Department of Pharmaceutical Biosciences, Division of Toxicology, Biomedical Center, Uppsala University, 75124 Uppsala, Sweden
Received June 12, 2003; accepted July 22, 2003
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ABSTRACT |
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Key Words: cadmium; zinc; apoptosis; DNA damage.
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INTRODUCTION |
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Cadmium produces oxidative modifications of DNA, such as the formation of 8-hydroxydeoxyguanosine, and the generation of strand breaks in different cell types, for example,. liver and kidney cells (Forrester et al., 2000; Littlefield and Hass, 1995
). Oxidative DNA damage produced by cadmium has been associated with an increased production of reactive oxygen species (ROS) (Ochi et al., 1987
), and interactions between this metal and DNA repair enzymes (Assmus et al., 2000; Waalkes, 2000
). Interestingly, there is evidence suggesting that Cd2+ binds covalently to N7 centers of adenine and guanine, and that it can form intrastrand bifunctional adenine-thymine (AT) adducts, suggesting a direct attack on the DNA molecule (Hossain and Huq, 2002
).
At the cellular level, cadmium also induces different biochemical changes, which are typically associated with apoptosis (Robertson and Orrenius, 2000). Apoptosis is a widespread and morphologically distinct process of cell death that plays an important role during normal embryonic development, for example, in modeling structures; regulating cell number; and eliminating abnormal, misplaced, nonfunctional, or harmful cells (Manova et al., 1998
). This process is particularly important in the development of the central nervous system, where neurons compete for a limited amount of survival factors (neurotrophic factors) and for the development and maintenance of the immune system (Collins et al., 1993; DSa-Eipper et al., 2000
). In human lymphoma cells, cadmium has been shown to cause apoptosis by two independent pathways: the Ca2+calpain and the caspasemitochondria pathways (Li et al., 2000
), indicating that apoptosis could play an important role in acute and chronic toxicity from this metal.
Zinc supplementation prior to cadmium administration prevents several of the effects observed when cadmium is added alone (Dreosti, 2001; Ferm and Carpenter, 1967). Thus it has been shown that zinc inhibits the apoptotic protease caspase-3 (Truong-Tran et al., 2001
), stabilizes the structure of p53 and DNA repair proteins (Chai et al., 1999
), acts as an antioxidant by decreasing ROS production in cell cultures (Dally and Hartwig, 1997
; Szuster-Ciesielska et al., 2000
), and prevents the gross teratogenic effects of cadmium by restoring normal development (Warner et al., 1984
).
One approach to investigating complex developmental processes and coordinating them with genetic regulation has been to disrupt morphogenesis with specific teratogens and study their consequences at the molecular and morphological levels. In this paper we have studied primary DNA damage by performing the alkaline version of the comet assay, a cadmium-induced alteration in the expression levels of genes by using the reverse transcription-polymerase chain reaction (RT-PCR), and pro-caspse-3 activation by western blot. The genes under study (p53, p21, Bcl-2, Bax, and Bad) were selected according to their implication in cell-cycle regulation and in the apoptotic pathway. In addition, the protective effects of zinc on cadmium-induced teratogenicity were studied in an experimental group of dams, which received a zinc pretreatment injection before cadmium exposure.
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MATERIALS AND METHODS |
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The animal experiments performed in this study were ethically approved by the Uppsala Djurförsökssetiska Nämnd, no. C 196/0 (as of September 29, 2000).
Cell viability.
Before the comet assay, cell viability was always evaluated, using the trypan blue exclusion assay. In brief, a 50-µl aliquot of single-cell suspension was collected from each treatment group and mixed with 50 µl of 0.25% trypan blue (Sigma St. Loius, MO). From this mixture a total volume of 10 µl was visualized in a Burker chamber under a microscope. Trypan blue-stained cells were considered nonviable.
Alkaline comet assay.
For the comet assay (Singh et al., 1988), 10 embryos from each treatment group were placed into phosphate buffered saline (PBS) and passed through a 0.4-µm diameter sieve in order to get a single-cell suspension. In the same way, one group of embryos was placed in freshly prepared 100-µM hydrogen peroxide at 37°C for 5 min, as a positive control. After centrifugation at 1100 rpm for 5 min, the cells were resuspended in 200 µl of PBS. Ten-µl cell suspension was added to 70 µl of 0.6% low-melting-point agarose in PBS, and 60 µl of this mixture was layered on top of a microscope slide precoated with 0.8% low-melting-point agarose in water. After the agarose had set, the slides were placed in a lysing solution (2.5-M NaCl, 100-mM Na2-ethylenediaminetetraacetic acid [Na2-EDTA], 10-mM Trizma base, 1% sodium lauryl sarcosinate, pH adjusted to 10 with NaOH, with 1% Triton X-100 and 10% dimethyl sulfoxide added before use) for 1 h at 4°C. From here on, all of the steps were performed in a cold room and under yellow light. The slides were then drained and placed in an electrophoresis unit (Sigma horizontal dual mode) containing an electrophoresis buffer (1-mM Na2-EDTA and 300-mM NaOH, pH > 13) for 40 min, to allow DNA unwinding. Thereafter, electrophoresis was performed for 5 min, using a field strength of 0.7 V/cm (300 mA, 25 V). After electrophoresis, the slides were neutralized with 0.4 M Trizma buffer (pH 7.5) for 15 min, dried at room temperature avoiding dust and particles, and kept in a sealed container until the day of image analysis (Vaghef et al., 1996
).
