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Depression of glutathione by cold-restraint in mice
Toxicology, 61 (1990) 59-71 Elsevier Scientific Publishers Ireland Ltd.
Depression of glutathione by cold-restraint in mice Henry F. Simmons a, Robert C. James b, Raymond D. Harbison b, and S t e p h e n M. R o b e r t s °'* ~Department of Pharmacology and Toxicology, University of Arkansas for Medical Sciences, Little Rock, AR, and bCenter for Environmental Toxicology, University of Florida, Gainesville, FL (U. S. A) (Received June 29th, 1989; accepted October 17th, 1989)
Summary The effects of cold-restraint as a physiological stressor on the glutathione (GSH) content of the liver and other tissues were examined in male mice. Mice of the ICR, NIH, ND/4, and B6C3F1 strains subjected to cold-restraint for 2 or 3 h experienced a loss of hepatic GSH concentrations ranging from approximately 15 to 50%. Though 3 of these strains (ICR, NIH, and B6C3F1) experienced hypothermia as result of the cold-restraint treatment, with average decreases in core body temperature ranging from 3.3 to 9.8°C, hepatic GSH levels were depressed in the ND/4 mouse in the absence of changes in core body temperature. The ability of cold-restraint as a stressor to diminsh hepatic GSH therefore could not be attributed simply to hypothermia. The decrease in hepatic GSH from cold-restraint in ND/4 mice was paralleled by a decrease in non-protein sulfhydryl (NPSH) content of the liver. In addition to its effects on liver GSH and NPSH concentrations, 1.5 h of cold-restraint stress significantly depressed plasma, heart, kidney, and lung NPSH concentrations. The extent of NPSH depression was equivalent to the GSH depression in the liver, heart, and kidney, despite the observation that the normal contribution of GSH to total NPSH content in these tissues ranged from a high of 89% (liver) to a low of 49% (heart). These results with cold-restraint in the ND/4 mouse suggest that other stressors may significantly depress cellular concentrations of GSH and other thiols, and may thereby render the affected tissues more susceptible to the toxicity of free radicals, electrophilic xenobiotic metabolites, or reactive oxygen species.
Key words: Glutathione; Cold-restraint; Stress; Non-protein sulfhydryl content
Introduction In 1888, glutathione or y-glutamyl-cysteinyl-glycine was discovered in yeast. Forty-seven years later its structure was elucidated. This tripeptide is found in virtually every living cell with the possible exception of some bacteria. Quite probably, it is the most abundant low molecular weight thiol that occurs naturally, with many tissue concentrations in the millimolar range [1]. The bulk of the Address all correspondence to: Dr. Stephen M. Roberts, Center for Environmental Toxicology, University of Florida, One Progress Blvd., Mailbox 17, Alachua, FL 32615, U.S.A. 0300-483X/90/$03.50 (~) 1990 Elsevier Scientific Publishers Ireland Ltd. Printed and Published in Ireland
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reduced glutathione (GSH) in most cells is found in the cytosol where it plays a role in numerous biochemical functions. These functions include the scavenging of free radicals, electrophilic metabolites and reactive oxygen species; the synthesis of deoxyribonucleotide precursors; and the reduction of various disulfide linkages [21. Because of the importance of GSH in the detoxification of a number of compounds, a significant depletion of GSH may increase the toxicity of these agents. There are several examples in the literature where a chemically-induced depletion of GSH (e.g. with diethyl maleate) has resulted in a potentiation of the toxicity of a compound (e.g. acetaminophen [3], bromobenzene [4], doxorubicin [5], and cocaine [6]). Depression of GSH by physiological means can also apparently influence toxicity as indicated by the effects of diurnal fluctuations in GSH levels on 1,1-dichloroethylene toxicity [7]. One of the more interesting, and perhaps important, non-chemically induced means of depressing GSH concentrations is through physiological stress. Studies spanning decades have shown that rats or mice subjected to various stressors have diminished hepatic levels of GSH, or of non-protein sulfhydryl (NPSH) content of which GSH is the principal component. Examples of stressors observed to cause a reduction in hepatic GSH or NPSH content include limb ligation trauma, scalding, bacterial endotoxins, tumbling trauma, hemorrhage, and insulin shock [8-11]. While changes have been observed occasionally in NPSH or GSH content of tissues other than the liver in stressed animals, there is little in the way of comprehensive information on stress-induced effects on GSH in extrahepatic tissues. In order to further examine the influence of stress on GSH, we have measured the effects of cold-restraint on tissue GSH concentrations in mice. Cold-restraint of laboratory rodents has been used for decades to study stress-related phenomena, and rats subjected to cold-restraint have been shown to develop gastric ulcers within a few hours [12,13]. Beck and Linkenheimer [8[ observed that cold exposure depressed hepatic NPSH content in mice, but cold plus restraint was found to be more effective in lowering hepatic NPSH content than cold alone in a study in rats [14]. Cold-restraint is a relatively humane stressor, and can be readily applied to varying extents to examine "dose"-response relationships. Initial experiments were directed to characterizing the effects of cold-restraint on hepatic GSH concentrations in mice, and later experiments examined the magnitude and temporal course of cold-restraint changes in NPSH and GSH concentrations in extra-hepatic tissues including spleen, stomach, lung, heart, kidney, and plasma. Materials and methods
Chemicals
Trichioroacetic acid, perchloric acid, disodium ethylenediamine tetraacetate, disodium hydrogen phosphate, potassium dihydrogen phosphate, and methanol (HPLC grade) were purchased from Fisher Chemical Company (Fair Lawn, NJ)
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as ASC grade. 5,5'-Dithiobis (2-nitrobenzoic acid), reduced glutathione, sodium chloride, monochloroacetic acid, and penicillamine (DL-isomeric mixture) were purchased from Sigma Chemical Company (St. Louis, MO). Sodium hydroxide (ACS grade) was purchased from MCB Manufacturing Chemists, Inc. (Cincinnati, OH), and sodium octyl sulfate was obtained from Eastman Kodak Company (Rochester, NY). Triple-distilled mercury was purchased from Bioanalytical Systems (West Lafayette, IN).
Animals and Treatments ICR, NIH, and ND/4 male mice all weighing 20-25 g were purchased from Harlan Sprague Dawley (Indianapolis, IN). Male B6C3F1 mice, 20-30 g, were obtained from the National Center for Toxicological Research (Jefferson, AR). The animals were housed 4-5 per cage in stainless steel and polycarbonate cages with hardwood chip bedding material. Aside from restraint periods, all animals had free access to food and water. A 12-h light/dark cycle was provided in the animal quarters, and the temperature was maintained at 22_+ I°C. Relative humidity ranged from 40 to 60%. Treatment and control group sizes ranged from a minimum of 5 to a maximum of 10 animals. All treatments were initiated in the morning, usually between 0700 and 0900 h. Sacrifice times varied with experimental design, but a control group was always sacrificed at the same time of day as a treatment group to preclude an influence of diurnal variation in GSH on experimental outcome. In studies determining strain responsiveness to cold-restraint and in studies in which the cold-restraint "dose"-response relationship was evaluated, treated animals were sacrificed by asphyxiation with carbon dioxide immediately post-cold-restraint. In the multiple tissue time course studies, groups of animals and their controls were asphyxiated immediately after termination of cold-restraint (0 h), or 1, 2, 3, or 5 h later. Cold-restraint was carried out in a draft-free portion of a refrigerated room maintained at 7.5 + 1°C. The restrainers used were modified 50 ml polystyrene centrifuge tubes with screw-on caps supplied by Fisher Scientific Company (Fair Lawn, N J). The wall of each tube was perforated with 4 rows of 44 2-mm holes. A single 5 mm perforation was drilled through the tapered tip. Five additional 2-mm holes were bored through the cap. Animals readily entered the tubes without shoving, orienting their heads at the tapered ends. An approximate 2:1 ratio of tube volume to body volume was achieved in most cases. In the cold room, each loaded tube was placed in a wooden rack that insured its isolation from other tubes. Rectal temperatures were measured with a flexible, small rodent rectal probe coupled to a calibrated electronic thermometer (YSI Instruments, Yellow Springs, OH).
