Serum myoglobin, but not lipopolysaccharides, is predictive of AMPH-induced striatal neurotoxicity

NeuroToxicology 37 (2013) 40–50

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NeuroToxicology

Serum myoglobin, but not lipopolysaccharides, is predictive of AMPH-induced striatal neurotoxicity Mark S. Levi a,1, Ralph E. Patton b, Joseph P. Hanig d, Karen M. Tranter b, Nysia I. George c, Laura P. James e,f, Kelly J. Davis b, John F. Bowyer a,* a

Division of Neurotoxicology, National Center for Toxicological Research, U.S. Food & Drug Administration, Jefferson, AR 72079-9502, USA Toxicologic Pathology Associates, National Center for Toxicological Research, U.S. Food & Drug Administration, Jefferson, AR 72079-9502, USA Division of Bioinformatics and Biostatistics, National Center for Toxicological Research, U.S. Food & Drug Administration, Jefferson, AR 72079-9502, USA d Center for Drug Evaluation and Research, U.S. Food & Drug Administration, Silver Spring, MD 20993, USA e Department of Pediatrics, University of Arkansas for Medical Sciences, 1 Children’s Way, Little Rock, AR 72202, USA f Arkansas Children’s Hospital Research Institute, 13 Children’s Way, Little Rock, AR 72202, USA b c

A R T I C L E I N F O

A B S T R A C T

Article history: Received 18 January 2013 Accepted 5 April 2013 Available online 19 April 2013

Determinants of amphetamine (AMPH)-induced neurotoxicity are poorly understood. The role of lipopolysaccharides (LPS) and organ injury in AMPH-induced neurotoxicity was examined in adult male Sprague-Dawley rats that were give AMPH and became hyperthermic during the exposure. Environmentally-induced hyperthermia (EIH) in the rat was compared to AMPH to determine whether AMPH-induced increases in LPS and peripheral toxicities were solely attributable to hyperthermia. Muscle, liver, and kidney function were determined biochemically at 3 h or 1 day after AMPH or EIH exposure and histopathology at 1 day after treatment. Circulating levels of LPS were monitored (via limulus amoebocyte coagulation assay) during AMPH or EIH exposure. Blood LPS levels were detected in 40–50% of the AMPH and EIH rats, but the presence of LPS in the serum had no effect on organ damage or striatal dopamine depletions (neurotoxicity). In both CR and NCTR rats, serum bound urea nitrogen and creatinine levels increased at 3 h after EIH or AMPH (2- to 3-fold above control) but subsided by 1 day. Alanine transaminase was increased (indicating liver dysfunction) by both AMPH and EIH at 3 h (2- to 10-fold above control) in CR rats, but the levels were not significantly different between the control and AMPH groups in NCTR animals. Mild liver necrosis was detected in 1 of 7 rats examined in the AMPH group and in 1 of 5 rats examined in the EIH group (only NCTR rats were examined). Serum myoglobin increased (indicating muscle damage) in both CR and NCTR rats at 3 h and was more pronounced with AMPH (5-fold above control) than EIH. Our results indicate that: (1) ‘‘free’’ blood borne LPS often increases with EIH and AMPH but may not be necessary for striatal neurotoxicity and CNS immune responses; (2) liver or kidney dysfunction may result from muscle damage; however, it is not sufficient nor necessary to produce, but may exacerbate, neurotoxicity; (3) AMPH-induced serum myoglobin release is a potential biomarker and possibly a factor in AMPH-induced toxicity processes. Published by Elsevier Inc.

Keywords: Amphetamine Neurotoxicity Lipopolysaccharides Muscle Liver Kidney

1. Introduction Abbreviations: ALT, alanine transferase; AST, aspartate transferase; AMPH, Damphetamine; BUN, bound urea nitrogen; CK, creatine kinase; CR, Charles River Laboratory; EIH, envionmentally-induced hyperthermia; LDH, lactate dehydorgenase; RBC, red blood cells; IL-1, interleukin-1; IACUC, Institute Animal Care and Use Committee; LPS, lipopolysaccharides; WBC, white blood cells. * Corresponding author at: NCTR, 3900 NCTR Road, HFT-132, Jefferson, AR 720799502, USA. Tel.: +1 870 543 7194; fax: +1 870 543 7745. E-mail addresses: [email protected] (M.S. Levi), [email protected] (R.E. Patton), [email protected] (J.P. Hanig), [email protected] (K.M. Tranter), [email protected] (N.I. George), [email protected] (L.P. James), [email protected] (K.J. Davis), [email protected] (J.F. Bowyer). 1 Present address: Center for Biologics Evaluation and Research, U.S. Food & Drug Administration, Kensington, MD 20895, USA. 0161-813X/$ – see front matter . Published by Elsevier Inc. http://dx.doi.org/10.1016/j.neuro.2013.04.003

Both amphetamine (AMPH) and methamphetamine (METH) can produce hyperthermia, hyperpyrexia, and hypertension, which are adverse effects on the body that are believed to contribute to the development of thrombotic and hemorrhagic strokes, rhabdomyolysis, neurotoxicity, and death in clinical (Ishigami et al., 2003; Kalant and Kalant, 1975; McGee et al., 2004; Miller and Coon, 2006; Richards et al., 1999; Rothrock et al., 1988; White, 2002; Winslow et al., 2007) and laboratory models of stimulant abuse (Bowyer et al., 1992, 1994, 1998b; Bowyer and Holson, 1995; Broening et al., 1997; Clausing et al., 1995; Fleckenstein et al., 2007; Miller and O’Callaghan, 1994; O’Callaghan and Miller, 1994,