The slides were stained with ethidium bromide (20 µg/ml, 35 µl/slide) and examined at x400 magnification, using a fluorescence microscope (excitation filter 515560 nm, barrier filter 590 nm) attached to a black and white video camera connected to a computer-based image analysis system. Fifty comets per slide (12 to 16 slides per group of treatment) were randomly captured at a constant depth of the gel. Care was taken to avoid debris, comets without an identifiable nucleus, comets that were superimposed, and comets at the edges of the slides (Hellman et al., 1995). The image analysis program (Aequitas 1A, version 1.22, DDL Ltd, Cambridge, UK) with its special application for the comet assay (Autocell, version 2.0/9E, Reppalon AB, Sweden) was used for automatic analysis of the digitized images. Two different parameters were used as indicators of DNA damage: the tail moment and the percentage of DNA in the tail (Wiklund and Agurell, 2003
). All calculations were based on absolute intensities, and all slides were coded before examination.
Reverse transcription (RT).
The embryonic part anterior from the otic vesicle, without including branchial arches and developing heart, was immediately homogenized in TRIzol reagent (Invitrogen, Groningen, The Netherlands), and the total RNA was isolated according to the manufacturers instructions. One µg of total RNA isolated from the control and cadmium-treated embryos with and without zinc pretreatment were treated with 1-U DNase I Amp grade (Life Technologies, Taby, Sweden) according to the manufacturers instructions and subjected to RT to produce cDNA. The RT was performed in a total volume of 40 µl/sample containing 0.1-µM oligo(dT) primer (Amersham Pharmacia Biotech, Uppsala, Sweden), 1 µl of 5 x RT buffer (Promega, Madison, WI), 1-mM dNTP, and 200-U Moloney murine leukemia virus reverse transcriptase (Promega, Madison, WI). The reactions were performed at 37°C for 60 min and stopped by heating at 75°C for 10 min.
Polymerase chain reaction assay (PCR).
Five µl of the cDNA samples were amplified by PCR in a total volume of 50 µl containing 0.25-µM 5' and 3' primers, 10 x PCR buffer (50-mM KCl, 10 -M TrisHCl, 0.1% Triton X-100, and 2.5 mM MgCl2), 0.2-µM dNTPs, and 1 µl of AdvanTaqTM DNA polymerase (Clontech, Palo Alto, CA). The samples were incubated at 94°C for 4 min, amplified (94°C for 15 s, 58°C for 30 s, and 68°C for 30 s), and ended with a final extension at 68°C for 7 min. PCR primers and the number of cycles shown in Table 1 were used to detect transcripts from p53, p21, Bcl-2, Bax, Bad, and Cyclophillin (Cyc).
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Western analysis.
For detection of caspase-3 protein activation, a total protein extract was made from neural tubes and craniofacial structures anterior to the otic vesicle of the three samples under study (control, cadmium, and zinc plus cadmium treatment) in the presence of a lysis buffer (PBS containing 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, 1-mM Pefabloc, 10-µg/ml aprotinin, and 1-mM sodium orthovanadate). After centrifugation at 10,000g for 10 min, the protein concentration in each supernatant was determined according to standard methods (bicinchoninic acid protein assay; Pierce, Rockford, IL). The protein extracts (50 µg/lane) were resolved by SDSpolyacrylamide gel electrophoresis, followed by transfer to a polyvinylidene difluoride membrane (Hybond P; Amersham Pharmacia Biotech, Uppsala, Sweden). The membrane was blocked in 10% nonfat dry milk in Tris-buffered saline (TBS)/0.1% Tween-20 (TBST) for 1 h at room temperature and incubated overnight at 4°C with primary antibodies (ab) diluted in 5% nonfat dry milk in TBST. The primary ab used were goat polyclonal anti-caspase-3 at 1:2500 (Sigma, St. Louis, MO) and monoclonal anti-actin at 1:5000 (Sigma, St. Louis, MO). HRP-linked anti-mouse or anti-goat secondary antibodies (Amersham Pharmacia Biotech, Uppsala, Sweden) were used at 1:5000 for 1 h, and the membranes were washed twice with TBST and three times with TBS. Antigenantibody complexes were visualized by development with an enhanced chemiluminescence system (Amersham Pharmacia Biotech, Uppsala, Sweden) according to manufacturers instructions. Bands were quantified as described for the PCR gels.