Assay for reduced glutathione (GSH) Tissues samples were excised, rinsed with cold saline, blotted dry, and weighed. These samples, weighing between 0.3 and 0.6 g, were homogenized in 3 ml of
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cold 0.3 M perchloric acid with 0.1% EDTA. Immediately after homogenization, precipitated protein was pelleted at 10 000 x g for 15 min in a refrigerated (4°C) centrifuge. A 50-/xl aliquot of the supernatant was pipetted into a 0.2-/xm pore-size microfiltration tube (Bioanalytical Systems, West Lafayette, IN) along with 250/xl cold 0.1% EDTA and 10/zl penicillamine internal standard solution (penicillamine 1 mg/ml in 0.1% EDTA). The microfiltration tube was capped, vortexed, and centrifuged to accomplish the mixing and filtration of the sample solution. The filtered samples were placed on ice for analysis the same day or stored frozen at -80°C for assay within 72 h. Previous stability studies revealed no significant differences in tissue GSH concentrations among samples analyzed immediately or stored for 5 days under these conditions (unpublished observations). GSH content of tissue samples was measured by HPLC with electrochemical detection in a manner similar to that described by Allison and Shoup [15]. A Bioanalytical Systems LC-448 Liquid Chromatograph was used equipped with modifications to exclude oxygen from the system including the mobile phase reservoir. The mobile phase consisted of 0.075 M monochloroacetic acid and 3% acetonitrile, adjusted to pH 2.8 with sodium hydroxide. Sodium octyl sulfate was included at a concentration of 348 mg/l as an ion-pairing agent. Chromatographic separation was achieved with a 3/xm, 3.2 x 100 mm reverse phase (ODS) column (Bioanalytieal Systems, West Lafayette, IN). The potential of the gold/mercury electrode vs. silver/silver chloride reference was 0.150 V.
Assay for non-protein sulfhydryl (NPSH) content NPSH content was measured by the method of Ellman [16]. Tissues were processed in the fashion described above for the measurement of GSH, with samples removed and homogenized under cold conditions with 0.3 M perchloric acid. Following cold (4°C) centrifugation of the tissue homogenate, protein-free supernatant was diluted with cold 0.1% EDTA. For heart tissue, 400 /xl of deproteinized supernatant was added to 100 txl of 0.1% EDTA prior to assay. For kidney, spleen, lung, and stomach, 200 /xl of supernatant was added to 100 txl 0.1% EDTA. The preparation of plasma for NPSH measurement differed from that of other tissues somewhat. Immediately following carbon dioxide asphyxiation, blood was collected from mice via cardiac puncture into a heparinized syringe. The blood was centrifuged at 13 000 x g for 3 min to recover the plasma. A 250-/xl aliquot of plasma was quickly pipetted into 500/xl cold 0.3 M perchloric acid in 0.1% EDTA and allowed to stand in ice for approximately 5 min. Following a 5-rain spin at 13 000 x g, the supernatant was removed for measurement of NPSH content.
Statistical analyses For most experiments in this study, each treatment group had a distinct, matching control group. For these experiments, the treatment and control groups were compared with an unpaired Student's t-test. In those experiments in which
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more than one treatment group was compared with a single control group sacrificed at the same time, statistical comparisons were made using the Dunnett's test. In all instances, statistical significance was considered to be P ~<0.05. Results In initial experiments, 3 mouse strains were evaluated for their susceptibility to cold-restraint induced depression of hepatic GSH, viz. the ICR, NIH, and N D / 4 strains. Later, response in a fourth strain (B6C3F1) was examined, albeit with a somewhat greater stressor period (3 vs. 2 h). Immediately following coldrestraint, hepatic G S H concentrations were significantly depressed in all strains (Table I). The depression in the ICR strain was modest, approximately 15%, while much greater depression (32.4-36.8%) was observed in mice of N I H or N D / 4 strains. The extent of hepatic G S H depression in B6C3F1 mice is not strictly comparable, since the duration of cold-restraint was longer than for the other strains. However, this strain appears to be at least as sensitive as the N I H and N D / 4 strains, and lost almost 50% of hepatic GSH after 3 h of cold-restraint. The ability to sustain core body temperature during cold-restraint was variable among the strains. The ICR, NIH, and B6C3F1 mice all became hypothermic (Table I). Rectal temperatures in cold-restrained ICR and N I H mice were approximately 3-4°C below those of control mice of the same strains housed unrestrained at room temperature, a statistically significant difference. While the body temperature changes in the B6C3F1 mouse are not strictly comparable to the other strains because of differences in cold-restraint duration, it is apparent that mice of this strain had difficulty maintaining body temperature when exposed to cold and became profoundly hypothermic. The N D / 4 strain, in contrast, suffered no significant loss of core temperature. In a subsequent experiment, the relationship between core temperature and hepatic G S H in the N D / 4 mouse was examined more closely, with rectal temperature and G S H measurements obtained for groups of N D / 4 mice subjectTABLE I E F F E C T O F C O L D - R E S T R A I N T ON H E P A T I C G L U T A T H I O N E L E V E L S IN V A R I O U S MOUSE STRAINS Mouse strain
Temperature change (°C)
Restraint time
Glutathione change ( % )
ICR NIH B6C3F1 ND/4
-3.8 -3.3 -9.8 -1.8
2h 2h 3h 2h
-15.7 -32.4 -48.4 -36.8
-+ 0.9* -+ 0.7* -+ 0.8* _+ 0.5
-+ 4.6* -+ 6.6* -+ 2.2* -+ 4.4*
Mice were restrained in a cold e n v i r o n m e n t (6.5°C) for the time periods indicated, and the results compared with control, unrestrained mice housed at room temperature (25°C). Results are expressed as m e a n _+ S.E.M., N = 5. Control G S H values were: ICR, 2.39 -+ 0.06; NIH, 1.82 -+ 0.10; B6C3F1, 3.10-+0.06; and N D / 4 , 2.85 -+0.08 m g / g liver. *Indicates significantly different from control as determined by Student's t-test, P < 0.05.
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ed to cold-restraint for intervals up to 3 h. At no time point was a significant change in core temperature observed (Fig. 1A). A significant depression of hepatic GSH concentration, over 20%, was observed at the earliest time point (0.5 h) (Fig. 1B). The extent of loss of hepatic GSH increased with increasing exposure to cold-restraint. At the longest cold-restraint period examined, 3 h, hepatic GSH concentrations were depressed greater than 40% compared to controls. To provide an indication of the rate at which hepatic GSH concentrations recovered, the time course of changes in NPSH concentration after 1.5 h of cold-restraint was measured in N D / 4 mice. Liver and plasma were measured together because plasma NPSH levels are known to be dependent in part on efllux from the liver [17]. As shown in Fig. 2, changes in plasma NPSH concentrations during recovery from cold-restraint paralleled those of the liver. A maximal decrease of approximately 25% was seen immediately after the stress was terminated and a significant depression of NPSH was still evident in both liver and plasma 3 h later. The time courses of NPSH changes in other tissues after cold-restraint treatment were more variable and did not mimic those of liver and plasma (Table II). For example, 1.5 h of cold-restraint significantly lowered kidney NPSH concentrations (0-h sample), but this effect was readily reversed and normal levels were measured at 1, 2, and 3 h post-stressor. No significant changes in splenic NPSH concentration were observed, and while lung NPSH concentrations were consistently less than control values at all time points, only the 1 h post-stressor group was significantly depressed. The heart showed a delayed but persistent decrease in NPSH concentrations, with significant decreases in NPSH concentration at 2 and 3 h after termination of cold-restraint. A later measurement of heart NPSH at 5 h post-cold-restraint found levels to be 83.9 ± 6.5% of control, which was also a significant depression ( P < 0 . 0 5 ) . Thus, the diminished NPSH in the heart appears to persist for at least 5 h after cold-restraint, and perhaps longer. Results in the stomach were variable, with some evidence of a transient rise in NPSH levels. GSH comprises the vast majority of the NPSH in the liver [18], and significant losses of NPSH in this organ can be presumed to reflect losses in GSH. Because the G S H / N P S H relationship in other tissues is less clearly defined it was of interest to determine if the NPSH changes observed in extra-hepatic tissues in response to cold-restraint were in fact due to, or reflective of, changes in G S H levels. In order to test this, samples from tissues with depressed NPSH levels were also assayed for GSH content at the time point of lowest NPSH content. In the case of the heart, this was the 2-h post-cold-restraint samples, for kidney it was the samples taken at the cessation of cold-restraint (0 h), and for lung NPSH and GSH concentrations were compared in 1-h samples (Table III). The corresponding tissue samples from control mice at these same time intervals were also assayed for NPSH and G S H content to provide the control values presented in Table III. There were substantial differences among tissues in NPSH and GSH content, with l i v e r > k i d n e y > spleen > l u n g > heart. Griffith and Meister [19], also
64
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C o l d - R e s t r a i n t T i m e (hrs) Fig. I. Rectal temperatures and hepatic G S H changes in cold-restrained N D / 4 Mice. (A) Rectal temperatures were m e a s u r e d immediately following cold-restraint (7.5°C environmental temperature) for periods ranging from 0.5 to 3 h. Results are expressed as m e a n -+ S.E.M., N / > 5 per group. None of the cold-restrained groups had body temperatures significantly different from those of control mice housed unrestrained at room temperature. (B) Hepatic G S H concentrations were m e a s u r e d immediately following cold-restraint of varying duration. Data are expressed as a percent of the m e a n G S H concentration of a control group housed unrestrained at room temperature (2.45 -+ 0.11 mg G S H / g liver, m e a n -+ S.E.M., N = 6). T h e timing of the initiation of cold-restraint treatments were staggered such that groups treated for varying intervals and the control group were all sacrificed at the same time of day. For each t r e a t m e n t group, the m e a n percent + S . E . M is plotted, with N / > 5 per group. *Indicates significantly different from control mice kept unrestrained at r o o m temperature.
