* Department of Pathology, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2561; and Department of Pathology, University of Washington, Seattle, Washington 98104-2499
Received May 7, 2004; accepted June 3, 2004
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ABSTRACT |
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Key Words: copper; demyelination; disulfiram; dithiocarbamate; isoprostanes; lipid peroxidation; myelin.
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INTRODUCTION |
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Previous studies have demonstrated that subchronic administration of either DEDC or disulfiram to rats can produce peripheral neurotoxicity characterized by segmental demyelination, splitting of myelin lamellae, and formation of fluid- and debris-filled vacuoles in the Schwann cell cytoplasm (Tonkin et al., 2000, 2003
). Accumulation of copper in the nervous system presents a possible mechanism for this neurotoxicity, because copper is potentially cytotoxic through its ability to facilitate the formation of reactive oxygen species (Bremner, 1998
). For example, in patients with the autosomal recessive disorder Wilson's disease, accumulation of copper in the brain and liver is associated with neurological lesions including cystic changes in brain stem nuclei, loss of myelin, and astrocytic hyperplasia and with hepatoxicity resulting in progressive liver failure (Strausak et al., 2001
). Similarly, intracellular transport of copper by DEDC and disulfiram has been demonstrated in vitro to increase cytotoxicity in primary rat astrocytes and rat thymocytes (Chen et al., 2000
; Orrenius et al., 1996
; Wilson and Trombetta, 1999
). In those studies, dithiocarbamate-mediated cell death could be ameliorated by preventing accumulation of intracellular copper, either by incubating the cells in copper-free media or by coadministration of a hydrophilic, non-cell permeable copper chelator.
Although accumulation of copper in the brain and spinal cord in animals exposed to DEDC or disulfiram has been documented in several reports, the effect of DEDC and disulfiram on copper levels in peripheral nerve has not been reported. Similarly, the effect of these dithiocarbamates on copper localization at the cellular and subcellular level is also unknown. Histochemical localization of copper in the brain of normal rats has supported copper being localized to glial cells, rather than neurons, which is consistent with the known ability of astroglia to accumulate and detoxify metals (Szerdahelyi and Kasa, 1986; Tiffany-Castiflioni and Qian, 2001
). Alterations in the cellular distribution of copper has been reported in the liver during copper intoxication, in Wilson's disease, and in the brain of Long-Evans cinnamon (LEC) rats, a rat model of Wilson's disease (Bremner, 1998
; Haywood et al., 1985
; Sato et al., 1994
). Some reports have suggested that copper accumulation following administration of DEDC localizes in particular brain regions, including the cortex, hippocampus, and brain stem (Iwata et al., 1970
; Lakomaa et al., 1982
), but the effect of DEDC exposure on cellular localization of copper in brain is not well understood. Additionally, unlike the distinct myelin lesions reported for disulfiram and DEDC in peripheral nerve, a well-defined and commonly accepted lesion within the central nervous system resulting from DEDC or disulfiram administration has not been identified.
If elevated levels of copper resulting from abnormal copper homeostasis in peripheral nerve and accompanying oxidative stress are responsible for demyelination, then conditions that enhance DEDC-mediated accumulation of copper are expected to also enhance oxidative stress and the severity of lesions observed. In this study an initial assessment of the role of copper accumulation and lipid peroxidation in DEDC and disulfiram-mediated peripheral neuropathy is presented. Rats were administered either DEDC or deionized water by ip osmotic pumps and fed either a normal diet or diet containing elevated copper. Following the exposures, lipid peroxidation products and the severity of lesions in peripheral nerve and brain were assessed using GC/MS and light microscopy, respectively. Brain, peripheral nerve and liver sections were also stained by silver autometallography to characterize copper localization in these tissues, and samples of brain, liver, kidney, sciatic nerve, and serum samples were analyzed for metals by atomic absorption spectroscopy (AAS) and inductively coupled plasma-atomic emission spectroscopy (ICPAES) analysis.