Detection of programmed cell death (PCD).
The embryos dissected out from the control and treated animals were fixed in 4% paraformaldehyde (PF) in PBT (0.1% Tween-20 in PBS) overnight, processed into 100% methanol, and stored at -20°C until use. PCD detection was performed with minor changes following the manufacturers instructions using the in situ Cell Death Detection Kit, AP (Roche Diagnostics, Bromma, Sweden). Briefly, the embryos were permeabilized for 10 min in 10 µg/ml of proteinase K followed by 4% PF for 10 min before inactivation of the endogenous peroxidase with H2O2. Incubation of the embryos with 50 µl of TUNEL reaction mixture for 1 h at 37°C performed the labeling of the 3'-OH DNA ends. For the colorimetric detection of the apoptotic cells, an AP converter was used, and the embryos were "developed" at room temperature with NTB/BCIP until the desired intensity was achieved.
Statistics.
A chi-square test was used in Table 2 for comparing two proportions. A one-way ANOVA (analysis of variance) test was used to evaluate the differences between the control, cadmium, and zinc plus cadmium treatment, which were done in at least triplicate, and all contained a minimum of three experiments. Results were considered significant at p < 0.05 (*), p < 0.01 (**), and p < 0.001 (***).
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RESULTS |
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DNA Damage in Mouse Embryos
Cell viability, expressed as the number of cells not stained with trypan blue divided by the total number of cells counted, was constantly found to be >85% in all treatment groups (Fig. 2). Tail moment and content of DNA in the tail were used as indicators of DNA damage. Tail moment was defined as the distance between the center of mass of the tail and the center of mass of the head, in microns, multiplied by the percentage of DNA in the tail. It is, as compared to the tail length parameter, better to use since it takes into account both tail length and intensity (Bowden et al., 2003
).
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Activation of Caspase-3
Caspase-3 exists in all cells as an inactive pro-enzyme that requires cleavage at specific aspartate cleavage sites to yield active subunits of 17 and 12 kilodaltons (kD). The antibody used is directed against the N terminus of caspase-3, thus recognizing the intact pro-caspase-3 of 32 kD and the 17-kD cleaved fragment of caspase-3 by western blot. The levels of endogenous pro-caspase-3 were very similar between the treatments under study (Fig. 5). On the contrary, the 17-kD subunit was significantly increased, as compared to the controls in extracts prepared from the heads of cadmium-treated and in zinc plus cadmiumtreated embryos. Zinc pretreatment seemingly could significantly decrease the activation of pro-caspase-3 caused by cadmium.
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DISCUSSION |
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The most prominent malformation induced by cadmium in mammalian embryos is a failure of the anterior neural pore to close, leading to exencephalic embryos (Christley and Webster, 1983; Ferm, 1971
; Webster and Messerle, 1980
). In our study, 80 % of the embryos had an open neural tube 24 or 72 h after an injection of 4 mg/kg bw of cadmium. The anterior defects preferentially compromised the mid- and hindbrain structures, as was previously seen by Nakashima et al.(1988)
. Embryos were also growth-retarded when compared to the controls, as measured by CR length and somite number. Pretreatment with zinc (molar excess 2.7) before cadmium administration has been shown to have protective effects on cadmium teratogenicity (Ferm and Carpenter, 1968
). On the other hand, 8 mg/kg bw of zinc by itself did not have any obvious deleterious effects on embryonic development.
Programmed cell death (apoptosis) involves diverse input signaling pathways originating from the plasma membrane, mitochondria, the nucleus, or the cytoskeleton, and all these signals are critical for the involvement of apoptosis in normal brain morphogenesis (DSa-Eipper et al., 2001; Ferri and Kroemer, 2001). Twenty-four h after cadmium exposure, an increased apoptosis was observed in the mouse embryo (this paper and other, unpublished data; Christley and Webster, 1983
; Mirkes and Little, 2000
). Interestingly, several teratogens causing neural tube defects (NTDs) increased apoptosis in the tip of the neural folds (Mirkes and Little, 2000
). These are areas where physiological apoptosis also takes place (Sah et al., 1995
), being necessary for neural tube closure (Nonn et al., 2003
). There are some controversies as to whether this increased apoptosis is part of the mechanism by which neural folds do not elevate and fuse, or if it is just a parallel phenomenon for a number of teratogens (Weil et al., 1997
). In this paper we showed an increased apoptotic cells, detected by TUNEL, in cadmium-treated embryos as compared to the controls. On the contrary, these apoptotic areas were similar between control and zinc-pretreated embryos exhibiting normal embryogenesis.