65
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Fig. 2. Time course of hepatic and plasma NPSH concentrations after cold-restraint. ND/4 mice were subjected to cold-restraint (7.5°C environmental temperature) for 1.5 h. Immediately following cold-restraint, mice were sacrificed at intervals up to 5 h. Data are expressed as a percent of the mean control hepatic or plasma NPSH concentration for each time interval. For each group, the mean +-S.E.M. is plotted, with N ~ 5 per group. Mean NPSH concentrations in control groups declined over the observation period, ranging from 2.99 to 2.41 mg/g in liver, and from 0.0824 to 0.0413 mg/ml in plasma. *Indicates significantly different from control mice.
noted this relationship in tissue concentrations when total glutathione (reduced plus oxidized) was measured in NCS male mice. Their observed values for total glutathione in untreated mice for these tissues (liver, 2.35 rag/g; kidney, 1.27 m g / g ; spleen, 1.05 m g / g ; lung, 0.46 mg/g; and heart, 0.414 m g / g tissue) were similar to G S H values measured in control N D / 4 mice (Table III). Modest differences between studies might be attributed to strain variations in tissue glutathione content, as evident in the comparison of control hepatic GSH levels among 4 strains in Table I, and to the differences inherent in the nature of the measurement (i.e. total glutathione vs. reduced glutathione only in the present study). In addition to differences among tissues in NPSH and GSH concentration, there were also substantial differences among tissues with respect to the portion of total NPSH content that is GSH. In control samples G S H accounted for almost 90% of the measurable hepatic NPSH, while in the heart it was only about 50% of NPSH levels. In kidney and lung control tissue samples, approximately three-quarters of the N P S H was GSH. The ratios of G S H / N P S H in control vs. treated (cold-restrained) mice indicated that the extent of depression occurred equally for NPSH and GSH in heart, kidney, and liver. While the depression of
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T A B L E II I N F L U E N C E OF C O L D - R E S T R A I N T ON T H E N O N - P R O T E I N S U L F H Y D R Y L C O N T E N T O F V A R I O U S TISSUES IN T H E N D / 4 M O U S E Time after termination of cold-restraint treatment (h) Tissue (group)
(0)
(1)
Spleen Control Treatment
100 -+ 7.9 97.9-+ 5.5
100 116
Kidney Control Treatment
(2)
(3)
-+ 6.8 -+ 12.2
100 ± 10 87.0± 9.5
100 124
± 13.0 ± 8.5
100 -+ 5.1 79.5-+ 1.8"
100 ± 2.5 98.1-+ 3.3
100 -+ 4.5 97.1± 3.5
100 112
-+ 6.3 ± 4.1
Heart Control Treatment
100 ± 8.4 95.7± 7.7
100 113
100 -+ 5.4 73.1± 6.8*
100 -+ 6.1 85.5-+ 2.1"
Lung Control Treatment
100 ± 2 4 . 0 94.2 -+ 23.0
100 ± 16.0 80.1 -+ 3.6*
100 ± 19.0 88.9 ± 13.0
100 -+ 18.0 94.6 -+ 3.9
Stomach Control Treatment
100 -+ 6.6 90.4+ - 9.7
100 119
100 120
100 ± 9.8 97.6± 6.3
-+ 8.2 ± 4.4
-+ 9.6 -+ 6.9*
± 11.0 ± 13.0
Mice were restrained in a cold environment (6.5°C) (treatment groups) or housed unrestrained at ambient temperatures (25°C) (control groups) for 1.5 h. Immediately after cold-restraint (0 h), or 1, 2, or 3 h after treatment, mice were sacrificed and tissues harvested for measurement of non-protein sulfhydryl (NPSH) content. Results are expressed as a percent of the mean control NPSH content for each time point + S.E.M., N = 5. Mean control NPSH values (mg/g tissue) ranged from 0.858 to 1.119 for spleen, 1.071 to 1.262 for kidney, 0.393 to 0.453 for heart, 0.575 to 0.762 for lung, and 0.911 to 1.205 for stomach. *Indicates significantly different from control as determined by Student's t-test, P < 0.05.
GSH in response to cold-restraint in the lung appeared to be less than that of other thiols comprising the NPSH pool, the unusually large variability in NPSH levels measured in this tissue make the reliability of this observation uncertain. Discussion The capacity of cold exposure to suppress the hepatic NPSH content of male CAF1 mice was first observed decades ago [8]. In 1953, Bartlett and Register confirmed this finding in rats, and demonstrated that concommitant restraint of the animal by placing it in a wire mesh cylinder enhanced this effect [14]. Simple chilling of rats at 0°C for 2 - 4 h depressed hepatic NPSH concentrations only about 5%, while the addition of restraint resulted in 4-fold greater decreases, i.e. to levels that were 20% lower than those of control animals. We have also observed a marked enhancement of hepatic G S H depression when cold exposure
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T A B L E III C O M P A R I S O N OF N O N - P R O T E I N S U L F H Y D R Y L A N D G L U T A T H I O N E C O N T E N T OF S E L E C T E D TISSUES F R O M C O N T R O L A N D C O L D - R E S T R A I N E D N D / 4 M I C E
Tissue content (mg/g tissue) Tissue (group)
NPSH
GSH
(GSH/NPSH)
Ratio
Heart Control Treatment
0.540 _+ 0.029 0.396 _+ 0.027
0.264 _+ 0.009 0.20l +_ 0.014
0.49 0.51
Kidney Control Treatment
1.22 _+0.06 0.964 -+ 0.017
0.940 + 0.058 0.704 + 0.009
0.77 0.73
Liver Control Treatment
2.99 + 0.17 2.16 _+ 0.12
2.65 _+ 0.15 1.81 + 0.06
0.89 0.84
0.762 -+ 0.121 0.610 _+ 0.061
0.571 _+ 0.002 0.545 _+ 0.008
0.75 0.89
Lung
Control Treatment
Tissue samples were divided and assayed for non-protein sulfhydryl (NPSH) and reduced glutathione ( G S H ) content as described in Methods. Samples for each tissue were taken from the time of maximal depression of NPSH after cold-restraint: heart, 2 h; kidney, 0 h; liver, 0 h; and lung, I h. Results are expressed as mean _+ S.E.M., N - 5 . The NPSH and GSH content of tissues from treated (coldrestrained) mice were significantly different from the corresponding controls, P < 0.05, in all cases except the GSH values for lung.