2001; Quinton and Yamamoto, 2006; Seiden et al., 1976; Seiden and Sabol, 1995; Tokunaga et al., 2006). The neurotoxicity of both AMPH and METH are in animal models are dependent on the magnitude of hyperthermia occurring during exposure (Bowyer et al., 1992, 1994, 1998b, 2008; Broening et al., 1997; Clausing and Bowyer, 1999; Eisch et al., 1998; Miller and O’Callaghan, 1994; O’Callaghan and Miller, 1994; Schmued and Bowyer, 1997; Yuan et al., 2006). The animal models yielding these results employ relatively acute exposures (less than 12 h) that can produce life threatening hyperthermia, which subsequently result in extensive dopamine terminal damage in the striatum and sporadic neurodegeneration in the cortex, striatum, and thalamus. This occurs rapidly within 3–12 h after AMPH exposure. Further increases in neurotoxicity are minimal to nonexistent within 24 h of exposure. Both neurodegeneration and long-term striatal dopamine depletions produced by AMPH and METH are nearly identical with respect to dose and magnitude of damage in rodent (Bowyer et al., 1998a, 2008; Cass et al., 1998; Eisch et al., 1998; Levi et al., 2012; Miller and O’Callaghan, 1994; O’Callaghan and Miller, 1994). Although multiple mechanisms have been proposed to explain how pronounced hyperthermia (body temperatures above 40 8C) is involved in the AMPH and METH neurotoxicity process, some have yet to be fully elucidated. Potential mechanisms that contribute to dopamine terminal damage during either AMPH or METH exposure include ROS mediated mitochondrial damage, protein misfolding and Hsp70 induction (Goto et al., 1993; Jayanthi et al., 2004; Lu and Das, 1993; Miller et al., 1991), and the possibility of inappropriate microglial/ immune system activation (Hess et al., 1990; Thomas and Kuhn, 2005; Thomas et al., 2004b; Yamaguchi et al., 1991). The development of hyperthermia may enhance the effects of these mechanisms. Both EIH and AMPH can induce gene expression pattern changes in pathways/systems important in scavenging ROS, minimizing protein damage, and enhancing the recovery of damaged proteins (Thomas et al., 2009, 2010). Additionally, factors such as prolonged stress and extended glucocorticoid exposure can enhance neurotoxicity by either exacerbating hyperthermia (stress) (Kelly et al., 2012) or possibly inappropriately activating the immune system response. It is not clear why AMPH exposure and EIH (to a lesser extent) produce expression patterns in brain and blood, indicating that a pronounced immune response occurs when extreme hyperthermia results {(Kelly et al., 2012; Sriram et al., 2006; Thomas et al., 2009, 2010), Bowyer et al. unpublished data GEO, NCBI; data file GSE29733}. It is known that LPS can interact with METH to alter or normally enhance immune responses of macrophages and microglia in humans and laboratory animals (Bailey et al., 2006; Dunn et al., 2003; Raghubeer and Matches, 1990; Selkirk et al., 2008; Thomas et al., 2004a). However, it is not known whether and how endogenous LPS levels are generated in vivo during METH or AMPH exposure and if they then circulate systemically through blood borne mechanisms. Gram negative bacteria (e.g. Escherichia (E.) coli) residing in the intestine could be a source of LPS. The lysis of bacteria resulting from EIH- or AMPH-induced hyperthermia could release LPS into the systemic circulation. However, this may not be likely due to E. coli’s resistance to elevated temperatures (Raghubeer and Matches, 1990). Alternatively, the intestinal vasculature may become ‘‘leaky’’ and allow entry of bacteria into the blood as this has been seen during heat stress and heat stroke (Bailey et al., 2006; Dunn et al., 2003; Selkirk et al., 2008). Previous studies indicate that administration of an interleukin-1 (IL-1) receptor antagonist prevented METH lethality (Bowyer et al., 1994) suggesting that bacteria or LPS from bacteria may trigger an inflammatory response during amphetamine-related hyperthermia. It is also possible that the muscle damage produced by rhabdomyolysis/malignant hyperthermia can be reproduced by

AMPH and METH (Bowyer and Holson, 1995) and may, in some manner, trigger an immune response. Rhabdomyolysis results in muscle breakdown, which can lead to myoglobin release and is capable of causing renal dysfunction, renal damage, and possibly hepatic injury (Groebler et al., 2011). The present study was initiated to determine whether an acute AMPH or EIH exposure in rats results in increased circulating levels of LPS in the blood, in muscle damage, or in peripheral organ dysfunction, and to determine how this relates to striatal neurotoxicity. Male Sprague-Dawley rats obtained from both the National Center for Toxicological Research (NCTR) breeding colony and Charles River (CR) Laboratories were tested. Systemic blood was obtained through cardiac puncture 3 h or 1 day after exposure so that (1) LPS levels in blood could be determined using the limulus amoebocyte coagulation assay and (2) blood chemistry could be used to indirectly determine dysfunction or damage to skeletal muscle, kidney, liver, heart, and pancreas. Histopathological damage or changes were determined in formalin-perfused animals 1 day post EIH or AMPH. 2. Materials and methods 2.1. Animals Male Sprague-Dawley rats were obtained from CR Laboratories [Crl:CD(SD)] and the NCTR breeding colony animals were originally obtained from Charles River Laboratories in 1974 [Crl: COBS CD (SD) BR Rat]. A total of 55 rats obtained from CR and 65 rats from NCTR were used in the experiments. An explanation for using the two lines of Sprague-Dawley is found in the Discussion. Studies were carried out in accordance with the Declaration of Helsinki and the Guide for the Care and Use of Laboratory Animals as adopted and promulgated by the National Institutes of Health. Approval of the study by the IACUC at NCTR was granted. Prior to testing, rats were housed 2 per cage with food and water available ad libitum. Rats were on a daily 12 h light cycle with lights on at 6:00 am and off at 6:00 pm. During housing, the temperature (23 8C) and humidity (53%) were controlled. The rats were 83–87 days of age at the time of testing. The rats were singly housed for 1 day prior to and during dosing to facilitate behavioral observations. Food and water were returned to the rats 4 h after the end of dosing, after which they were allowed overnight access to 15 g of rat chow and water ad libitum. The food and water restriction ensured that alterations in physiology and gene expression in the AMPH groups were not merely the result of dehydration or fasting. 2.2. Dosing Dosing commenced at 8:00 a.m. and ended at 2:00 p.m. AMPHexposed animals (22 CR and 28 NCTR rats) were given 4 doses of amphetamine comprised of a sequential exposure to 5, 7.5, 10, and 10 mg/kg AMPH subcutaneously with 2 h between each dose. The D-amphetamine-sulfate (Sigma–Aldrich, St. Louis, MO) dose was dissolved in normal saline. The saline control animals received 4 injections of 1 ml/kg. The EIH groups were also given 4 doses of 1 ml/kg normal saline in a 39–40 8C environment. Rats that were unlikely to survive AMPH exposure (5 CR and 4 NCTR rats) or EIH (3 CR and 4 NCTR rats) were removed from the study. Animals were deemed not likely to survive if they had (1) signs of hind limb paralysis, (2) persistent body tremors or myoclonus, or (3) respiratory distress and/or body temperatures below 35.0 8C. In all groups, the behavior and body temperature of each rat were monitored hourly during saline or AMPH exposure and monitoring continued until at least 3 h after the last dose. All animals in the EIH or AMPH groups had pronounced hyperthermia except for one CR rat sacrificed at the 3 h time point (see supplementary table). The