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MATERIALS AND METHODS |
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Animals and exposures. All exposures were conducted in accordance with the National Institutes of Health Guide for Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee. Male Sprague-Dawley rats obtained from Harlan Bioproducts (Indianapolis, IN) were caged in a room on a diurnal light cycle and given food and water ad libitum. Body weights were determined prior to surgery and then weekly during the course of the experiment. The average weight of the animals at the time of the initial surgery was 300 ± 5 g. For morphological assessments and tissue copper levels four exposure groups were used: two DEDC-exposed groups receiving 0.35 mmol/kg/day NaDEDC and two control groups. One of the control groups (n = 4) and one of the DEDC groups (n = 4) were fed normal rodent diet containing 13 ppm copper, while the other control (n = 5) and DEDC (n = 5) groups were fed rodent diet supplemented with 200 ppm copper sulfate.
Animals were exposed to DEDC via 2ML4 Alzet® osmotic pumps surgically implanted in the abdomen under deep anesthesia (90 mg/kg ketamine HCl and 7.5 mg/kg xylazine). After 4 weeks, the pumps were replaced by the same procedure to extend the exposure period to 8 weeks. DEDC was delivered as an aqueous solution of NaDEDC, the concentration of which was determined from the UV absorbance at 282 nm ( = 13,000 l/mol/cm) prior to filling the pumps. Control pumps contained deionized water. Solutions were sterilized prior to filling the pumps by filtering through a 0.22 µm-syringe filter. At the end of the exposure period neuromuscular function was assessed by measuring hind limb grip strength using a digital force gauge (Meyer et al., 1979
), and the animals were perfused for morphological examination of peripheral nerve, liver, and brain.
Identical exposure groups (n = 4 per group), exposure durations, and exposure levels to those described above were used for the animals analyzed for F2-isoprostanes and F4-neuroprostanes. At the end of the 8-week exposures the animals were deeply anesthetized, exsanguinated, and the sciatic nerves and brains immediately frozen in liquid nitrogen.
Preparation of tissue for morphological assessment. Tissue was prepared according to the method of Danscher with some modifications (Danscher, 1984). Animals were perfused through the left ventricle of the heart with 0.1% Na2S in 0.15 M sodium phosphate buffer (pH 7.4) for 2 to 3 min, then with 3% glutaraldehyde in 0.15 M phosphate buffer (pH 7.4) for 10 to 15 min, followed by the Na2S solution for 7 min. After perfusion, liver, brain, kidney, and sciatic nerve were dissected out and immersed in 3% glutaraldehyde for 1 h, then transferred to 0.15 M phosphate buffer. Sciatic nerve sections were post-fixed with osmium and embedded in Epon; thick sections were cut and stained with toluidine blue. Thick (1-mm) sections of sciatic nerve tissue were evaluated by light microscopy on an Olympus BX41 microscope equipped with an Olympus C-3040 Camedia digital camera. Brain and liver sections were embedded in paraffin and stained with hematoxylin and eosin (H&E) by the Vanderbilt Medical Center Neuropathology Laboratory and also evaluated by light microscopy. Sections of sciatic nerve, brain, and liver were embedded and stained by silver autometallography at NeuroScience Associates (Knoxville, TN) (Ross et al., 1996). Free floating MultiBrain sections were rinsed twice in Na acetate, and then placed into the acidic physical developer (pH 5) of Gallyas for several (46) min. Development was stopped by transferring the sections to 1% acetic acid. Following a 30-s rinse in 1% potassium thiosulfate the sections were rinsed in water and mounted. After air-drying the slide-mounted sections were lightly counterstained with neutral red.
Blood collection. Prior to perfusion, 34 ml of blood was collected from anesthetized animals via cardiac puncture and 0.51 ml used to isolate globin for analysis by HPLC. Blood was separated into plasma and hemolysate as described (Erve et al., 2000) and 100 µl of 1 N ascorbic acid added to each 1 ml hemolysate. The resulting solution was added dropwise to 10 ml of 2.5% oxalic acid in acetone. Globin was allowed to precipitate on ice for 30 min, then centrifuged at 10,000 x g for 10 min at 4°C and washed in 5 ml ice-cold acetone. The pellet, containing crude globin, was dried under a stream of N2 and stored at 80°C. Dried globin was solubilized with 0.1% trifluoroacetic acid (TFA) to produce a solution for HPLC analysis.