and restraint are combined as stressors. In ICR mice, exposures of unrestrained mice to cold (8.5°C) for periods up to 5 h had no effect on hepatic G S H concentrations (unpublished observations) while a 2-h exposure of restrained mice under similar conditions resulted in a significant loss of GSH (15.7% decrease, P < 0.05; Table I). Restraint without cold was an insufficient stressor to produce a change in hepatic GSH concentrations (unpublished observations). Cold-restraint was employed in this study as a stressor, with the hope that the results would be applicable to responses to stressors in general. It was therefore important that the response to cold-restraint not be dependent upon a unique feature of this stressor such as hypothermia. When Beck and Linkenheimer first reported a fall in the hepatic NPSH content of CAF1 mice in mice exposed to cold (5-10°C for 5 h), they noted only a 1-2°C drop in rectal temperatures compared to pre-stress levels [8]. Bartlett and Register [14] had a different experience with cold-restrained ( 2 - 4 h at 0°C) S p r a g u e - D a w l e y rats, and found that a significant decline in hepatic GSH took place in animals whose core temperatures had fallen between 15 and 25°C. In our comparisons of hepatic GSH response to cold-restraint among 4 m o u s e strains, it was apparent that depression of hepatic GSH levels was independent from effects on core body temperature. The N D / 4 mouse was able to sustain core temperature during cold-restraint for
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periods up to 3 h, during which hepatic GSH concentrations declined to approximately 50% of control levels. The GSH depression, at least in the ND/4 mouse, therefore cannot be attributed simply to hypothermia and may reflect a process common to a variety of stressors. The effects of cold-restraint were not confined to the liver, and plasma NPSH concentrations mirrored those in the liver over time following cold-restraint. If subsequent animal experiments bear out this relationship, these observations suggest that plasma GSH or NPSH measurements might serve as a useful, relatively non-invasive means of studying hepatic GSH or NPSH changes in response to stressors in humans. Plasma GSH does not always mimic hepatic GSH changes however, and plasma GSH concentrations in animals subjected to exhaustive exercise or administered oxidant drugs have been observed to remain the same or increase while hepatic levels were diminished [20,21]. These differences in plasma-liver GSH relationships suggest different mechanisms for loss of GSH in the liver. Rather than the oxidation of GSH which appears to occur with exercise or oxidant drug exposure, stress may diminish hepatic GSH through other means such as decreased synthesis or increased intracellular catabolism. Cold-restraint effects on tissues other than liver and plasma were somewhat more variable. The most lasting response was the depression observed in the heart, which showed a delayed decline in NPSH content that persisted through the last time interval examined, 5 h post-treatment. Effects in the kidney were far more transient, with the effects of cold-restraint lasting less than 1 h. Other tissues showed changes that were typically significant only at 1 time point. Though these results suggest that tissues such as the lung may also be susceptible to NPSH depression from cold-restraint, the temporal displacement of the decrease observed in these experiments and their transient nature require that they be interpreted with caution. The approximate 20% increase in NPSH in stomach observed 1 and 2 h after cold-restraint is also worth noting, though variability caused only the rise at 1 h to be statistically signficant. Other studies of the effects of stressors on extra-hepatic NPSH or GSH have been conducted in rats, and the results show some similarities and differences to the changes seen here in mice. For example, Reichard and coworkers observed significant declines in NPSH concentrations in both rat kidney and spleen (as well as liver) 1.5 h after whole body trauma [11], and a significant decrease in renal NPSH concentration has been observed in rats stressed by scalding [9]. Rats subjected to cold-restraint for 8-12 h had a 27% decline in gastric GSH [22]. It appears, through comparisons of N P S H / G S H ratios in tissues from control and cold-restrained mice, that changes in these 2 thiol parameters generally occurred in remarkably parallel fashion. In the case of the liver, this is not surprising since approximately 90% of the NPSH content is GSH. However GSH is only about 50% of the NPSH in the heart, and stress-induced changes in GSH in this tissue appear to be matched by changes in other soluble, non-protein thiols. Equivalent effects on GSH and non-GSH components of total cellular NPSH were also observed in the kidney. These observations suggest that either the mechanism producing the lowering of tissue GSH in response to cold-restraint stress operates equally with other soluble, non-protein (and perhaps protein)
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sulfliydryls, or that GSH is in a relatively rapid equilibrium with other soluble cellular sulfhydryls. In conclusion, application of cold-restraint as a stressor in mice produced a depression in hepatic GSH that was independent of hypothermia and increased in magnitude with increasing duration. Though the effects were generally less pronounced, GSH changes were also observed in other tissues. The extent of loss of GSH was probably not in itself sufficient to cause tissue injury. However, this loss might be sufficient to enhance the toxic insult produced by free radicals, electrophilic xenobiotic metabolites or reactive oxygen species dependent upon GSH for detoxification. Therefore, the results of this study suggest that a stress response may significantly depress cellular GSH/thiol concentrations in the liver and other tissues and thereby render these tissues more susceptible to certain chemical-induced toxicities.
Acknowledgements This research was supported in part by a grant from the National Institutes of Health (ES 05216), and by the Medical Research Endowment Fund, University of Arkansas for Medical Sciences.
References 1 A. Meister, Metabolism and functions of glutathione. TIBS, September (1981) 231. 2 N. Kaplowitz, The regulation of hepatic glutathione. Annu. Rev. Pharmacol. Toxicol,, 25 (1985) 715. 3 J.R. Mitchell, D.J. Jollow, W.Z. Potter, J.R. Gillette and B.B. Brodie, Acetaminophen-induced hepatic necrosis. IV. Protective role of glutathione. J. Pharmacol. Exp. Ther., 187 (1973) 211. 4 D.J. Jollow, J.R. Mitchell, N. Zampaglione and J.R. Gillette, Bromobenzene-induced liver necrosis. Protective role of glutathione and evidence for 3,4-bromobenzene epoxide as the hepatotoxic metabolite. Pharmacology, 11 (1974) 151. 5 P.G. Wells, R.C. Boerth, J.A. Oates and R.D. Harbison, Toxicological enhancement by a combination of drugs which deplete hepatic glutathione: acetaminophen and doxorubicin (Adriamycin). Toxicol. Appl. Pharmacol., 54 (1980) 197. 6 M.A. Evans and R.D. Harbison, Cocaine-induced hepatotoxicity in mice. Toxicol. Appl. Pharmacol., 45 (1978) 739. 7 R.J. Jaeger, R.B. Conolly and S.D. Murphy, Diurnal variation of hepatic glutathione concentration and its correlation with 1,1-dichloroethylene inhalation toxicity in rats. Res. Commun. Chem. Pathol. Pharmacol., 6 (1973) 465. 8 L.V. Beck and W. Linkenheimer, Effects of shock and cold on mouse liver sulfhydryl. Proc. Soc. Exp. Biol. Med., 81 (1952) 291. 9 L.V. Beck, W. Linkenheimer and A. Marraccini, Effects of tumbling trauma, scalding, and hemorrhage on rat tissue non-protein sulfhydryl. Proc. Soc. Exp. Biol. Med., 86 (1954) 823. 10 L.V. Beck and V.D. Rieck, Control mechanisms for trauma induced DLSH reaction (Decrease in mouse liver non-protein sulfhydryl). Proc. Soc. Exp. Biol. Med., 98 (1958) 466. 11 S.M. Reichard, N.M. Bailey and M.J. Galvin, Alterations in tissue glutathione levels following traumatic shock. Adv. Shock Res., 5 (1981) 37. 12 D.A. Brodie and L.S. Valitski, Production of gastric hemorrhage in rats by multiple stresses. Proc. Soc. Exp. Biol. Med., 113 (1963) 998. 13 L. Buchel and D. Gallaiare, Ulcere de contrainte chez le rat. a. Influence, sur la frequence des ulceres, du jeune et de la temperature de l'environment associes a des immobilisations de durees variables. Arch. Sci. Physiol., 21 (1967) 527.
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14 R.G. Bartlett and U.D. Register, Effect of cold and restraint on blood and liver non-protein sulfhydryl compounds. Proc. Soc. Exp. Biol. Med., 83 (1953) 708. 15 L.A. Allison and R.E. Shoup, Dual electrode liquid chromatography detector for thiols and disulfides. Anal. Chem., 55 (1983) 8. 16 G.L. Ellman, Tissue sulfhydryl groups. Arch. Biochem. Biophys., 82 (1959) 70. 17 M. Inoue, Y. Saito, E. Hirata, Y. Morino and S. Nagase, Regulation of redox states of plasma proteins by metabolism and transport of glutathione and related compounds. J. Protein Chem., 6 (1987) 207. 18 N.S. Kosower and E.M. Kosower, The glutathione status of cells. Int. Rev. Cytol., 54 (1978) 109. 19 O.W. Griffith and A. Meister, Glutathione: Interorgan translocation, turnover, and metabolism. Proc. Natl. Acad. Sci. U.S.A., 76 (1979) 5606. 20 H. Lew, S. Pyke and A. Quintanilha, Changes in the glutathione status of plasma, liver and muscle following exhaustive exercise in rats. FEBS Lett., 185 (1985) 262. 21 J.D. Adams, B.H. Lauterberg and J.R. Mitchell, Plasma glutathione disulfide as an index of oxidant stress in vivo: effects of carbon tetrachloride, dimethylnitrosamine, nitrofurantoin, metronidazole, doxorubicin, and diquat. Res. Commun. Chem. Pathol. Pharmacol., 46 (1984) 401. 22 S.C. Boyd, H.A. Sasame and M.R. Boyd, Effects of cold restraint stress on rat gastric and hepatic glutathione: a potential determinant of response to chemical carcinogens. Physiol. Behav., 27 (1981) 377.