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M.S. Levi et al. / NeuroToxicology 37 (2013) 40–50

lethal effects of hyperthermia and hyperpyrexia in the AMPH and EIH groups (when body temperatures exceeded 41.4 8C) were prevented by placing the animals unrestrained on crushed ice for 15–30 min in a clean, wood chip free cage to allow their temperatures to drop below 40.0 8C. Animals in the EIH group were not placed back into the hot environment until they regained mobility. 2.3. Animal sacrifice and tissue harvest Rats were sacrificed at 3 h or 1 day (22 h after the 4th dose) for blood chemistry and LPS analyses, striatal dopamine, 5-HT and metabolite levels, and/or histological analysis. All rats were administered 300 mg/kg of pentobarbital by intraperitoneal injection. Upon loss of consciousness and response to pain (tail pinch), 5 ml of blood was slowly withdrawn via cardiac puncture using an 18 gauge needle for blood chemistry and LPS analysis. The blood was aliquoted into 3 fractions. Five hundred microliters of whole blood was frozen immediately on dry ice and stored at 80 8C. Three hundred microliters was added to a heparin-coated blood storage tube and inverted 5 times to mix the sample for determining WBC and RBC counts. The remaining (3–4 ml) was added to a BD Red Top SST Vacutainer1. This sample was allowed to clot and the serum for blood chemistry analysis was obtained by centrifugation, 500 ml of serum was added to an Eppendorff tube and stored at 4 8C while the remaining serum was placed in an Eppendorff tube and frozen at 20 8C. Rats were then processed for determining striatal dopamine, 5-HT, metabolite levels (NCTR and Charles River animals), or histological analysis (NCTR rats). Animals used for histological evaluation of heart, kidney, and liver were prepared for fixation after respiration ceased (see below for details). Animals used strictly for determining only the striatal dopamine and 5-HT and metabolite levels (all the CR and most of the NCTR rats) were decapitated immediately after blood withdrawal. For animals whose brain and internal organs were harvested, once decapitated, the carotid arteries were clamped off and the animals were perfused as described in Section 2.5. 2.4. Analysis of dopamine, serotonin and metabolite levels in striatum The striatum was harvested by removing the surrounding parietal and frontal skull bones and dissecting away the dura mater. The brains were then rapidly but carefully removed without damaging the meninges and chilled in ice-cold normal saline for 5 min. After the meninges and associated vasculature were harvested from the cerebrum, the parietal cortex and striatum were harvested. These tissues were immediately frozen on dry ice and stored at 70 8C. The harvested striata were used to determine dopamine and 5-HT and metabolite levels. One striatum from each animal was placed in ice-cold 0.2 N perchloric acid pH  1.0 (10 buffer volume to tissue wt.) and homogenized by ultrasonication for 15–20 s. The resulting homogenate was centrifuged at 12,000  g for 10 min at 5 8C. One hundred microliters of the resulting supernatant was pipetted into a costar1 Spin-X Tube filter (0.45 mm) and centrifuged at 2000  g for 10 min at 5 8C. The filtrate was removed and diluted 3-fold with 200 ml of HPLC buffer just prior to high pressure liquid chromatography (HPLC). HPLC and electrochemical detection methods were used for quantifying dopamine, 5-HT and metabolite levels as described by Stephans with modifications (Stephans and Yamamoto, 1994). HPLC retention times were as follows: 5.05 min (DOPAC), 7.05 min (dopamine), 8.55 min (5-HIAA), 14.40 min (HVA), 17.44 min (5HT) and 19.65 min (3-methoxytyramine). Tissue preparation and HPLC analysis was completed within 4 h for each sample to avoid the loss of the 5-HT and 5-HIAA signal.

2.5. Histological evaluation of heart, liver, and kidney After blood collection, the body cavity was opened and the animals were perfused through the left ventricle and aorta with 70 ml normal saline (25 ml/min) followed by 300 ml (30 ml/min) of 4% formaldehyde in 0.1 M sodium phosphate buffer (pH 7.4). The perfusion-fixed heart, kidneys, and liver were collected and immersed in 4% formaldehyde in 0.1 M sodium phosphate buffer (pH 7.4). The collected tissues were trimmed, routinely processed, embedded in paraffin-based infiltrating media, sectioned at approximately 5 mm, stained with hematoxylin and eosin, and examined by light microscopy. Lesions were graded as 1 (minimal), 2 (mild), 3 (moderate), or 4 (marked). 2.6. Serum chemistry analysis Blood cell counts were determined in whole blood collected in EDTA. White blood cell (WBC) counts and red blood cell (RBC) counts were determined on an ABX Pentra 60 C+ analyzer (ABX, Irvine CA). Maintenance and calibration was done according to the manufacturer’s recommendations. Three levels of assayed controls were included in daily analyses as internal controls. Blood chemistry methods used to determine lactate dehydrogenase (LDH), creatinine kinase (CK), aspartate serum transaminase (AST), alanine transaminase (ALT), creatinine, bound urea nitrogen (BUN), and myoglobin were performed on serum. Clinical chemistry analyses were conducted on an Alfa Wassermann ALERA (West Caldwell, NJ). Alfa Wassermann reagents were used for ALT, AST, CK, LDH, BUN, creatinine, amylase, and lipase. Pointe Scientific, Inc. (Canton, MI) reagents were used for CK-MB. All maintenance was done according to the manufacturer’s recommendations. The instrument was calibrated daily and 2 levels of assayed controls were included in daily analyses as internal controls. 2.7. LPS analysis LPS levels were determined from frozen blood samples stored at 70 8C since sacrifice. Endotoxin was measured by a limulus amebocyte lysate (LAL) assay using the commercially available chromogenic LAL endpoint QCL 1000 Kit from Lonza (Walkersville, MD) according to the manufacturer’s instructions. Inhibiting and enhancing components were eliminated by diluting the sample 1:10 in pyrogen-free water and heating at 75 8C for 10 min before the assay was performed. The plates were read on an ELx800 Universal Microplate Reader (Bio-Tek, Winooski, VT). A standard curve using the manufactured supplied standard was run with each batch of samples and results were calculated by the reader’s software. All maintenance was done according to the manufacturer’s recommendations. 2.8. Statistical analysis Tests of statistical difference between treatment groups for serum ALT levels, striatal dopamine levels, and myoglobin levels were performed using a two-way analysis of variance (ANOVA) (treatment  sacrificial time point) followed by contrast analysis. Analyses were carried out using the GLM procedure of SAS 9.2. Pearson’s correlation coefficient was used to examine the linear association between various biochemistry parameters e.g. temperature and ALT, myoglobin and striatal dopamine, etc. The blood LPS levels varied greatly (from 0.2 to 10 units) between animals; see Supplementary Table 1 for individual animal LPS values. Animals with detected levels of LPS were classified as LPS+ and animals with undetectable LPS levels (0.1 units) were classified as LPS . The nonparametric Mann–Whitney was used to evaluate the

