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University of California, Berkeley, CA 94720 and Children’s Hospital Oakland Research Institute, Oakland, CA 94609
3To whom correspondence should be addressed. E-mail: bames{at}chori.org.
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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KEY WORDS: • zinc • DNA damage • microarray • p53
The essential role of zinc has been shown in a wide range of cellular processes including electron transport, cell proliferation, reproduction, immune functions and defense against free radicals (1,2) as well as genetic stability and function (3,4). Approximately 25% of the zinc in rat liver is found in the cell nucleus and a large amount of zinc supplied in vitro is incorporated in the nuclei (5). Zinc content has a marked mechanistic impact on DNA as a component of chromatin structure, DNA replication and transcription and DNA repair (6). Zinc is a component of more than 1000 proteins, including electron transport proteins, DNA-binding proteins with zinc fingers, Cu/Zn superoxide dismutase (CuZnSOD) and several proteins involved in DNA repair such as p53, which is mutated in 50% of human tumors.
Deficits in certain essential vitamins and minerals can induce DNA damage by mechanisms similar to known environmental mutagens such as ionizing radiation (7–9). The role of zinc in DNA integrity and cancer has recently received increasing attention. Zinc status is compromised in cancer patients compared with healthy controls (10,11). Zinc deficiency causes oxidative DNA damage (12) and chromosome breaks have been reported in animals fed a zinc-deficient diet (13,14). In rats, dietary zinc deficiency causes an increase in esophageal tumor incidence compared with zinc-adequate controls (15) and zinc-deficient rats are more susceptible to tumor development when exposed to carcinogenic compounds (16–18). Epidemiological studies show that many adults and children may not be getting enough zinc in their diets. Zinc intake in 10% of the United States population is less than one-half of the recommended level (19), which could put them at greater risk for DNA damage and cancer.
We have previously shown in a rat glioma cell line that low zinc induces oxidative stress and increases DNA damage and p53 expression; however, the binding of the p53 transcription factor to DNA is markedly decreased (20). Thus, zinc deficiency impairs antioxidant defenses and compromises DNA repair mechanisms, making the cell highly susceptible to oxidative DNA damage. The current study examined the effects of zinc deficiency on DNA integrity in a human cell culture model and explored the potential mechanisms. The lung is an organ that is exposed to chronic oxidative conditions and could be sensitive to zinc depletion and cancer. Therefore, human lung fibroblasts, a primary cell line, was used in this study. Two distinct approaches to modulate intracellular zinc status were used: 1) chemically-induced Zn deficiency using the intracellular zinc chelator, N,N,N',N'-tetrakis-(2-pyridylmethyl)ethylenediamine (TPEN, a membrane permeable metal chelator with a high specificity for zinc) and 2) cell growth in zinc-deficient medium.
To understand cell response to zinc deficiency and the potential impact on DNA integrity, the overall gene expression changes with both TPEN and growth of cells in zinc-deficient medium were examined. Zinc modulates gene expression through both direct and indirect mechanisms. The use of microarrays allows the performance of large-scale gene profiling experiments revealing the overall response of the cell to zinc deficiency. Although microarrays give extensive information on gene expression changes, they offer no information regarding the actual physiological response of the cell. Therefore, to complement microarray results and to determine the physiological effects of zinc deficiency on DNA integrity, several functional tests of oxidative stress, DNA damage and DNA repair protein expression were also performed.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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IMR90 cells were obtained from American Tissue Culture Collection (Manassas, VA). Cells were grown in DMEM (Gibco Life Technology, Carlsbad, CA) and 10% CO2 at 37°C. For all experiments, a population doubling < 35 was used. Zinc-deficient media were prepared using a chelation method. Fetal bovine serum (FBS) was mixed at 4°C with 10% chelex-100 overnight. Mineral levels were monitored by inductively coupled plasma spectroscopy (ICP). IMR90 cells were seeded in 100-mm plates and grown in control medium (DMEM + 10% FBS), zinc-adequate media (ZnAD, DMEM + 10% chelex FBS + 4 µmol/L ZnCl2) or zinc-deficient medium (ZnDF, DMEM + 10% chelex FBS). Media were replaced every 2–3 d. For TPEN treatment cells were treated with 40 µmol/L of TPEN for 2 h. Cells were washed with PBS and harvested for further analysis. Cell viability was assessed using the methylthiazolydiphenyl-tetrazolium bromide assay as described by Hansen et al. (21). Cell counts were performed by use of a Beckman Coulter Z2 counter.
Mineral concentration.
Mineral concentrations of calcium, magnesium, iron, copper and zinc were determined using inductively coupled plasma-absorption emission spectrometry (ICP-AES) (Jarell-Ash Thermospec, Franklin, MA) with a slight modification of a previously reported method (22). Briefly, either 1 mL of medium or cell pellets (1 x 107 cells) were incubated with 1 mL of 40% ultrapure nitric acid (VWR, West Chester, VA) overnight. Following incubation, samples were diluted with deionized water to an 8% acid solution and analyzed by ICP-AES.
