The sub-G1 fraction varies with treatment time and HA concentration in a pattern similar to that of TUNEL-positive cells. Over 40% of cells underwent apoptosis in response to 120 hr of treatment with 100 µg/ml of HA, whereas only insignificant level of apoptosis was observed in cells receiving 36 hr of treatment at all the HA concentrations used.

Increased Oxidative Stress and 8-OHdG Formation in HA-Treated HFF Cells.
We previously demonstrated that HA induces oxidative damages to erythrocytes (20). To determine whether HA also elicited oxidative stress in nucleated cells, we loaded HFF cells with dichlorofluorescin, and examined them after 24 hr of HA treatment. H2DCF has been used in a number of studies to monitor overall oxidative stress in single cells (49–51). Once oxidized, the leuco form H2DCF is converted into fluorescent DCF. If an oxidative insult occurs, these dye-loaded cells will take on a greenish appearance under fluorescence microscope. As shown in Figure 3Go, increased fluorescence of DCF, and hence increased oxidative stress, was observed in HA-treated HFF cells, whereas only a very low level (close to the background in unstained cells) of fluorescence was detected in untreated control. Of note, the fluorescence level increased with HA concentration (Fig. 3Go, F–J). Experiments with another fluorogenic dye dihydrorhodamine 123 gave similar results (data not shown). Moreover, as compared with the untreated control (Fig. 3AGo), a number of HA-treated cells showed slender morphology with greatly extended, spike-like cellular processes (Fig. 3Go, D and E), which was indicative of a stressful intracellular milieu.



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  		Figure 3. H2DCF staining for reactive oxygen species in HA-treated HFF cells. HFF cells were treated with 0 µg/ml (A and F), 10 µg/ml (B and G), 25 µg/ml (C and H), 50 µg/ml (D and I), and 100 µg/ml (E and J) of HA for 24 hr. After a change of medium, the cells were loaded with 10 µM H2DCFDA for 5 min. The samples were examined under inverted (A–E) and fluorescence microscope (F–J). The representative photographs shown here are taken from one out of five experiments (original magnification: x200).

 
It is plausible that HA-induced oxidative stress causes severe damage to macromolecules, in particular DNA, leading to growth arrest and cell death. Therefore, we measured the extent of oxidative DNA damage in HFF cells treated in different ways. The level of 8-OHdG, a reliable marker for assessment of oxidative DNA damage, was detected using HPLC-electrochemical method (30, 52). Figure 4Go shows representative chromatograms of 8-OHdG and total deoxyguanosine in DNA from cells treated with or without 50 µg/ml HA. The magnitude of oxidative DNA damage induced by HA was considerable (Table IIGo). Treatment of cells with 100 µg/ml for 120 hr produced about 57 8-OHdG adducts per 106 dG molecules, an over 4.5-fold increase as compared with the untreated control. It was evident that 8-OHdG formation increased with the HA dosage and duration of treatment. The 8-OHdG/dG ratios for cells incubated with 50 and 100 µg/ml for 72 hr increased by nearly 2- and 3-fold, respectively; after an additional 48-hr treatment, the corresponding values were 3.15 and 4.68 times that of control.



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  		Figure 4. HPLC analysis of 8-OHdG and dG in DNA of HA-treated HFF cells. Chromatograms for 8-OHdG (A) and dG (D) standards are shown here. HFF cells were untreated (B and E) or treated with 50 µg/ml of HA for 72 hr (C and F). Cellular DNA was isolated and processed as described in the "Materials and Methods" section for quantification of 8-OHdG (B and C) and dG (E and F). The amounts of 8-OHdG and dG were calculated by measuring the area under the corresponding peaks in chromatograms. These representative chromatograms are taken from one out of three experiments.

 

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  		Table II. HA-Induced Formation of 8-OHdG in HFF Cells
 
Prevention of HA-Induced 8-OHdG Formation and Cellular Changes by Vitamin E.
Vitamin E, a potent lipid peroxyl radical scavenger, has been shown to reduce oxidant-induced nuclear damages (53). Accordingly, we examined the possibility of using vitamin E to prevent the deleterious effect of HA. Pretreatment of HFF cells with vitamin E greatly diminished the HA-induced 8-OHdG adduct formation (Table IIGo). Thirty minutes of incubation with 200 µg/ml vitamin E could reduce the 8-OHdG/dG ratios by more than 50% in experimental groups treated with 100 µg/ml HA for 72 and 120 hr.