significance of biochemistry parameters between LPS+ and LPS animals. Correlation and the Mann–Whitney test were performed using R statistical software (http://www.r-project.org). P-values less than 0.05 were deemed statistically significant unless otherwise noted. Separate analyses were carried out for NCTR and CR animals due to the experimental design. Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.neuro.2013.04.003. 3. Results 3.1. Body temperatures during EIH and AMPH exposure All saline control animals survived dosing and were sacrificed either 3 h or 1 day after the 4th dose. Sixteen of the 19 rats from CR and 15 of 19 rats from NCTR in the EIH groups that were deemed likely to survive and recover were sacrificed at either the 3 h or 1 day time points. Seventeen of the 22 rats from CR and 24 of 28 rats from NCTR in the AMPH groups, also deemed likely to survive and recover, were sacrificed at either the 3 h or 1 day time point. Data from the animals determined not likely to survive and recover due to EIH or AMPH were excluded from the study. Temperature profiles for the NCTR and CR groups sacrificed 3 h post exposure are shown in Fig. 1. The overall hyperthermic profile of the EIH groups for CR and NCTR were very similar. Body temperature for the CR EIH group tended to be lower toward the end of exposure because many did not regain their full motor activity (exhibited some ataxia) and were not put back into the hot environment at the later time points. With respect to overall behavior, the CR EIH group appeared more adversely affected than the NCTR EIH group. Although body temperatures of the NCTR and CR groups exposed to AMPH were very similar after the 3 h time point, AMPH increased body temperature more after the 1st and 2nd dose (within the first 3 h) in the CR group. In previous studies we used a room temperature of 23.5–24.5 8C, which is slightly above the normal housing temperature of 22–23 8C, to ensure that the neurotoxicity was maximized in NCTR animals. We intentionally reduced the environmental temperature during AMPH exposure in the present study to get a broader range of neurotoxicity (dopamine depletions). Unexpectedly, AMPH increased body temperature to a much greater extent than expected after the

Fig. 1. Body temperature profiles resulting from saline, EIH, or AMPH exposure. A time course of the mean core body temperatures (S.E.M.) are presented for both the NCTR and CR Sprague-Dawley rats during control, EIH, or AMPH exposure for animals sacrificed 3 h after the end of dosing. The means with S.E.M. for each group are shown. The number of NCTR rats per group was: control, n = 9; EIH, n = 7; AMPH, n = 9. There were n = 9 rats in each CR treatment group. * Indicates time at which saline, EIH, or AMPH was injected.

1st and 2nd dose (within the first 3 h of treatment) in the CR group compared to the NCTR group. 3.2. Dopamine depletions and hyperthermic profiles The striatal dopamine levels (and metabolites Supplementary Figure 1) of the various treatments for both NCTR and CR rats at 3 h and 1 day after exposure are shown in Supplementary Table 1 and Fig. 2. Striatal dopamine and metabolite levels were not depleted by EIH compared to controls in either NCTR or CR rats (shown in Tables 1a and 1b and Supplementary Figure 1). Interestingly, the CR rats in the 3 h and 1 day control and EIH groups had lower striatal dopamine levels than the NCTR rats. However, at the 3 h and 1 day time points, both the NCTR and CR groups exposed to AMPH had pronounced and significant striatal dopamine decreases (Fig. 2). Although the peak body temperatures of CR rats exposed to AMPH were not significantly different from the NCTR AMPH groups (Tables 1a and 1b), the CR AMPH groups had higher body temperatures for the first 3 h after the start of dosing/exposure (Fig. 1). The early occurrence of body temperatures above 40 8C in the CR rats may explain why the 3 h striatal dopamine levels are less than the NCTR AMPH group since this is at a relatively early time point in AMPH-induced striatal damage. Striatal dopamine decreases of 80% in NCTR and 90% in CR were seen in rats 1 day after AMPH Supplementary Figure 1. However, these levels would have been a bit higher (less dopamine depletion) if the animals had been sacrificed 1 week after AMPH. Nonetheless, the dopamine depletions would have still been very pronounced at 1 week post AMPH exposure (75% for NCTR and 80% for CR, as extrapolated from unpublished data from Bowyer et al.). At 1 day after AMPH the 5-HT levels were also somewhat decreased with striatal levels at about 60% of control Supplementary Figure 2.

Fig. 2. Scatterplot of peak core body temperature and striatal dopamine. Peak core body temperatures and striatal dopamine levels are plotted for individual rats given AMPH and sacrificed either 3 h or 1 day after exposure for NCTR and CR SpragueDawley rats.

M.S. Levi et al. / NeuroToxicology 37 (2013) 40–50

44 Table 1a Serum and brain chemistry analysis. Time point

Treatment

3h

Control

Rat source

n

Peak body temp. (8C)

Striatal dopaminea

ALT (units/L)

BUN (mg/dL)

Myoglobin (ng/dL)

White blood cells (103/mm3)

Red blood cells (106/mm3)

NCTR Charles River NCTR Charles River NCTR Charles River

8 9 7 10 9 9 8b

38.2  0.7 38.1  0.4 41.9  0.9 42.4  0.5 41.4  0.4 41.6  0.8 41.8  0.3

1254  53.6 925  55.9 1224  70.8 999  58.9 484  92.6d,e 228  58.9d,e 171  19.1

72  10.7 60  7.6 47  8.2 1008  543.4c 111  14.9 212  63.5e 227  22.5

14  0.9 16  1.6 26  2.2 32  3.4c 38  5.5d 42  6.4d,e 45  2.3

107  21.9 110  42.0 173  108.6 206  42.0 504  130.8d,e 556  75.4d,e 602  67.5

8  1.0 7  0.9 7  1.6 7  1.1 10  1.9 8  1.4

(n = 8) (n = 8) (n = 6) (n = 9) (n = 8) (n = 5)

8  0.4 8  0.2 10  0.8 9  0.6 10  0.6 10  0.3

(n = 8) (n = 8) (n = 6)c (n = 9)c (n = 8) (n = 5)d

NCTR Charles River NCTR Charles River NCTR Charles River

8 5 8 6 16 8

37.8  0.5 38.4  0.3 41.8  0.9 42.0  0.3 41.5  0.4 41.9  0.2

1263  69.1 1006  28.5 1177  50.2 1007  44.3 261  61.2d,e 78  22.3d,e

68  14.9 47  5.9 56  6.6 180  124.8 128  35.4 116  16.7e

18  0.8 16  0.8 20  1.5 20  2.3 32  8.9 19  0.6

102  31.7 63  21.2 94  28.7 194  134.9 230  102.9 273  119.2

9  1.8 11  2.7 8  1.7 8  1.5 8  0.5 5  0.8

(n = 6) (n = 5) (n = 7) (n = 5) (n = 15) (n = 7)d

9  0.4 8  0.1 9  0.5 8  0.2 9  0.3 9  0.4

(n = 6) (n = 5) (n = 7) (n = 5) (n = 15) (n = 7)

group

EIH AMPH

1 day

Control EIH AMPH

Values are given as mean  standard error of the mean (SEM). Alanine transaminase (ALT), bound urea nitrogen (BUN), creatinine, creatine kinase (CK), lactate dehydrogenase (LDH), and myoglobin were determined in serum. Note that for white and red blood cell counts, the n values were different compared to the rest of the parameters listed in the table. a Dopamine levels are listed in ng/100 mg striatal tissue. b Indicates that the one CR rat with minimal hyperthermic response to AMPH was removed from mean and SEM calculations. c Indicates EIH group significantly differs from control. d Indicates AMPH group significantly differs from control. e Indicates AMPH group significantly differs from EIH.