71
Depression of glutathione by cold-restraint in mice Henry F. Simmons a, Robert C. James b, Raymond D. Harbison b, and S t e p h e n M. R o b e r t s °'* ~Department of Pharmacology and Toxicology, University of Arkansas for Medical Sciences, Little Rock, AR, and bCenter for Environmental Toxicology, University of Florida, Gainesville, FL (U. S. A) (Received June 29th, 1989; accepted October 17th, 1989)
Summary The effects of cold-restraint as a physiological stressor on the glutathione (GSH) content of the liver and other tissues were examined in male mice. Mice of the ICR, NIH, ND/4, and B6C3F1 strains subjected to cold-restraint for 2 or 3 h experienced a loss of hepatic GSH concentrations ranging from approximately 15 to 50%. Though 3 of these strains (ICR, NIH, and B6C3F1) experienced hypothermia as result of the cold-restraint treatment, with average decreases in core body temperature ranging from 3.3 to 9.8°C, hepatic GSH levels were depressed in the ND/4 mouse in the absence of changes in core body temperature. The ability of cold-restraint as a stressor to diminsh hepatic GSH therefore could not be attributed simply to hypothermia. The decrease in hepatic GSH from cold-restraint in ND/4 mice was paralleled by a decrease in non-protein sulfhydryl (NPSH) content of the liver. In addition to its effects on liver GSH and NPSH concentrations, 1.5 h of cold-restraint stress significantly depressed plasma, heart, kidney, and lung NPSH concentrations. The extent of NPSH depression was equivalent to the GSH depression in the liver, heart, and kidney, despite the observation that the normal contribution of GSH to total NPSH content in these tissues ranged from a high of 89% (liver) to a low of 49% (heart). These results with cold-restraint in the ND/4 mouse suggest that other stressors may significantly depress cellular concentrations of GSH and other thiols, and may thereby render the affected tissues more susceptible to the toxicity of free radicals, electrophilic xenobiotic metabolites, or reactive oxygen species.
Key words: Glutathione; Cold-restraint; Stress; Non-protein sulfhydryl content
Introduction In 1888, glutathione or y-glutamyl-cysteinyl-glycine was discovered in yeast. Forty-seven years later its structure was elucidated. This tripeptide is found in virtually every living cell with the possible exception of some bacteria. Quite probably, it is the most abundant low molecular weight thiol that occurs naturally, with many tissue concentrations in the millimolar range [1]. The bulk of the Address all correspondence to: Dr. Stephen M. Roberts, Center for Environmental Toxicology, University of Florida, One Progress Blvd., Mailbox 17, Alachua, FL 32615, U.S.A. 0300-483X/90/$03.50 (~) 1990 Elsevier Scientific Publishers Ireland Ltd. Printed and Published in Ireland
59
reduced glutathione (GSH) in most cells is found in the cytosol where it plays a role in numerous biochemical functions. These functions include the scavenging of free radicals, electrophilic metabolites and reactive oxygen species; the synthesis of deoxyribonucleotide precursors; and the reduction of various disulfide linkages [21. Because of the importance of GSH in the detoxification of a number of compounds, a significant depletion of GSH may increase the toxicity of these agents. There are several examples in the literature where a chemically-induced depletion of GSH (e.g. with diethyl maleate) has resulted in a potentiation of the toxicity of a compound (e.g. acetaminophen [3], bromobenzene [4], doxorubicin [5], and cocaine [6]). Depression of GSH by physiological means can also apparently influence toxicity as indicated by the effects of diurnal fluctuations in GSH levels on 1,1-dichloroethylene toxicity [7]. One of the more interesting, and perhaps important, non-chemically induced means of depressing GSH concentrations is through physiological stress. Studies spanning decades have shown that rats or mice subjected to various stressors have diminished hepatic levels of GSH, or of non-protein sulfhydryl (NPSH) content of which GSH is the principal component. Examples of stressors observed to cause a reduction in hepatic GSH or NPSH content include limb ligation trauma, scalding, bacterial endotoxins, tumbling trauma, hemorrhage, and insulin shock [8-11]. While changes have been observed occasionally in NPSH or GSH content of tissues other than the liver in stressed animals, there is little in the way of comprehensive information on stress-induced effects on GSH in extrahepatic tissues. In order to further examine the influence of stress on GSH, we have measured the effects of cold-restraint on tissue GSH concentrations in mice. Cold-restraint of laboratory rodents has been used for decades to study stress-related phenomena, and rats subjected to cold-restraint have been shown to develop gastric ulcers within a few hours [12,13]. Beck and Linkenheimer [8[ observed that cold exposure depressed hepatic NPSH content in mice, but cold plus restraint was found to be more effective in lowering hepatic NPSH content than cold alone in a study in rats [14]. Cold-restraint is a relatively humane stressor, and can be readily applied to varying extents to examine "dose"-response relationships. Initial experiments were directed to characterizing the effects of cold-restraint on hepatic GSH concentrations in mice, and later experiments examined the magnitude and temporal course of cold-restraint changes in NPSH and GSH concentrations in extra-hepatic tissues including spleen, stomach, lung, heart, kidney, and plasma. Materials and methods
Chemicals
Trichioroacetic acid, perchloric acid, disodium ethylenediamine tetraacetate, disodium hydrogen phosphate, potassium dihydrogen phosphate, and methanol (HPLC grade) were purchased from Fisher Chemical Company (Fair Lawn, NJ)
60
as ASC grade. 5,5'-Dithiobis (2-nitrobenzoic acid), reduced glutathione, sodium chloride, monochloroacetic acid, and penicillamine (DL-isomeric mixture) were purchased from Sigma Chemical Company (St. Louis, MO). Sodium hydroxide (ACS grade) was purchased from MCB Manufacturing Chemists, Inc. (Cincinnati, OH), and sodium octyl sulfate was obtained from Eastman Kodak Company (Rochester, NY). Triple-distilled mercury was purchased from Bioanalytical Systems (West Lafayette, IN).
Animals and Treatments ICR, NIH, and ND/4 male mice all weighing 20-25 g were purchased from Harlan Sprague Dawley (Indianapolis, IN). Male B6C3F1 mice, 20-30 g, were obtained from the National Center for Toxicological Research (Jefferson, AR). The animals were housed 4-5 per cage in stainless steel and polycarbonate cages with hardwood chip bedding material. Aside from restraint periods, all animals had free access to food and water. A 12-h light/dark cycle was provided in the animal quarters, and the temperature was maintained at 22_+ I°C. Relative humidity ranged from 40 to 60%. Treatment and control group sizes ranged from a minimum of 5 to a maximum of 10 animals. All treatments were initiated in the morning, usually between 0700 and 0900 h. Sacrifice times varied with experimental design, but a control group was always sacrificed at the same time of day as a treatment group to preclude an influence of diurnal variation in GSH on experimental outcome. In studies determining strain responsiveness to cold-restraint and in studies in which the cold-restraint "dose"-response relationship was evaluated, treated animals were sacrificed by asphyxiation with carbon dioxide immediately post-cold-restraint. In the multiple tissue time course studies, groups of animals and their controls were asphyxiated immediately after termination of cold-restraint (0 h), or 1, 2, 3, or 5 h later. Cold-restraint was carried out in a draft-free portion of a refrigerated room maintained at 7.5 + 1°C. The restrainers used were modified 50 ml polystyrene centrifuge tubes with screw-on caps supplied by Fisher Scientific Company (Fair Lawn, N J). The wall of each tube was perforated with 4 rows of 44 2-mm holes. A single 5 mm perforation was drilled through the tapered tip. Five additional 2-mm holes were bored through the cap. Animals readily entered the tubes without shoving, orienting their heads at the tapered ends. An approximate 2:1 ratio of tube volume to body volume was achieved in most cases. In the cold room, each loaded tube was placed in a wooden rack that insured its isolation from other tubes. Rectal temperatures were measured with a flexible, small rodent rectal probe coupled to a calibrated electronic thermometer (YSI Instruments, Yellow Springs, OH).