Globin chains were separated by RPHPLC on a Phenomenex Jupiter 5-µm column (150 x 460 mm) using a Waters 2690 liquid chromatograph after adjusting sample concentration to a UV absorption of 1.0 ± 0.2 at 280 nm. Globins were separated using a linear gradient from 56% solvent A, 44% solvent B, to 30% solvent A, 70% solvent B, over 30 min followed by a linear gradient to 100% solvent B over 10 min. Solvent A was 20:80:0.1 acetonitrile:water:TFA, and solvent B was 60:40:0.08 acetonitrile:water:TFA. The elution of globin peaks was monitored by their UV absorption at 220 nm using a Waters 996 photodiode array detector.
The remaining blood not used to isolate globin was placed into a 15-ml conical tube and allowed to clot, then centrifuged to precipitate the clot, and the serum removed. 200-µl serum aliquots were analyzed at the Vanderbilt University Hospital Clinical Chemistry laboratory (Nashville, TN) for aspartate aminotransferase activity (AST/SGOT). The remaining serum was stored at 20°C and analyzed for copper concentration.
Analysis of tissue metal levels. Brain, liver, and kidney sections were analyzed by both inductively coupled plasma-atomic emission spectroscopy (ICPAES) and atomic absorption spectroscopy (AAS); sciatic nerve sections were analyzed by ICPAES alone. Samples for AAS analysis were taken from individual animals and analyzed at Kord Animal Disease Diagnostic Laboratory (Nashville, TN). Due to the limited amount of sciatic nerve tissue available, all tissue samples for ICPAES were taken from individual animals, then pooled by treatment group in order to obtain enough tissue for analysis. Brain tissue samples for ICPAES were approximately 1-mm midsagittal slices. Kidney sections for ICPAES were approximately 1-mm cross-sections through the center of a single kidney; liver sections were 1- to 2-mm3 pieces of tissue; sciatic nerve sections were 10-mm lengths of a single nerve. Average weights of the pooled tissue samples for each treatment group of brain, kidney, liver, and sciatic nerve were 0.79 ± 0.05 g, 0.34 ± 0.07 g, 0.11 ± 0.01 g, and 0.059 ± 0.006 g, respectively. Each tissue sample was analyzed in duplicate for mineral content by ICPAES at the Diagnostic Center for Population and Animal Health at Michigan State University (East Lansing, MI). Elements determined in this analysis were barium, calcium, copper, iron, magnesium, manganese, molybdenum, phosphorus, zinc, chromium, cadmium, sodium, nickel, vanadium, sulfur, and potassium.
Analysis of serum copper levels. Serum copper levels were measured spectrophotometrically based on described methods (Tietz, 1976). Briefly, serum was mixed with 2 M HCl and incubated at room temperature for 10 min; then 20% trichloroacetic acid was added, and the solution was mixed and allowed to stand at room temperature for 10 min. Samples were centrifuged for 10 min; then the supernatant was removed and combined with a solution of sodium pyrophosphate, sodium citrate, and ammonium hydroxide, followed by 0.5% cuprizone in 50% ethanol. The resulting solution was mixed and incubated at room temperature 20 min; then the absorbance at 620 nm was determined and compared to a standard solution of copper sulfate.
Analysis of F2-isoprostanes and F4-neuroprostanes. Measurements were performed on individual animals using two sciatic nerves (F2-isoprostanes) and the right cerebral hemisphere (F2-isoprostanes and F4-neuroprostanes) for each animal. F2-isoprostanes and F4-neuroprostanes were quantified in Folch extracts of cerebrum or sciatic nerves using gas chromatography with negative ion chemical ionization mass spectrometry with selective ion monitoring using a published method (Reich et al., 2001). Briefly, samples were extracted by the method of Folch, a stable isotope internal standard added, and then prepared for gas chromatography (GC) through a series of TLC and SepPak separations. Gas chromatography was performed using a 15 m, 0.25 mm diameter, 0.25 mm film thickness, DB1701 fused silica capillary column. The column temperature was programmed from 200 to 300°C at 20°C/min. Methane was used as the carrier gas at a flow rate of 1 ml/min. Ion source temperature was 250°C, electron energy was 70 eV, and filament current was 0.25 mA. Negative ion chemical ionization (NICI) mass spectrometry (MS) was performed using an Agilent Technologies G1789A GC/MSD instrument with a Hewlett-Packard computer system with ChemStation (NT).