Microarray analysis.
The stress and aging array was purchased from Operon Technologies (Alameda, CA) and contained 70mer oligos from 704 known genes implicated in DNA repair, cell cycle, stress response, inflammation and micronutrient metabolism, printed in triplicate. Total RNA was purified from 2.0 x 107 cells using an RNeasy Miniprep kit (QIAGEN, Valencia, CA), and RNA quality and quantity were determined by electrophoresis and spectroscopy. Poly(A)+ RNA was subsequently isolated from total RNA using an Oligotex mRNA purification kit (QIAGEN). RNA labeling was performed using reverse transcription and aminoallyl coupling of RNA as previously described (23). Data were quantified using GenePix Pro 3.0 (Axon Instruments, Foster City, CA). After background subtraction, low intensity spots and ratios of intensity lower than twofold were eliminated from further analysis. TPEN-treated samples were RNA purified from IMR90 cells after incubations with 40 µmol/L of TPEN labeled with Cy5 dye for 2 h. The control sample was RNA purified at the same time from saline-treated IMR90 cells and labeled with Cy3 dye. For zinc deficiency, the treated sample was IMR90 cells cultured in zinc-deficient medium for 5 d and labeled with Cy5 dye, and the control sample was IMR90 cells cultured in zinc-adequate medium for 5 d and labeled with Cy3 dye. For both studies, slides were done in duplicate, with Cy dyes switched to control for labeling bias.
Assessment of DNA damage.
Single-strand breaks in cells were determined by alkali single-cell gel electrophoresis (Comet assay) as described by Singh (24). The assay is based on the alkaline lysis of labile DNA at sites of damage (i.e., from oxidation). Cells were suspended in 0.5% agarose and applied to microscope slides. Cells were subsequently lysed in comet assay lysis buffer (Trevigen, Gaithersburg, MD) for 1 h. After lysis, DNA was allowed to unwind in alkali buffer for 20 min; samples then underwent electrophoresis. Nuclear material was then stained with Sybr-green (Molecular Probes, Eugene, OR). Fifty cells from four independent samples were scored for tail migration intensity. Sample identity was not known until after scoring to reduce bias. To increase the sensitivity, a modified comet assay was performed using Fapy glycosylase (Fpg). To determine Fpg sensitive sites, an identical procedure was performed as described above followed by incubation of agarose-embedded cells with Fpg (Trevigen) at 37°C for 90 min. After incubation, cells were washed and incubated in alkali buffer for 20 min. Electrophoresis and staining were performed as previously described.
Assessment of oxidant production.
Oxidant production was monitored using the fluorescent probe dichorofluoroscein (DCFH) (Molecular Probes). DCFH (10 µmol/L) was added to the cells after 5 d in control, ZnAD or ZnDF media. Cells were incubated for 15 min at 37°C, then washed, trypsinized and immediately analyzed by the use of a fluorescence activated cell sorter (FACSort; Becton Dickinson, San Jose, CA).
Superoxide dismutase (SOD) Activity.
SOD activity was determined as described by Fridovich (25). Briefly, cells were lysed in 0.1% Triton-X, underwent two subsequent freeze/rethaw cycles and were stored on ice. SOD activity was determined from the percentage inhibition of the cytochrome c, xanthine-xanthine oxidase assay. The reduction of cytochrome c by superoxide generated from xanthine and xanthine oxidase was monitored by absorption at 418 nm.
Western blot analysis.
One million cells were harvested by trypsinization after 5 d in experimental medium and lysed in a 1.5% of SDS lysis buffer. Samples were mixed with Laemmli buffer and boiled for 5 min. SDS electrophoresis was carried out under standard conditions. Protein was transferred from the SDS gel to polyvinylidene fluoride membranes (Millipore, Bedford, MA) at 350 mA for 2 h. Blots were blocked in 50 g/L nonfat dry milk for 1 h at room temperature. Blots were then incubated with p53 antibody that recognizes both wildtype and mutant forms of p53 (1:1000 dilution; DO-1; Santa Cruz Biotechnology, Santa Cruz, CA). The blots were washed five times with phosphate-buffered saline + 0.2% Tween 20, and then incubated with horseradish peroxidase-conjugated goat anti-rabbit IgG at a dilution of 1:5000 for 1 h. Protein was detected by enhanced chemiluminescence using enhanced chemiluminesce reagents (NEN Life Science, Boston, MA). Radiographic films were scanned and band density was quantified with the public domain image processing and analysis program NIH Image (www.rsb.info.nih.gov/nih-image).
Statistics.