Assay for reduced glutathione (GSH) Tissues samples were excised, rinsed with cold saline, blotted dry, and weighed. These samples, weighing between 0.3 and 0.6 g, were homogenized in 3 ml of
61
cold 0.3 M perchloric acid with 0.1% EDTA. Immediately after homogenization, precipitated protein was pelleted at 10 000 x g for 15 min in a refrigerated (4°C) centrifuge. A 50-/xl aliquot of the supernatant was pipetted into a 0.2-/xm pore-size microfiltration tube (Bioanalytical Systems, West Lafayette, IN) along with 250/xl cold 0.1% EDTA and 10/zl penicillamine internal standard solution (penicillamine 1 mg/ml in 0.1% EDTA). The microfiltration tube was capped, vortexed, and centrifuged to accomplish the mixing and filtration of the sample solution. The filtered samples were placed on ice for analysis the same day or stored frozen at -80°C for assay within 72 h. Previous stability studies revealed no significant differences in tissue GSH concentrations among samples analyzed immediately or stored for 5 days under these conditions (unpublished observations). GSH content of tissue samples was measured by HPLC with electrochemical detection in a manner similar to that described by Allison and Shoup [15]. A Bioanalytical Systems LC-448 Liquid Chromatograph was used equipped with modifications to exclude oxygen from the system including the mobile phase reservoir. The mobile phase consisted of 0.075 M monochloroacetic acid and 3% acetonitrile, adjusted to pH 2.8 with sodium hydroxide. Sodium octyl sulfate was included at a concentration of 348 mg/l as an ion-pairing agent. Chromatographic separation was achieved with a 3/xm, 3.2 x 100 mm reverse phase (ODS) column (Bioanalytieal Systems, West Lafayette, IN). The potential of the gold/mercury electrode vs. silver/silver chloride reference was 0.150 V.
Assay for non-protein sulfhydryl (NPSH) content NPSH content was measured by the method of Ellman [16]. Tissues were processed in the fashion described above for the measurement of GSH, with samples removed and homogenized under cold conditions with 0.3 M perchloric acid. Following cold (4°C) centrifugation of the tissue homogenate, protein-free supernatant was diluted with cold 0.1% EDTA. For heart tissue, 400 /xl of deproteinized supernatant was added to 100 txl of 0.1% EDTA prior to assay. For kidney, spleen, lung, and stomach, 200 /xl of supernatant was added to 100 txl 0.1% EDTA. The preparation of plasma for NPSH measurement differed from that of other tissues somewhat. Immediately following carbon dioxide asphyxiation, blood was collected from mice via cardiac puncture into a heparinized syringe. The blood was centrifuged at 13 000 x g for 3 min to recover the plasma. A 250-/xl aliquot of plasma was quickly pipetted into 500/xl cold 0.3 M perchloric acid in 0.1% EDTA and allowed to stand in ice for approximately 5 min. Following a 5-rain spin at 13 000 x g, the supernatant was removed for measurement of NPSH content.
Statistical analyses For most experiments in this study, each treatment group had a distinct, matching control group. For these experiments, the treatment and control groups were compared with an unpaired Student's t-test. In those experiments in which
62
more than one treatment group was compared with a single control group sacrificed at the same time, statistical comparisons were made using the Dunnett's test. In all instances, statistical significance was considered to be P ~<0.05. Results In initial experiments, 3 mouse strains were evaluated for their susceptibility to cold-restraint induced depression of hepatic GSH, viz. the ICR, NIH, and N D / 4 strains. Later, response in a fourth strain (B6C3F1) was examined, albeit with a somewhat greater stressor period (3 vs. 2 h). Immediately following coldrestraint, hepatic G S H concentrations were significantly depressed in all strains (Table I). The depression in the ICR strain was modest, approximately 15%, while much greater depression (32.4-36.8%) was observed in mice of N I H or N D / 4 strains. The extent of hepatic G S H depression in B6C3F1 mice is not strictly comparable, since the duration of cold-restraint was longer than for the other strains. However, this strain appears to be at least as sensitive as the N I H and N D / 4 strains, and lost almost 50% of hepatic GSH after 3 h of cold-restraint. The ability to sustain core body temperature during cold-restraint was variable among the strains. The ICR, NIH, and B6C3F1 mice all became hypothermic (Table I). Rectal temperatures in cold-restrained ICR and N I H mice were approximately 3-4°C below those of control mice of the same strains housed unrestrained at room temperature, a statistically significant difference. While the body temperature changes in the B6C3F1 mouse are not strictly comparable to the other strains because of differences in cold-restraint duration, it is apparent that mice of this strain had difficulty maintaining body temperature when exposed to cold and became profoundly hypothermic. The N D / 4 strain, in contrast, suffered no significant loss of core temperature. In a subsequent experiment, the relationship between core temperature and hepatic G S H in the N D / 4 mouse was examined more closely, with rectal temperature and G S H measurements obtained for groups of N D / 4 mice subjectTABLE I E F F E C T O F C O L D - R E S T R A I N T ON H E P A T I C G L U T A T H I O N E L E V E L S IN V A R I O U S MOUSE STRAINS Mouse strain
Temperature change (°C)
Restraint time
Glutathione change ( % )
ICR NIH B6C3F1 ND/4
-3.8 -3.3 -9.8 -1.8
2h 2h 3h 2h
-15.7 -32.4 -48.4 -36.8
-+ 0.9* -+ 0.7* -+ 0.8* _+ 0.5
-+ 4.6* -+ 6.6* -+ 2.2* -+ 4.4*
Mice were restrained in a cold e n v i r o n m e n t (6.5°C) for the time periods indicated, and the results compared with control, unrestrained mice housed at room temperature (25°C). Results are expressed as m e a n _+ S.E.M., N = 5. Control G S H values were: ICR, 2.39 -+ 0.06; NIH, 1.82 -+ 0.10; B6C3F1, 3.10-+0.06; and N D / 4 , 2.85 -+0.08 m g / g liver. *Indicates significantly different from control as determined by Student's t-test, P < 0.05.
63
ed to cold-restraint for intervals up to 3 h. At no time point was a significant change in core temperature observed (Fig. 1A). A significant depression of hepatic GSH concentration, over 20%, was observed at the earliest time point (0.5 h) (Fig. 1B). The extent of loss of hepatic GSH increased with increasing exposure to cold-restraint. At the longest cold-restraint period examined, 3 h, hepatic GSH concentrations were depressed greater than 40% compared to controls. To provide an indication of the rate at which hepatic GSH concentrations recovered, the time course of changes in NPSH concentration after 1.5 h of cold-restraint was measured in N D / 4 mice. Liver and plasma were measured together because plasma NPSH levels are known to be dependent in part on efllux from the liver [17]. As shown in Fig. 2, changes in plasma NPSH concentrations during recovery from cold-restraint paralleled those of the liver. A maximal decrease of approximately 25% was seen immediately after the stress was terminated and a significant depression of NPSH was still evident in both liver and plasma 3 h later. The time courses of NPSH changes in other tissues after cold-restraint treatment were more variable and did not mimic those of liver and plasma (Table II). For example, 1.5 h of cold-restraint significantly lowered kidney NPSH concentrations (0-h sample), but this effect was readily reversed and normal levels were measured at 1, 2, and 3 h post-stressor. No significant changes in splenic NPSH concentration were observed, and while lung NPSH concentrations were consistently less than control values at all time points, only the 1 h post-stressor group was significantly depressed. The heart showed a delayed but persistent decrease in NPSH concentrations, with significant decreases in NPSH concentration at 2 and 3 h after termination of cold-restraint. A later measurement of heart NPSH at 5 h post-cold-restraint found levels to be 83.9 ± 6.5% of control, which was also a significant depression ( P < 0 . 0 5 ) . Thus, the diminished NPSH in the heart appears to persist for at least 5 h after cold-restraint, and perhaps longer. Results in the stomach were variable, with some evidence of a transient rise in NPSH levels. GSH comprises the vast majority of the NPSH in the liver [18], and significant losses of NPSH in this organ can be presumed to reflect losses in GSH. Because the G S H / N P S H relationship in other tissues is less clearly defined it was of interest to determine if the NPSH changes observed in extra-hepatic tissues in response to cold-restraint were in fact due to, or reflective of, changes in G S H levels. In order to test this, samples from tissues with depressed NPSH levels were also assayed for GSH content at the time point of lowest NPSH content. In the case of the heart, this was the 2-h post-cold-restraint samples, for kidney it was the samples taken at the cessation of cold-restraint (0 h), and for lung NPSH and GSH concentrations were compared in 1-h samples (Table III). The corresponding tissue samples from control mice at these same time intervals were also assayed for NPSH and G S H content to provide the control values presented in Table III. There were substantial differences among tissues in NPSH and GSH content, with l i v e r > k i d n e y > spleen > l u n g > heart. Griffith and Meister [19], also
64
A. 40 35 =1
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0
I
I
1
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I
I
2
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3
C o l d - R e s t r a i n t T i m e (hrs) Fig. I. Rectal temperatures and hepatic G S H changes in cold-restrained N D / 4 Mice. (A) Rectal temperatures were m e a s u r e d immediately following cold-restraint (7.5°C environmental temperature) for periods ranging from 0.5 to 3 h. Results are expressed as m e a n -+ S.E.M., N / > 5 per group. None of the cold-restrained groups had body temperatures significantly different from those of control mice housed unrestrained at room temperature. (B) Hepatic G S H concentrations were m e a s u r e d immediately following cold-restraint of varying duration. Data are expressed as a percent of the m e a n G S H concentration of a control group housed unrestrained at room temperature (2.45 -+ 0.11 mg G S H / g liver, m e a n -+ S.E.M., N = 6). T h e timing of the initiation of cold-restraint treatments were staggered such that groups treated for varying intervals and the control group were all sacrificed at the same time of day. For each t r e a t m e n t group, the m e a n percent + S . E . M is plotted, with N / > 5 per group. *Indicates significantly different from control mice kept unrestrained at r o o m temperature.