Reduction in 8-OHdG/dG ratio was paralleled by abatement of the growth inhibitory effect of HA. As shown in Figure 5Go, the cell numbers in the vitamin E- and HA-treated groups were significantly higher than those in HA-treated groups. After 36 hr of incubation with 50 µg/ml HA, the cell number was about 54.65% ± 5.37% of the control without HA treatment; however, with vitamin E pretreatment, this increased to 85.40% ± 7.20% of the control (Fig. 5AGo). A similar protective effect of vitamin E was observed for 72 and 120 hr of treatment (Fig. 5Go, B and C). Consistent with this, vitamin E ameliorated the G1 arrest imposed by HA: the percentage of G1 cells was significantly lower in the group treated with vitamin E and 100 µg/ml HA than in the one with HA alone (Table IGo). More important, vitamin E abolished the HA-induced apoptosis (Fig. 6Go). At whatever concentration, HA failed to induce apoptosis in the presence of vitamin E. Apparently, vitamin E was so potent as to completely prevent apoptosis under the condition that usually led over 40% of cells to demise.



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  		Figure 5. Growth inhibitory effect of HA was alleviated by vitamin E. HFF cells were incubated with or without 200 µg/ml vitamin E for 30 min, and then treated with the indicated concentrations of HA for 36 hr (A), 72 hr (B), or 120 hr (C). The cell number in each treatment group is expressed relative to that of control without HA treatment, which is assigned 100%. The data are presented as mean ± SD of four separate experiments. *P < 0.005 vs. the corresponding group without vitamin E pretreatment.

 


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  		Figure 6. Vitamin E-protected HFF cells from HA-induced apoptosis. HFF cells were incubated with or without 200 µg/ml vitamin E for 30 min, and then treated with the indicated concentrations of HA for 36, 72, or 120 hr. The extent of apoptosis was subsequently measured by PI staining and flow cytometric analysis. The data are presented as mean ± SD from four separate experiments. #P < 0.05; *P < 0.005 vs. the corresponding group with vitamin E pretreatment.

 

   Discussion
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
References
 
Previous studies have implicated dysfunction of endothelial cells in the arteriosclerotic changes associated with Blackfoot disease (17, 21, 22). However, this does not fully explain the persistence of chronic ulcers at the extremities of patients. The pathogenesis of chronic ulcers is multifactorial (54). There has not been any study examining the effect of HA on human fibroblast, a crucial player in wound repair. Here, our results demonstrate that HA instigated growth arrest and apoptosis in fibroblasts. These changes were associated with increased oxidative stress, and could be prevented, to a large extent, by vitamin E treatment.

Apparently, there is an intimate relationship between HA and ROS. Being a polyphenolic compound, HA has been shown to generate ROS such as superoxide anion (55). It is believed that the di-, trihydroxylphenolic, and quinone structures in HA can give rise to stable benzosemiquinone radicals that may directly attack cellular components or indirectly do so through generation of secondary free radicals. In erythrocytes, it causes a depletion of glutathione and several antioxidant enzymes (20). Clinically, the levels of serum lipid peroxides are significantly higher in patients with Blackfoot disease (56). Our results demonstrate that addition of HA caused a dose-dependent increase in DCF fluorescence in H2DCF-loaded cells (Fig. 3Go). Moreover, HA significantly increased 8-OHdG adduct formation in cellular DNA (Table IIGo). These findings clearly show that HA generates oxidative stress and inflicts oxidative damages on human fibroblasts. It follows that preincubation with a chain-breaking antioxidant, vitamin E, was able to reduce the oxidative DNA damage (Table IIGo). Indeed, these data are consistent with previous findings that HA can induce lipid peroxidation in erythrocytes (20), chondrocytes (57), and endothelial cells (58). Change in cell morphology is known to be a consequence of oxidative stress. Oxidants can modulate cytoskeletal proteins, resulting in actin cytoskeleton reorganization and changes in cell shape (59). A number of HFF cells receiving short-term HA treatment assumed a slender appearance with spike-like processes, instead of a normal flat morphology (Fig. 3Go). The mechanism for such change remains obscure. The ROS may effect through the direct alteration of the redox state of cytoskeletal proteins such as actin and its regulators (59–61), or may indirectly do so via other signaling pathways (62–64). More important, such morphological change and the accompanying change in cell-matrix contact may represent major alteration in cell physiology, for instance, a switch from growth to quiescence (65).