Table 1b Serum chemistry analysis continued. Time point

Treatment group

Rat source

n

LDH (units/L)

CK (units/L)

Creatinine (mg/dL)

Amylase (units/L)

Lipase (units/L)

3h

Control

NCTR Charles River NCTR Charles River NCTR Charles River

8 9 7 10 9 9 8a

1339  224.2 1149  155.0 1420  97.8 11,451  6376.2b 2213  553.3 4631  1509.8 5653  1176.3

1084  123.5 1003  280.4 2114  974.5 3390  1088.2b 2286  352.4 5534  1044.0c,d 5650  1176.3

0.29  0.04 0.30  0.03 0.53  0.07b 0.73  0.17b 0.67  0.08c 0.83  0.10c 0.86  0.11

792  40.0 (n = 8) 1098  80.4 (n = 9) 1322  218.0 (n = 7) 2050  337.1 (n = 10)b 1177  122.6 (n = 9) 1264  78.9 (n = 9)d

19  3.2 20  5.3 24  1.3 33  6.5 33  5.2 18  4.1

(n = 8) (n = 9) (n = 7) (n = 10) (n = 9)c (n = 7)d

NCTR Charles River NCTR Charles River NCTR Charles River

8 5 8 6 16 8

1368  199.1 923  95.0 1651  264.4 1176  371.2 2028  359.9 1317  112.8

1389  271.5 1555  219.6 966  181.5 1407  449.4 2264  609.9 1386  223.0

0.32  0.02 0.39  0.04 0.38  0.07 0.45  0.07 0.34  0.04 0.29  0.04

813  44.9 (n = 7) 884  117.2 (n = 5) 971  46.9 (n = 7) 955  84.6 (n = 6) 1167  281.1 (n = 15) 958  157.8 (n = 8)

27  3.9 25  2.4 29  2.5 18  3.3 28  1.5 23  5.9

(n = 7) (n = 5) (n = 7) (n = 6) (n = 15) (n = 8)

EIH AMPH

1 day

Control EIH AMPH

Values are given as mean  standard error of the mean (SEM). Creatinine, creatine kinase (CK), lactate dehydrogenase (LDH), amylase, and lipase were determined in serum. a Indicates that the one CR rat with minimal hyperthermic response to AMPH was removed from mean and SEM calculations. b Indicates EIH group significantly differs from control. c Indicates AMPH group significantly differs from control. d Indicates AMPH group significantly differs from EIH.

Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.neuro.2013.04.003. 3.3. Blood chemistry affected by EIH and AMPH The blood chemistry parameters most affected by EIH and AMPH (compared to control) are summarized in Tables 1a and 1b. Blood chemistry, blood cell counts, and all other parameters measured in this study are reported for each animal in Supplementary Table 1. Figs. 1–7 are presented to facilitate an understanding of how EIH and AMPH affect selected blood chemistry parameters and LPS from blood and to determine how these measures relate to striatal neurotoxicity, as determined by dopamine depletions. In general, with the exception of LPS levels, EIH and AMPH had much greater effects on the parameters measured at 3 h than at 1 day. The serum ALT levels at 3 h post exposure are plotted against core body temperature in Fig. 3 to enable a visual inspection of whether body temperature correlates with liver

injury (represented by serum ALT) in the CR and NCTR rats exposed to either EIH or AMPH. Inspection of Fig. 3 also shows that the ALT levels in CR rats were affected to a much greater extent by both AMPH and EIH than the NCTR animals. There was a non-significant, weak, and positive correlation between body temperature and ALT in the CR rats exposed to AMPH (r = 0.30) and a significant, strong, and positive correlation (r = 0.87; p = 0.00) for NCTR rats. However, with respect to EIH exposure, neither CR nor NCTR rats had a statistically significant correlation between ALT and body temperature. The relationship between ALT and striatal dopamine levels at 3 h for control, EIH, and AMPH in both NCTR rats (top panel) and CR rats (bottom panel) is shown in Fig. 4. In regard to NCTR rats, there was a significant strong negative correlation (r = 0.84; p = 0.01) between ALT and dopamine levels at 3 h post AMPH only. However, unexpectedly, there was not a statistically significant difference in ALT levels between the NCTR control and AMPH groups. Furthermore, there was very little correlation (r = 0.13) between ALT and dopamine levels in the NCTR control group (Fig. 4

Fig. 3. Scatterplot of peak core body temperature with serum ALT from EIH or AMPH exposure. Peak core body temperatures and serum ALT levels are plotted for individual rats exposed to EIH or AMPH at the 3 h time point. Fig. 5. Scatterplot of serum BUN levels versus striatal dopamine levels after EIH or AMPH exposure at 3 h after dosing. Serum BUN and striatal dopamine levels for individual animals are plotted for both NCTR (top graph) and CR Sprague-Dawley (bottom graph) rats for the three treatment groups at the 3 h time point.

and Tables 1a and 1b). In stark contrast to the NCTR animals, in both the CR AMPH and CR EIH groups, there were big increases in ALT in many of the animals compared to controls (CR AMPH and EIH were significantly different). Changes in serum AST paralleled the ALT changes seen within treatment groups and within animals for both NCTR and CR groups (Tables 1a and 1b). The relationship between BUN levels and striatal dopamine at 3 h for control, EIH, and AMPH exposure groups for NCTR (top graph) and CR (bottom graph) is shown in Fig. 5 and Tables 1a and 1b. Changes in BUN levels correlated negatively (r = 0.68; p = 0.06) with dopamine levels in the NCTR animals given AMPH, but not in the control (r = 0.04) or EIH (r = 0.05). There was a non-significant correlation between BUN and dopamine levels in the CR EIH and AMPH groups. BUN levels were positively correlated with dopamine levels for the control group at 3 h (r = 0.66; p = 0.08). The serum myoglobin levels in the EIH group was about 60% higher than controls for both NCTR and CR animals at 3 h, but this increase was not statistically significant. Additionally, there was not a significant correlation between myoglobin and dopamine levels in the control and EIH groups for NCTR or CR (Supplementary Figure 1, Fig. 6 and Tables 1a and 1b). Myoglobin levels were significantly different at 3 h between AMPH and control and AMPH and EIH for NCTR and CR. Also, there was a significant negative correlation between myoglobin and striatal dopamine levels in the NCTR (r = 0.78; p = 0.02) and CR (r = 0.86; p = 0.01) AMPH animals. Fig. 4. Scatterplot of serum ALT levels versus striatal dopamine levels after EIH or AMPH exposure at 3 h after dosing. Serum ALT and striatal dopamine levels for individual animals are plotted for both NCTR (top graph) and CR Sprague-Dawley (bottom graph) rats for the three treatment groups at the 3 h time point.