One-way ANOVA was performed to assess the differences among control, TPEN and Zn+TPEN groups or control, ZnAD and ZnDF groups. Differences in means among treatments were calculated by use of Tukey’s test. Differences were considered significant at P < 0.05.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Both methods had a significant influence on intracellular zinc status. Using ICP analysis, both short-term TPEN exposure and growth of cells in zinc-deficient medium for 5 d resulted in an 
50% drop in cellular zinc levels (Fig. 1). Other trace elements such as iron and copper were not affected. At these time points, cell viability and growth were not affected (data not shown).
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Although the gene expression changes were consistent with our hypothesis that low intracellular zinc status affects DNA integrity, little is known about the actual physiological response of the cell. The expression of p53 was also examined with both TPEN treatment and zinc deficiency in the culture medium. Western blot analysis revealed that both methods caused a significant induction of p53 protein (Fig. 2) in which upregulation may be in response to the DNA damage induced by either deficiency.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Microarray analysis showed changes in gene expression that account for the induction of oxidative DNA damage with the loss of zinc in human lung fibroblasts (Tables 1, and 2). A targeted stress and aging array was used to confirm a dysregulation of cellular processes leading to oxidative stress and DNA damage. In both models of zinc deficiency, gene expression profile changes were consistent with an increase in oxidative stress, DNA damage and a possible impairment of DNA repair. Although similar classes of genes were affected by both zinc-deficiency methods, only three genes were common to both treatments. These differences in response may be attributed to the fact that zinc depletion by TPEN was short term (2 h) and more drastic, whereas the medium deficiency method was completed over a 5-d period. Although cellular zinc levels were comparable between the two treatments, the response to the rapid zinc depletion with TPEN induced a different response. If cells were followed over a longer period of time, with a lower concentration of TPEN, more comparable changes in metallothionein and other variables may have been evident.
Similarities were evident in the classes of genes affected with both TPEN zinc depletion and cell growth in a zinc-deficient medium despite the differences resulting from acute and chronic exposures. Both up- and downregulation were seen in genes involved in DNA damage and repair. An upregulation of certain DNA damage and repair genes confirmed that both TPEN and zinc-deficient medium induced DNA damage. On the other hand, downregulation of DNA damage and repair genes suggested that some DNA repair mechanisms may be compromised (Tables 1, and 2). Some decreases in several proteins involved in both stress response and protein degradation were observed. The downregulation of these factors would alter the cells’ stress response and indirectly affect transcription of DNA damage/repair genes. Impairment in both stress response and proteolytic processing would also be directly detrimental to the cell.
Zinc deficiency also downregulated several mitochondrial electron transport chain proteins. Several studies have shown that an impairment of mitochondrial electron transport chain components can increase oxidant release and oxidative stress (32–35). Zinc is a component of many proteins in the mitochondrial transport chain and deficiency could result in the release of oxidants (36,37). Thus, mitochondrial disruption may account for the source of increased oxidative stress with zinc loss. It is clear that zinc deficiency had a deleterious effect on the DNA integrity of the cell, but the primary mechanism remains to be determined.
The expression profiles gave some clues in clarifying the mechanisms by which zinc deficiency increases oxidant levels and alters DNA integrity. However, gene expression changes need to be confirmed by alternative methods such as northern blot analysis or quantitative polymerase chain reaction. The goal of the current study was to identify gene expression changes in zinc-deficient cells accounting for increased DNA damage. The extent of the impairment to stress response and DNA repair mechanisms and quantification with alternative methods remains to be determined. Microarrays have been used previously to monitor gene expression profiles in zinc-deficient rat intestine and liver and have provided confirmatory data to the results in this study (38,39).
Approximately10% of the United States population ingests <50% of the RDA for zinc and are at risk for at least marginal zinc deficiency (19). Vegetarians and individuals consuming foods such as cereals and legumes containing zinc-binding phytates are at the highest risk for developing zinc deficiency. Through the use of genomics combined with functional assays, we observed that zinc deficiency induced oxidative stress and DNA damage, but at the same time compromised the cells’ ability to deal with this stress. A large portion of the population possibly being at risk for developing cancer as a result of zinc deficiency underscores the importance of proper nutrition in its prevention.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 This work was supported by National Foundation for Cancer Research Grant 00-CHORI, Wheeler Fund for the Biological Sciences at the University of California Berkeley, the Ellison Medical Foundation Grant SS-042–99, NIEHS Center Grant (BNA) and Training Grant NIH 5 T32 ES07075 (EH).
4 Abbreviations used: CuZnSOD, copper zinc superoxide dismutase; DCFH, dichorofluoroscein; FBS, fetal bovine serum; Fpg, Fapy glycosylase; ICP, inductively-coupled plasma; ICP-AES, inductively-coupled plasma absoption emission spectroscopy; SOD, superoxide dismutase; TPEN, N,N,N',N'-tetrakis-(2-pyridylmethyl)ethylenediamine; ZnAD, zinc-adequate; ZnDF, zinc-deficient.
Manuscript received 14 January 2003. Initial review completed 26 February 2003. Revision accepted 29 April 2003.
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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