Free radicals have been implicated in regulating cell growth and death (66). HFF cells exhibited a diminished capacity to grow upon exposure to HA (Fig. 1Go). They underwent cell cycle arrest in G1 phase (Table IGo). In agreement with this, cyclin-dependent kinase inhibitors, p21Cip1 and p27Kip1, the expression of which was associated with G1 arrest (41–45), accumulated in HA-treated HFF cells (Fig. 1CGo). This was accompanied by reduction in the level of Rb phosphorylation (Fig. 1CGo). As mentioned earlier, cell geometry is closely related to growth control (67, 68). The change in cell morphology, together with a reduction in the area of cell-matrix contact, correlated well with their diminished proliferation and/or survival. Long-term treatment of cells with HA resulted in apoptosis (Figs. 2 and 6GoGo). A similar finding on endothelial cells has been recently reported (69). Vitamin E significantly relieved HA-treated HFF cells of growth inhibition, and completely blocked their apoptosis (Figs. 5 and 6GoGo), suggesting the involvement of ROS in these processes. Our results have several implications. First, HA induces G1 growth arrest as a protective mechanism. Cell cycle checkpoints are believed to allow time for cells to repair oxidative damage to macromolecules, in particular DNA, before their re-entering the cell cycle and completing such events as DNA synthesis and mitosis. In this way, genome integrity is preserved (70). Second, HA elicits apoptosis as an alternative mechanism to deal with irreparably damaged cells. Persistent oxidative stress during long-term HA incubation may cause overloading of repair system. The level of damage then exceeds the threshold required to trigger apoptosis. Furthermore, the complete blockage of HA-induced cell death by vitamin E suggests the involvement of lipid peroxidation in initiation or execution phase of apoptosis (71).

The finding that HA induced growth cessation and apoptosis in fibroblasts sheds light into the pathogenesis of Blackfoot disease. Chronic ulcers result from complex interaction between various factors, for instance, arterial insufficiency, inflammation, and neuropathy (19, 72). Our previous studies have shown that HA causes injuries to endothelial cells (21, 22), and erythrocytes (20), possibly accounting for the reduced blood supply to the extremities of Blackfoot patients. However, the picture is far from complete. Chronic wounds also represent the inability of epithelial cells and fibroblasts to carry out healing process (23–27). Being a critical player in wound healing, dermal fibroblasts proliferate actively during formation of granulation tissue (23–27). The present study clearly demonstrates that HA inflicts serious oxidative damage on human dermal fibroblasts, and impairs their ability to grow. This probably explains, in part, the persistence of chronic ulcers in Blackfoot patients.

Increased oxidative DNA damage in HA-treated cells is intriguing in cancer biology. It has been known for a long time that the humic substances isolated from artesian well water in Blackfoot endemic region are mutagenic (73). There are high incidences of skin, liver, lung, and bladder cancer in the same region (9, 74). Elevated 8-OHdG adduct formation in HA-treated cells provides a connection between HA and carcinogenesis. 8-Hydroxy-2'-deoxyguanosine is mutagenic itself: depending on its conformation, 8-OHdG can pair with dC or dA, and if unrepaired properly, causes G:C to T:A mutation (75, 76). As a marker of oxidative DNA damage, increased 8-OHdG formation indicates an overt oxidative attack on DNA. In addition to 8-OHdG, other base oxidation products, adducts, and even DNA strand breaks may be formed (77–82). The strand break lesions frequently cause various kinds of chromosomal aberrations, such as translocation and deletion (83, 84). Interestingly, there exist increased frequencies of chromosomal aberrations in lymphocytes isolated from cancer patients who inhabit the Blackfoot endemic area (85). Our findings also suggest that HA may be an etiological factor of cancer, and that HA-induced oxidative damage can be an underlying mechanism of carcinogenesis in the Blackfoot endemic region.

The current study clearly demonstrates that HA induces oxidative stress in dermal fibroblasts, leading to growth arrest and apoptosis. This suggests that HA-induced dysfunction of fibroblasts is implicated in the pathogenesis of Blackfoot disease. Nonetheless, more studies should be conducted before we apply our findings to the more complex clinical situation. Intriguingly, that vitamin E can reduce HA-elicited oxidative raises the possibility of using antioxidants as therapeutic intervention of Blackfoot disease.


   Footnotes
 
This study was supported by the National Science Council Grants NSC 90-2320-B182-029 and NSC 89-2314-B182-069 and by Chang Gung Memorial Hospital Grants CMRP1048 and CMRP1024.

M.-L.C. and H.-Y.H. contributed equally to this paper.

1 To whom requests for reprints should be addressed at Graduate Institute of Medical Biotechnology and School of Medical Technology, Chang Gung University, 259 Wen-Hua 1st Road, Kwei-san, Tao-yuan, Taiwan. E-mail: dtychiu{at}mail.cgu.edu.tw Back


   References
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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Received for publication June 20, 2002. Accepted for publication December 11, 2002.





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