3.4. Leukocyte and LPS levels after EIH and AMPH RBC count seemed to increase at the 3 h time point after EIH and AMPH in both NCTR and CR animals when compared to controls

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M.S. Levi et al. / NeuroToxicology 37 (2013) 40–50 Table 2 Incidence of occurrence of LPS in blood after EIH or AMPH. Treatment group

Rat source

Control

NCTR Charles River NCTR Charles River NCTR Charles River

EIH AMPH

3 h post exposure

1 day post exposure

Number tested

Number LPS+

Number tested

Number LPS+

8 9 7 10 9 9

0 2 2 5 5 4

6 5 7 6 15 8

0 1 2 2 7 3

Detectable levels of LPS presence varied more than 50-fold among the animals. Levels ranged from just above a threshold of 0.2 to over 10.5 EU/ml. See Section 2 for details on measurement of LPS levels and Supplementary Table 1 for details on the variability of LPS levels among individual rats.

Fig. 6. Scatterplot of serum myoglobin levels versus striatal dopamine levels after EIH or AMPH exposure at 3 h after dosing. Serum myoglobin and striatal dopamine levels for individual animals are plotted for both NCTR (top graph) and CR SpragueDawley (bottom graph) rats for the three treatment groups at the 3 h time point.

(possibly due to dehydration during exposure). The increases reached statistical significance in NCTR and CR animals for EIH and also in CR animals for AMPH. The WBC count in blood was not affected by EIH or AMPH in NCTR animals while WBC counts in the CR AMPH rats were only affected at the 1 day time point, where they unexpectedly decreased (Table 2). The LPS levels in blood for the animals tested are summarized in Table 2 and the individual animal levels of LPS can be found in Supplementary Table 1. The levels of detected LPS ranged widely among the animals that were LPS+; LPS levels ranged from 0.2 EU/ml to 10.5 EU/ml. The presence or absence of LPS did not indicate the severity of ALT or BUN increases in either CR or NCTR AMPH groups at either 3 h or 1 day (data not shown). As can be seen in Fig. 7 the presence of LPS also did not coincide with either the severity of dopamine decreases or myoglobin increases in the NCTR and CR AMPH groups at either 3 h or 1 day (Fig. 8).

Fig. 7. The presence of LPS in serum does not potentiate the striatal dopamine depletions or serum myoglobin increases produced by AMPH exposure. Striatal dopamine levels versus serum myoglobin are plotted for both NCTR and CR Sprague-Dawley at the 3 h (left panel) and 1 day (right panel) time points for rats exposed to AMPH. A plus sign inside the symbol indicates that LPS was detected in the animal’s serum.

with marked glycogen depletion. Interestingly, this animal also had 95% striatal dopamine depletion, greatly elevated levels of ALT, BUN, and myoglobin, but no detectable LPS levels in the blood (see Supplemental Table 1). The animal in the AMPH group that had marked glycogen depletion in the liver, normal ALT levels, and no detectable LPS levels, but its BUN and myoglobin levels were elevated above the 1 day saline controls at 1.7- and 4.0-fold, respectively. The EIH animal with mild liver necrosis had nearly normal ALT, BUN, and myoglobin levels, but had high blood levels of LPS (5.4 EU/ml) and did not have glycogen depletion in the liver (Supplemental Table 1). Unfortunately, blood analysis for the EIH animal with marked glycogen depletion in the liver could not be determined. Although the EIH animal with mild liver necrosis and the EIH animal with glycogen depletion in the liver were clearly very hyperthermic (Tables 1a and 1b), their peak body temperatures were not as high as the other animals in this EIH group, which was very unexpected. 4. Discussion

Fig. 8. Histological evaluation of liver toxicity produced by AMPH and EIH. Panel A (upper) shows a normal (control) liver section stained with hematoxylin & eosin and Panel B (lower) shows a section of liver stained with hematoxylin & eosin from the one animal (out of 7) with necrosis and glycogen depletion attributed to AMPH treatment. The one animal from the five EIH animals that had mild necrosis is not shown but its liver necrosis was similar to that seen in Panel B. Panel A: The green arrow in the center is pointing at a central vein. There are several other central veins present that are not labeled because they are difficult to visualize due to the attenuation of sinusoids and central veins by hepatocytes filled with glycogen (represented by irregularly shaped clear cytoplasmic spaces). The black arrow is pointing at a hepatic portal vein in a portal triad. Panel B: The red arrows point to areas of centrilobular hepatocellular necrosis. Inflammatory cell infiltrates are present within the areas of centrilobular necrosis. A diffuse depletion of glycogen from the cytoplasm of hepatocytes is also present (compare to Panel A). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

3.5. Histological evaluation of heart, liver, and kidney after EIH and AMPH Histological analysis of heart, liver, and kidney of the controls (n = 5) was compared to EIH (n = 5) and AMPH (n = 7) groups in NCTR rats 1 day after exposure. There was no histological evidence that either the heart or kidney was damaged by EIH or AMPH. Some histological effects were noted in the liver after both EIH and AMPH exposure. Two of 7 animals in the AMPH group (designated JB15 and JB25 in Supplemental Table 1) and 1 of 5 animals in the EIH group (designated JB23 in Supplemental Table 1) had marked glycogen depletion in the liver. The glycogen depletion involved all hepatocytes in all zones of the hepatic lobule. One animal in the EIH group (designated JB24 Supplemental Table 1) and one animal in the AMPH group (designated JB15 Supplemental Table 1) had mild coagulative necrosis of hepatocytes surrounding or adjacent to central/sublobular veins. Cellular infiltrates of macrophages and/or polymorphonuclear cells were often present in the areas of coagulative necrosis. The animal in the AMPH group that had mild liver necrosis was also one of the two animals in the AMPH group