Statistical analysis. Analysis of variance (ANOVA) with Bonferroni's post-hoc test, the nonparametric Kruskal-Wallis test (for ranked scoring of lesion severity), and the Student's t-test were performed using GraphPad Prism software. Statistical significance was taken to be p < 0.05 unless otherwise noted.
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RESULTS |
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Analysis of Tissue Metal Levels
Copper levels in the brain, liver, and kidney determined by ICPAES and AAS are shown in Table 2. Consumption of the 200-ppm copper diet alone had no effect on copper tissue levels relative to the normal copper diet controls. Exposure to DEDC alone resulted in a significant increase in copper levels in brain (AAS and ICPAES), sciatic nerve (ICPAES), and liver (ICPAES) relative to controls. Exposure to DEDC and the 200-ppm copper diet also resulted in elevated copper levels relative to controls in brain (AAS and ICPAES), liver (AAS and ICPAES), and sciatic nerve (ICPAES). The level in liver (AAS and ICPAES), brain (ICPAES), and sciatic nerve (ICPAES) for the combined treatment was statistically greater than DEDC treatment alone. No significant changes in the levels of kidney copper were seen with any treatment group. Similarly, no significant differences in serum copper levels were observed for any treatment group. For controls, serum copper was 130 ± 19.7 µg/dl; for the DEDC-exposed the copper level was 150 ± 13.6 µg/dl; for the high copper diet-exposed the level was 128 ± 7.2 µg/dl; and for the high copper diet plus DEDC exposure group, the level was 142 ± 9.3 µg/dl.
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DISCUSSION |
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Peripheral nerve lesions were observed both in animals exposed to DEDC alone and in animals exposed to DEDC in combination with a high copper diet, but it was not possible to distinguish a difference between these two exposure groups based on quantification of lesion severity. The most severely affected individuals were in the DEDC normal copper diet exposure group, but the median level of lesion severity was identical for both DEDC exposure groups. Although this result may suggest that elevated copper does not enhance DEDC toxicity, the small increase in copper observed in the sciatic nerve from the high copper diet may not have been sufficient to produce a significant difference in lesion severity given the sample size and the variation in lesion severity among individuals. Therefore, it is not possible based on the present study to determine whether further elevation of nervous system copper levels would enhance DEDC neurotoxicity, suggesting a cytotoxic effect of copper, or decrease the neurotoxicity, suggesting that DEDC may be acting through sequestering of critical copper cofactors.
Deposition of metal was observed by metallography staining in the liver, sciatic nerve, and brain of animals exposed to DEDC and elevated dietary copper. This staining technique relies on precipitation of silver on metal sulfides in tissue sections and is thought to stain zinc and copper sulfides most efficiently (Danscher, 1984). The lack of DEDC-mediated increases in metals other than copper in any of the treatment groups suggests that the silver reaction product observed in tissue sections is due to the presence of copper, although precipitate formation due to some other compound-related effect cannot be completely ruled out. In the liver, staining was observed in a periportal distribution, consistent with the observed localization of copper in Wilson's disease and in copper toxicosis (Bremner, 1998
; Haywood et al., 1985
). This precipitate was not associated with hepatotoxicity, however, based on the absence of elevated AST/SGOT activity and lack of significant histopathological changes in liver sections. The copper observed in these sections may have been effectively detoxified through binding to metallothionein; rats are relatively tolerant of copper excess, and hepatotoxicity typically does not occur below doses of 500 mg/kg/day (Bremner, 1998
). Regional precipitate was also observed in brain sections from animals exposed to both a high copper diet and DEDC, but not in control animals on either diet. Staining was observed in the hippocampus, the caudate, the thalamus, and layer VI of the cortex, but not in the rest of the cortex or the majority of the cerebellum. Previous studies have demonstrated localization of copper following DEDC exposure to the hippocampus, cortex, and brain stem by AAS analysis of these regions, although the results have not been consistently replicated among investigators (Ho et al., 1985
; Lakomaa et al., 1982
; Szerdalheyi and Kasa, 1987
). The absence of lesions as assessed by thick sections of representative brain regions in the present and previous studies suggests that the accumulation of copper is not associated with morphological changes in this tissue, and deposition of copper may reflect storage or detoxification sites for copper, rather than sites of damage, as was observed here for the liver. However, deposition of copper in sciatic nerve of rats exposed to DEDC was associated with myelin lesions. Interestingly, the staining observed in sciatic nerve appeared consistent with copper being distributed along the length of the axon, possibly in the myelin sheath or associated with the axon, whereas the staining in brain was punctate, suggesting it may be localized to a specific cell type.