65
i
120 1
• liver I 0 plasma
110 1
O0
=
O 90
•
o I~
r~
$
V
60 so
I
I
I
i
i
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2
3
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Hours After Termination
of Cold-Restraint
Fig. 2. Time course of hepatic and plasma NPSH concentrations after cold-restraint. ND/4 mice were subjected to cold-restraint (7.5°C environmental temperature) for 1.5 h. Immediately following cold-restraint, mice were sacrificed at intervals up to 5 h. Data are expressed as a percent of the mean control hepatic or plasma NPSH concentration for each time interval. For each group, the mean +-S.E.M. is plotted, with N ~ 5 per group. Mean NPSH concentrations in control groups declined over the observation period, ranging from 2.99 to 2.41 mg/g in liver, and from 0.0824 to 0.0413 mg/ml in plasma. *Indicates significantly different from control mice.
noted this relationship in tissue concentrations when total glutathione (reduced plus oxidized) was measured in NCS male mice. Their observed values for total glutathione in untreated mice for these tissues (liver, 2.35 rag/g; kidney, 1.27 m g / g ; spleen, 1.05 m g / g ; lung, 0.46 mg/g; and heart, 0.414 m g / g tissue) were similar to G S H values measured in control N D / 4 mice (Table III). Modest differences between studies might be attributed to strain variations in tissue glutathione content, as evident in the comparison of control hepatic GSH levels among 4 strains in Table I, and to the differences inherent in the nature of the measurement (i.e. total glutathione vs. reduced glutathione only in the present study). In addition to differences among tissues in NPSH and GSH concentration, there were also substantial differences among tissues with respect to the portion of total NPSH content that is GSH. In control samples G S H accounted for almost 90% of the measurable hepatic NPSH, while in the heart it was only about 50% of NPSH levels. In kidney and lung control tissue samples, approximately three-quarters of the N P S H was GSH. The ratios of G S H / N P S H in control vs. treated (cold-restrained) mice indicated that the extent of depression occurred equally for NPSH and GSH in heart, kidney, and liver. While the depression of
66
T A B L E II I N F L U E N C E OF C O L D - R E S T R A I N T ON T H E N O N - P R O T E I N S U L F H Y D R Y L C O N T E N T O F V A R I O U S TISSUES IN T H E N D / 4 M O U S E Time after termination of cold-restraint treatment (h) Tissue (group)
(0)
(1)
Spleen Control Treatment
100 -+ 7.9 97.9-+ 5.5
100 116
Kidney Control Treatment
(2)
(3)
-+ 6.8 -+ 12.2
100 ± 10 87.0± 9.5
100 124
± 13.0 ± 8.5
100 -+ 5.1 79.5-+ 1.8"
100 ± 2.5 98.1-+ 3.3
100 -+ 4.5 97.1± 3.5
100 112
-+ 6.3 ± 4.1
Heart Control Treatment
100 ± 8.4 95.7± 7.7
100 113
100 -+ 5.4 73.1± 6.8*
100 -+ 6.1 85.5-+ 2.1"
Lung Control Treatment
100 ± 2 4 . 0 94.2 -+ 23.0
100 ± 16.0 80.1 -+ 3.6*
100 ± 19.0 88.9 ± 13.0
100 -+ 18.0 94.6 -+ 3.9
Stomach Control Treatment
100 -+ 6.6 90.4+ - 9.7
100 119
100 120
100 ± 9.8 97.6± 6.3
-+ 8.2 ± 4.4
-+ 9.6 -+ 6.9*
± 11.0 ± 13.0
Mice were restrained in a cold environment (6.5°C) (treatment groups) or housed unrestrained at ambient temperatures (25°C) (control groups) for 1.5 h. Immediately after cold-restraint (0 h), or 1, 2, or 3 h after treatment, mice were sacrificed and tissues harvested for measurement of non-protein sulfhydryl (NPSH) content. Results are expressed as a percent of the mean control NPSH content for each time point + S.E.M., N = 5. Mean control NPSH values (mg/g tissue) ranged from 0.858 to 1.119 for spleen, 1.071 to 1.262 for kidney, 0.393 to 0.453 for heart, 0.575 to 0.762 for lung, and 0.911 to 1.205 for stomach. *Indicates significantly different from control as determined by Student's t-test, P < 0.05.
GSH in response to cold-restraint in the lung appeared to be less than that of other thiols comprising the NPSH pool, the unusually large variability in NPSH levels measured in this tissue make the reliability of this observation uncertain. Discussion The capacity of cold exposure to suppress the hepatic NPSH content of male CAF1 mice was first observed decades ago [8]. In 1953, Bartlett and Register confirmed this finding in rats, and demonstrated that concommitant restraint of the animal by placing it in a wire mesh cylinder enhanced this effect [14]. Simple chilling of rats at 0°C for 2 - 4 h depressed hepatic NPSH concentrations only about 5%, while the addition of restraint resulted in 4-fold greater decreases, i.e. to levels that were 20% lower than those of control animals. We have also observed a marked enhancement of hepatic G S H depression when cold exposure
67
T A B L E III C O M P A R I S O N OF N O N - P R O T E I N S U L F H Y D R Y L A N D G L U T A T H I O N E C O N T E N T OF S E L E C T E D TISSUES F R O M C O N T R O L A N D C O L D - R E S T R A I N E D N D / 4 M I C E
Tissue content (mg/g tissue) Tissue (group)
NPSH
GSH
(GSH/NPSH)
Ratio
Heart Control Treatment
0.540 _+ 0.029 0.396 _+ 0.027
0.264 _+ 0.009 0.20l +_ 0.014
0.49 0.51
Kidney Control Treatment
1.22 _+0.06 0.964 -+ 0.017
0.940 + 0.058 0.704 + 0.009
0.77 0.73
Liver Control Treatment
2.99 + 0.17 2.16 _+ 0.12
2.65 _+ 0.15 1.81 + 0.06
0.89 0.84
0.762 -+ 0.121 0.610 _+ 0.061
0.571 _+ 0.002 0.545 _+ 0.008
0.75 0.89
Lung
Control Treatment
Tissue samples were divided and assayed for non-protein sulfhydryl (NPSH) and reduced glutathione ( G S H ) content as described in Methods. Samples for each tissue were taken from the time of maximal depression of NPSH after cold-restraint: heart, 2 h; kidney, 0 h; liver, 0 h; and lung, I h. Results are expressed as mean _+ S.E.M., N - 5 . The NPSH and GSH content of tissues from treated (coldrestrained) mice were significantly different from the corresponding controls, P < 0.05, in all cases except the GSH values for lung.