The results of this work show that although increases in blood/ LPS occurred in about 50% of the animals during EIH or AMPH exposure, LPS presence in blood does not appear to be necessary for the occurrence of neurotoxicity and muscle, liver, or kidney damage. Additionally, since LPS was only detected in 50% of the AMPH animals, it also may not be linked to the activation of genes related to immune system alterations/activation that is normally seen in brain, meninges, and blood in almost all animals exposed to the AMPH regimen used in our studies (Thomas et al., 2009, 2010). Our studies also indicate that heart, kidney, and liver damage are not necessary for striatal neurotoxicity but could exacerbate it. The occurrence of myoglobin in blood was much more pronounced in both the NCTR and CR AMPH groups than in the control and EIH groups and it cannot not be discounted as a factor involved in somehow triggering or exacerbating vascular or striatal neurotoxicity. However, this is only a correlative finding and not a causal/ mechanistic one. Nonetheless, significantly increased serum myoglobin has the potential of being a biomarker for striatal neurotoxicity after AMPH or METH exposure and may persist for up to 1 day after exposure. 4.1. Comparisons between NCTR and Charles River animal responses to EIH and AMPH CR Sprague-Dawley rats were used in these experiments as well as the NCTR line so that the neurotoxicity and other somatic toxicities of EIH and AMPH could be roughly compared between the two animal lines. We have previously published our work using the NCTR line of Sprague-Dawley rats. The NCTR line was originally started from Charles River rats obtained in 1974 but has been bred in isolation since then. Thus, it was not certain prior to this study how similarly the NCTR strain responded to toxic insults of AMPH or EIH compared to Sprague-Dawley rats from commercial vendors. Our results clearly show that both lines show similar overall toxicity profiles. A comparison of how AMPH and EIH affected striatal dopamine levels, serum ALT, and serum myoglobin is presented in Supplementary Figure 1. However, direct statistical comparisons of the treatment effects on biochemistry parameters and muscle toxicity could not be made between the two lines because animals from both lines were not ‘‘same-day’’ tested and the CR rats’ hyperthermia was somewhat more pronounced. Testing both the NCTR and CR lines of rat enabled us to determine that (1) systemic organ damage/impairment in the absence of AMPH does not produce striatal damage (CR line) and (2) that liver, kidney and, heart damage are not necessary, but may potentiate striatal damage (NCTR line).

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4.2. Origin of LPS in blood, immune system activation, and toxicity The origin of the LPS in blood in our studies is not clear. Gram negative bacteria, such as Escherichia coli (E. coli) present in the intestine during AMPH exposure or EIH could have been a source of LPS if they were killed/lysed by greatly elevated body temperatures produced by EIH or AMPH. However, this appears to be an unlikely event (Raghubeer and Matches, 1990). Alternatively, the intestinal vasculature may become ‘‘leaky’’ and allow the entry of bacteria into the blood which apparently can occur in humans and laboratory animals during heat stress, heat stroke, or stress (Bailey et al., 2006; Dunn et al., 2003; Selkirk et al., 2008). It is possible that the mechanical irritation to the colon due to temperature monitoring was sufficient to have produced some intestinal vasculature permeability in our studies in a few of the CR, but not the NCTR, rats. The explanation for this difference is not readily apparent. In turn, it is possible the intestinal leakiness is significantly exacerbated when this mechanical disturbance is coupled with the extreme hyperthermia produced by EIH or AMPH. Studies determining whether the intensity of mechanical trauma to the colon under control conditions would be necessary to resolve the question of whether measurements of temperature via rectal thermometers could result in intestinal bacteria entering the circulating blood. In any event, if such trauma occurred, it had no effect on neurotoxicity because LPS in serum had no effect on neurotoxicity. Whatever the source, the LPS detected in circulating blood in our studies was only seen in approximately 50% of the CR and NCTR animals exposed to AMPH. However, the measure of striatal, muscle (myoglobin), liver (ALT), and kidney (BUN) toxicity was not significantly different in animals when LPS was present in circulating blood of the AMPH groups. Additionally, pronounced dopamine depletions were not dependent on presence or absence of LPS in blood. Thus, striatal neurotoxicity is probably not dependent on LPS in blood. The presence of LPS in blood also had no effect on the circulating levels of leukocytes at 3 h or 1 day after either EIH or AMPH. However, we did not find that the overall WBC count significantly changed with either EIH or AMPH compared to controls at the 3 h time point. This was unexpected because previous data indicated that gene expression and other immune-related data showed alterations in the mRNA and protein levels of genes related to immune function in brain and the meninges and associated vasculature (Hess et al., 1990; Kelly et al., 2012; O’Callaghan and Miller, 1994; Thomas et al., 2004a,b, 2009, 2010; Thomas and Kuhn, 2005; Yamaguchi et al., 1991). Changes in mRNA levels of immuno-related genes in circulating blood are present at 3 h and 1 day, particularly those related to macrophages, specific T-cells and/ or possibly vasculature such as IL-1b, Cd8a, Cxcr2, Itgam, and Tnfrsf1a (unpublished data in GEO, NCBI; data file GSE29733). However, these changes in brain and associated vasculature may be triggered by damage to the neurons and vasculature present and not related to alterations/activation of the innate immune system blood of laboratory animals. The significant decrease in the WBC counts at 1 day in the CR animals given AMPH was unexpected. 4.3. Liver and kidney damage or dysfunction in relation to striatal neurotoxicity In NCTR rats, AMPH and EIH exposure induced histopathologically detectable damage at 1 day in only 2 animals. Mild liver damage was only observed in 1 animal and/or significant glycogen depletion was only seen in 2 of the 7 AMPH animals. Pronounced (70–80% depletions) long-term dopamine depletions would be expected from the animals’ hyperthermic profiles and/or from the 1 day striatal dopamine levels observed in all 7 rats in the AMPH