Isoprostanes are products of free radical damage to arachidonic acid, and the measurement of the major class of isoprostanes, F2-isoprostanes, has been used widely to assess free radical-mediated lipid peroxidation in vivo (Roberts, 2000). Within the nervous system, arachidonic acid is homogeneously distributed throughout gray and white matter. Neuroprostanes are generated from free radical damage to docosohexanoic acid that, unlike arachidonic acid, is enriched in the gray matter of the CNS, providing more specific localization of free radical damage (Reich et al., 2001
). Significant elevations of F2-isoprostanes were observed in sciatic nerve for the DEDC-treated rats but no elevations in either F2-isoprostanes or F4-neuroprostanes were observed in brain for any treatment group. Elevated isoprostanes and myelin lesions in sciatic nerve without similar changes in brain support a correlation of lipid peroxidation with DEDC-mediated myelin lesions, although it cannot be determined from the present study whether lipid peroxidation is a contributing mechanism or a secondary effect of the myelin injury. These results are consistent with the lack of observable morphological lesions in the brain, despite high copper deposition in this tissue, suggesting that the peripheral nervous system is more susceptible to DEDC-mediated demyelination. An increased susceptibility of the peripheral nervous system may arise from inherent differences between Schwann cells and oligodendrocytes and their associated myelin or possibly result from a lack of astrocytes that may sequester excess copper and protect oligodendrocytes and myelin in the central nervous system.
The results of the current study demonstrate that ip administration of DEDC increases copper levels in sciatic nerve, demonstrating a role for DEDC in elevating peripheral as well as central nervous system copper levels. But significant differences were observed for DEDC-mediated myelin injury, lipid peroxidation, and the distribution of copper in the peripheral nervous system relative to the central nervous system. Further studies are required to delineate the cellular and subcellular sites of copper deposition in the peripheral and central nervous systems to evaluate the role of copper deposition and lipid peroxidation in DEDC neurotoxicity and to determine the basis for the relative sensitivities of the central and peripheral nervous systems. The design of experimental methods or dithiocarbamate analogs that can more effectively modulate the ability of dithiocarbamates to modify copper levels in the nervous system will aid in determining the role of copper elevation in disulfiram neurotoxicity. Relatively few neurotoxicants have been identified that produce injury to Schwann cells and myelin as a primary rather than secondary effect (Morell and Toews, 1996), and for those that have been identified, the mechanism is often poorly understood. Therefore, elucidating the mechanism of action for DEDC and disulfiram will aid in understanding how xenobiotics can interfere with the normal functioning of the peripheral nervous system and advance our understanding of the biology of this system. Additionally, determining the mechanisms responsible for dithiocarbamate-mediated demyelination will aid in the development of structure-activity relationships useful for predicting agents that may act through similar mechanisms, formulating safer dithiocarbamates and identifying potential effects of long-term low-level exposures.
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SUPPLEMENTARY DATA |
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed at Department of Pathology, 1401 21st Ave. S., Annex to MCS Rm 108, Vanderbilt University Medical Center, Nashville, TN, 372322561. Fax: (615) 343-9825. E-mail: bill.valentine{at}vanderbilt.edu.
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