and restraint are combined as stressors. In ICR mice, exposures of unrestrained mice to cold (8.5°C) for periods up to 5 h had no effect on hepatic G S H concentrations (unpublished observations) while a 2-h exposure of restrained mice under similar conditions resulted in a significant loss of GSH (15.7% decrease, P < 0.05; Table I). Restraint without cold was an insufficient stressor to produce a change in hepatic GSH concentrations (unpublished observations). Cold-restraint was employed in this study as a stressor, with the hope that the results would be applicable to responses to stressors in general. It was therefore important that the response to cold-restraint not be dependent upon a unique feature of this stressor such as hypothermia. When Beck and Linkenheimer first reported a fall in the hepatic NPSH content of CAF1 mice in mice exposed to cold (5-10°C for 5 h), they noted only a 1-2°C drop in rectal temperatures compared to pre-stress levels [8]. Bartlett and Register [14] had a different experience with cold-restrained ( 2 - 4 h at 0°C) S p r a g u e - D a w l e y rats, and found that a significant decline in hepatic GSH took place in animals whose core temperatures had fallen between 15 and 25°C. In our comparisons of hepatic GSH response to cold-restraint among 4 m o u s e strains, it was apparent that depression of hepatic GSH levels was independent from effects on core body temperature. The N D / 4 mouse was able to sustain core temperature during cold-restraint for
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periods up to 3 h, during which hepatic GSH concentrations declined to approximately 50% of control levels. The GSH depression, at least in the ND/4 mouse, therefore cannot be attributed simply to hypothermia and may reflect a process common to a variety of stressors. The effects of cold-restraint were not confined to the liver, and plasma NPSH concentrations mirrored those in the liver over time following cold-restraint. If subsequent animal experiments bear out this relationship, these observations suggest that plasma GSH or NPSH measurements might serve as a useful, relatively non-invasive means of studying hepatic GSH or NPSH changes in response to stressors in humans. Plasma GSH does not always mimic hepatic GSH changes however, and plasma GSH concentrations in animals subjected to exhaustive exercise or administered oxidant drugs have been observed to remain the same or increase while hepatic levels were diminished [20,21]. These differences in plasma-liver GSH relationships suggest different mechanisms for loss of GSH in the liver. Rather than the oxidation of GSH which appears to occur with exercise or oxidant drug exposure, stress may diminish hepatic GSH through other means such as decreased synthesis or increased intracellular catabolism. Cold-restraint effects on tissues other than liver and plasma were somewhat more variable. The most lasting response was the depression observed in the heart, which showed a delayed decline in NPSH content that persisted through the last time interval examined, 5 h post-treatment. Effects in the kidney were far more transient, with the effects of cold-restraint lasting less than 1 h. Other tissues showed changes that were typically significant only at 1 time point. Though these results suggest that tissues such as the lung may also be susceptible to NPSH depression from cold-restraint, the temporal displacement of the decrease observed in these experiments and their transient nature require that they be interpreted with caution. The approximate 20% increase in NPSH in stomach observed 1 and 2 h after cold-restraint is also worth noting, though variability caused only the rise at 1 h to be statistically signficant. Other studies of the effects of stressors on extra-hepatic NPSH or GSH have been conducted in rats, and the results show some similarities and differences to the changes seen here in mice. For example, Reichard and coworkers observed significant declines in NPSH concentrations in both rat kidney and spleen (as well as liver) 1.5 h after whole body trauma [11], and a significant decrease in renal NPSH concentration has been observed in rats stressed by scalding [9]. Rats subjected to cold-restraint for 8-12 h had a 27% decline in gastric GSH [22]. It appears, through comparisons of N P S H / G S H ratios in tissues from control and cold-restrained mice, that changes in these 2 thiol parameters generally occurred in remarkably parallel fashion. In the case of the liver, this is not surprising since approximately 90% of the NPSH content is GSH. However GSH is only about 50% of the NPSH in the heart, and stress-induced changes in GSH in this tissue appear to be matched by changes in other soluble, non-protein thiols. Equivalent effects on GSH and non-GSH components of total cellular NPSH were also observed in the kidney. These observations suggest that either the mechanism producing the lowering of tissue GSH in response to cold-restraint stress operates equally with other soluble, non-protein (and perhaps protein)
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sulfliydryls, or that GSH is in a relatively rapid equilibrium with other soluble cellular sulfhydryls. In conclusion, application of cold-restraint as a stressor in mice produced a depression in hepatic GSH that was independent of hypothermia and increased in magnitude with increasing duration. Though the effects were generally less pronounced, GSH changes were also observed in other tissues. The extent of loss of GSH was probably not in itself sufficient to cause tissue injury. However, this loss might be sufficient to enhance the toxic insult produced by free radicals, electrophilic xenobiotic metabolites or reactive oxygen species dependent upon GSH for detoxification. Therefore, the results of this study suggest that a stress response may significantly depress cellular GSH/thiol concentrations in the liver and other tissues and thereby render these tissues more susceptible to certain chemical-induced toxicities.
Acknowledgements This research was supported in part by a grant from the National Institutes of Health (ES 05216), and by the Medical Research Endowment Fund, University of Arkansas for Medical Sciences.
References 1 A. Meister, Metabolism and functions of glutathione. TIBS, September (1981) 231. 2 N. Kaplowitz, The regulation of hepatic glutathione. Annu. Rev. Pharmacol. Toxicol,, 25 (1985) 715. 3 J.R. Mitchell, D.J. Jollow, W.Z. Potter, J.R. Gillette and B.B. Brodie, Acetaminophen-induced hepatic necrosis. IV. Protective role of glutathione. J. Pharmacol. Exp. Ther., 187 (1973) 211. 4 D.J. Jollow, J.R. Mitchell, N. Zampaglione and J.R. Gillette, Bromobenzene-induced liver necrosis. Protective role of glutathione and evidence for 3,4-bromobenzene epoxide as the hepatotoxic metabolite. Pharmacology, 11 (1974) 151. 5 P.G. Wells, R.C. Boerth, J.A. Oates and R.D. Harbison, Toxicological enhancement by a combination of drugs which deplete hepatic glutathione: acetaminophen and doxorubicin (Adriamycin). Toxicol. Appl. Pharmacol., 54 (1980) 197. 6 M.A. Evans and R.D. Harbison, Cocaine-induced hepatotoxicity in mice. Toxicol. Appl. Pharmacol., 45 (1978) 739. 7 R.J. Jaeger, R.B. Conolly and S.D. Murphy, Diurnal variation of hepatic glutathione concentration and its correlation with 1,1-dichloroethylene inhalation toxicity in rats. Res. Commun. Chem. Pathol. Pharmacol., 6 (1973) 465. 8 L.V. Beck and W. Linkenheimer, Effects of shock and cold on mouse liver sulfhydryl. Proc. Soc. Exp. Biol. Med., 81 (1952) 291. 9 L.V. Beck, W. Linkenheimer and A. Marraccini, Effects of tumbling trauma, scalding, and hemorrhage on rat tissue non-protein sulfhydryl. Proc. Soc. Exp. Biol. Med., 86 (1954) 823. 10 L.V. Beck and V.D. Rieck, Control mechanisms for trauma induced DLSH reaction (Decrease in mouse liver non-protein sulfhydryl). Proc. Soc. Exp. Biol. Med., 98 (1958) 466. 11 S.M. Reichard, N.M. Bailey and M.J. Galvin, Alterations in tissue glutathione levels following traumatic shock. Adv. Shock Res., 5 (1981) 37. 12 D.A. Brodie and L.S. Valitski, Production of gastric hemorrhage in rats by multiple stresses. Proc. Soc. Exp. Biol. Med., 113 (1963) 998. 13 L. Buchel and D. Gallaiare, Ulcere de contrainte chez le rat. a. Influence, sur la frequence des ulceres, du jeune et de la temperature de l'environment associes a des immobilisations de durees variables. Arch. Sci. Physiol., 21 (1967) 527.
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14 R.G. Bartlett and U.D. Register, Effect of cold and restraint on blood and liver non-protein sulfhydryl compounds. Proc. Soc. Exp. Biol. Med., 83 (1953) 708. 15 L.A. Allison and R.E. Shoup, Dual electrode liquid chromatography detector for thiols and disulfides. Anal. Chem., 55 (1983) 8. 16 G.L. Ellman, Tissue sulfhydryl groups. Arch. Biochem. Biophys., 82 (1959) 70. 17 M. Inoue, Y. Saito, E. Hirata, Y. Morino and S. Nagase, Regulation of redox states of plasma proteins by metabolism and transport of glutathione and related compounds. J. Protein Chem., 6 (1987) 207. 18 N.S. Kosower and E.M. Kosower, The glutathione status of cells. Int. Rev. Cytol., 54 (1978) 109. 19 O.W. Griffith and A. Meister, Glutathione: Interorgan translocation, turnover, and metabolism. Proc. Natl. Acad. Sci. U.S.A., 76 (1979) 5606. 20 H. Lew, S. Pyke and A. Quintanilha, Changes in the glutathione status of plasma, liver and muscle following exhaustive exercise in rats. FEBS Lett., 185 (1985) 262. 21 J.D. Adams, B.H. Lauterberg and J.R. Mitchell, Plasma glutathione disulfide as an index of oxidant stress in vivo: effects of carbon tetrachloride, dimethylnitrosamine, nitrofurantoin, metronidazole, doxorubicin, and diquat. Res. Commun. Chem. Pathol. Pharmacol., 46 (1984) 401. 22 S.C. Boyd, H.A. Sasame and M.R. Boyd, Effects of cold restraint stress on rat gastric and hepatic glutathione: a potential determinant of response to chemical carcinogens. Physiol. Behav., 27 (1981) 377.
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