As indicated in the present study, cadmium induced an upregulation in p53 expression levels, expected to stop cell cycle and allowing DNA repair mechanisms to act. p53 has been implicated in the regulation of both normal embryonic development and in the prevention of developmental defects after teratogenic exposure (Polyak et al., 1997; Sah et al., 1995
). Developmental abnormalities occur in mice with loss of p53 as well as with overexpression of p53, suggesting that p53 levels are critical for normal cellular processes (Lozano and Liu, 1998
). Specifically, p53- mice display defects in neural tube closure, resulting in an overgrowth of neural tissue in the region of the midbrain (exencephaly) (Sah et al., 1995
). In parallel, we also showed an increased primary DNA damage in cells that were exposed to cadmium. What still remains to be shown is the order of these events. (i) If cadmium, through a process of apoptosis, leads to the activation of endonucleases, the inactivation of DNA repair enzymes, and, as a consequence, the production of DNA strand breaks; or (ii) if the primary effect of cadmium is at the DNA level, arresting the cell cycle to allow repair of the DNA damage and secondarily activating apoptotic pathways due to the severe damage. Recent studies demonstrated that cadmium can produce DNA adducts interacting directly with adenine or guanine (Hossain and Huq, 2002
). Zinc administered 2 h before cadmium exposure was sufficient to maintain p53 expression levels and DNA damage as in the control embryos. Zinc has been shown to inhibit cadmium-induced apoptosis and the production of ROS in cell cultures (Szuster-Ciesielska et al., 2000
). It is also well known that zinc is an essential metal involved in zinc-finger proteins that are factors controlling cell proliferation, differentiation, and apoptosis through the regulation of gene expression (Urrutia, 1997
). The specific DNA-binding domain of p53 has a complex tertiary structure that is stabilized by zinc (Dreosti, 2001
). Cadmium competes and displaces zinc from its normal localization, thus altering zinc homeostasis and its physiological function (Hartwig, 2001
).
p53 directly interacts with the transcription of p21 that will in turn inactivate cyclinCdk complexes inhibiting the elongation step in DNA replication (Bunz et al., 1998). Here we show that p21 expression levels were also significally upregulated by cadmium. Besides, p53 regulates other apoptotic effector proteins interacting with members of the Bcl-2 family of proteins, including anti-apoptotic Bcl-2 and Bcl-XL, and pro-apoptotic Bax and Bad proteins (Sionov and Haupt, 1999
). The activity of some of these proteins can be regulated by phosphorylation and by the ratio of inhibitors to activators, since various family members can dimerize with one another, antagonizing or enhancing the function of the other (Gross et al., 1999
). Interactions between Bcl-2 and Bax regulate cytochrome c release from mitochondria and establish baseline sensitivity to apoptotic stimuli. In this study we observed a significant downregulation of Bcl-2 and a concurrent upregulation of Bax at the transcription level. It is thus likely that a decreased Bcl-2/Bax ratio promotes apoptosis signaling to be activated (Oltvai et al., 1993
). By western blot we also showed activation of pro-caspase-3, the executioner caspase, after cadmium administration.
The injection of zinc before cadmium treatment can keep the expression levels of the genes under study at basal levels. Zinc supplementation thus maintained the expression levels of p53 and p21, as well as the Bcl-2/Bax ratio. Pro-caspase-3 activation could not be kept at basal levels, but its activation was not as dramatic as was observed in the cadmium-treated embryos. Anyway, there was a significant decrease in pro-caspase-3 activation between zinc-pretreated and cadmium-treated embryos. It should be said that several different pathways leading to the activation of pro-caspse-3 could be involved, and only a few were taken into consideration in this study.
Protective effects by zinc may be related to metallothionein (MT) activation. It is well known that zinc administration induced MT expression levels in different tissues (liver, kidney, etc.) (Shimoda et al., 2003). A role of MT is to detoxify heavy metals, such as cadmium and mercury, and MT induction can prevent apoptosis induced by them (Klaassen et al., 1999
). In our study, MT induction by zinc prior to cadmium administration could account for the ameliorative effects observed.
In the present study we have clearly shown that cadmium can be a potent inducer of primary DNA damage in embryonic cells, and that it can activate several transcription factors implicated in the apoptotic pathway, such as p53, Bcl-2, and Bax. We have also shown that treatment with zinc could ameliorate the effects induced by cadmium, supporting previous data implicating that the mechanisms of cadmium teratogenicity are in some way related to zinc homeostasis.
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ACKNOWLEDGMENTS |
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NOTES |
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2 To whom correspondance should be addressed at Department of Pharmaceutical Biosciences, Division of Toxicology, Biomedical Center, Uppsala University, P.O. Box 594, SE-751 24 Uppsala, Sweden. Fax: +46-18-471-4253. E-mail: Lennart.Dencker{at}farmbio.uu.se.
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