group. LPS was not detected in the blood of rats with mild liver damage 1 day after either AMPH or EIH. Thus, histopathological damage is not necessary for striatal neurotoxicity, and LPS presence in the blood is probably not necessary for AMPH- or EIH-induced liver histopathology. Histological analysis of the CR animals was not performed, but it is unlikely that liver necrosis was present in the majority of them. Six of the 9 CR animals had ALT levels within the NCTR range at the 3 h time point and there was no apparent difference between the ALT levels in CR and NCTR rats given AMPH at the 1 day time point. However, two of the 13 NCTR rats evaluated had ALT serum levels over 300 U/L indicating the possibility of liver damage. Also, all of the CR rats sacrificed 1 day after treatment had ALT levels that were 50% or lower than those seen in the one NCTR animal that had mild to moderate liver necrosis. Thus, it is unlikely that our Charles River animals sustained more permanent liver toxicity (necrosis) than the NCTR animals in our experiment. Our results show that the extent of ALT and BUN elevations in the AMPH and EIH groups was comparable in the CR line. Since the EIH group did not have striatal neurotoxicity (dopamine or metabolite depletions), it is plausible to conclude that that alterations in liver and kidney function related to elevated serum levels of ALT and BUN do not trigger striatal neurotoxicity. As well, liver dysfunction is not necessary for striatal neurotoxicity since ALT levels in the NCTR group given AMPH were not different than the respective controls. With regard to kidney dysfunction and striatal neurotoxicity, our results are not as clear. It is not believed that elevated BUN levels cause striatal neurotoxicity because in both the NCTR and CR line, EIH significantly elevates BUN levels without producing striatal neurotoxicity. There is some overlap in BUN levels between a few animals in the AMPH and control groups in both lines, but the BUN levels in the AMPH group significantly differed from control in both NCTR and CR animals. Overall, there was no compelling evidence indicating that liver dysfunction is necessary for striatal neurotoxicity. However, this does not rule out the possibility that liver damage exacerbates striatal neurotoxicity since the animals with greatest dopamine depletions had elevated ALT and BUN levels. It has been reported by others (Halpin and Yamamoto, 2012) that liver damage may result in increased levels of ammonia in blood and that this may be necessary for striatal neurotoxicity. However, in those studies a significantly greater necrosis occurred, which did not happen in our study. Thus, in the present study, we do not expect that significant increases in ammonia levels occurred due to significant long-lasting liver damage (e.g. liver necrosis) since the ALT levels in over 80% of the NCTR and CR animals were within normal range 1 day after AMPH. 4.4. Serum myoglobin levels relevance to striatal neurotoxicity and immune response It is clear (as can be visualized in Supplemental Figure 1 and the data analysis in Section 3) that elevated myoglobin correlates much more closely with dopamine depletions than ALT. It is unlikely that myoglobin breakdown would lead to elevated levels of ammonia in blood or liver toxicity. However, the myoglobin breakdown product, ferrihemate, is nephrotoxic and has been linked to rhabdomyolysis, kidney dysfunction, and immune responses in humans and laboratory animals (Bellomo et al., 1991; Groebler et al., 2011; Paller, 1988; Ramesh and Reeves, 2004). Rhabdomyolysis occurs in humans as a result of the abuse of amphetamines (White, 2002). Therefore, the possibility exists that the striatal neurotoxicity occurring in humans (Ricaurte and McCann, 2005; Volkow et al., 2001; Yuan et al., 2006) might be linked to myoglobin released from damaged muscle, subsequently causing kidney dysfunction. There is no clinical data at present relating serum myoglobin levels to a biomarker for the occurrence

of striatal neurotoxicity in humans. Although significant increases in serum myoglobin and BUN levels occurred in our studies due to AMPH, they did not lead to detectable histological signs of kidney damage. Nonetheless, increased levels of myoglobin in serum indicate that significant tissue damage has occurred, which could directly result in the release of damage associated molecular proteins from muscle and an activation of the immune system (Hirsiger et al., 2012; Tang et al., 2012). We believe that the release of myoglobin, creatine kinase, and other DAMPs from muscle in turn impacts the kidney and, to a lesser extent particularly in NCTR rats, the liver. It is still not clear whether activation of the innate immune system elements outside the brain would influence the striatal neurotoxicity produced by amphetamines. Moreover, as previously stated, there is no causal link between muscle damage or elevated serum levels of myoglobin and striatal neurotoxicity. In addition, the activation of the immune system in the MAV and striatum may not link to the periphery and may be solely dependent on cellular damage in these regions. 5. Conclusion Our results indicate that (1) ‘‘free’’ blood borne LPS increases in EIH and AMPH treated rats are likely not necessary for striatal neurotoxicity and CNS immune responses, and that there is not a notable increase in leukocytes in the circulation resulting from neurotoxic exposures to AMPH, (2) liver and kidney dysfunction by itself is not sufficient to produce striatal neurotoxicity since this occurred with EIH, which does not produce neurotoxicity, (3) it is possible that myoglobin release due to muscle damage may, in some way, be related to striatal neurotoxicity, (4) myoglobin is a potential serum biomarker for AMPH-induced striatal neurotoxicity, and (5) peripheral organ dysfunction is not necessary for, but may exacerbate, striatal neurotoxicity. Conflict of interest statement None declared. Disclaimer The findings and conclusions in this article have not been formally disseminated by the Food and Drug Administration and should not be construed to represent any Agency determination or policy. References Bailey MT, Engler H, Sheridan JF. Stress induces the translocation of cutaneous and gastrointestinal microflora to secondary lymphoid organs of C57BL/6 mice. J Neuroimmunol 2006;171:29–37. Bowyer JF, Davies DL, Schmued L, Broening HW, Newport GD, Slikker W Jr, et al. Further studies of the role of hyperthermia in methamphetamine neurotoxicity. J Pharmacol Exp Ther 1994;268:1571–80. Bowyer JF, Frame LT, Clausing P, Nagamoto-Combs K, Osterhout CA, Sterling CR, et al. Long-term effects of amphetamine neurotoxicity on tyrosine hydroxylase mRNA and protein in aged rats. J Pharmacol Exp Ther 1998a;286:1074–85. Bowyer JF, Holson RR. Methamphetamine and amphetamine neurotoxicity: characteristics, interactions with body temperature and possible mechanisms. In: Chang LW, Dyer RS, editors. Handbook of neurotoxicology. New York: Marcel Dekker Inc.; 1995, pp. 845–70. Bowyer JF, Peterson SL, Rountree RL, Tor-Agbidye J, Wang GJ. Neuronal degeneration in rat forebrain resulting from D-amphetamine-induced convulsions is dependent on seizure severity and age. Brain Res 1998b;809:77–90. Bowyer JF, Tank AW, Newport GD, Slikker W Jr, Ali SF, Holson RR. The influence of environmental temperature on the transient effects of methamphetamine on dopamine levels and dopamine release in rat striatum. J Pharmacol Exp Ther 1992;260:817–24. Bowyer JF, Thomas MT, Schmued LC, Ali SF. Brain region-specific neurodegenerative profiles showing the relative importance of amphetamine dose, hyperthermia, seizures and the blood–brain barrier. Ann N Y Acad Sci 2008;1139:127–39.

Bellomo R, Daskalakis M, Parkin G, Boyce N. Myoglobin clearance during acute continuous hemodiafiltration. Intensive Care Med 1991;17:509–10. Broening HW, Pu C, Vorhees CV. Methamphetamine selectively damages dopaminergic innervation to the nucleus accumbens core while sparing the shell. Synapse 1997;27:153–60. Cass WA, Manning MW, Dugan MT. Effects of neurotoxic doses of methamphetamine on potassium and amphetamine evoked overflow of dopamine in the striatum of awake rats. Neurosci Lett 1998;248:175–8. Clausing P, Bowyer JF. Time course of brain temperature and caudate/putamen microdialysate levels of amphetamine and dopamine in rats after multiple doses of D-amphetamine. Ann N Y Acad Sci 1999;890:495–504. Clausing P, Gough B, Holson RR, Slikker W Jr, Bowyer JF. Amphetamine levels in brain microdialysate, caudate/putamen, substantia nigra and plasma after dosage that produces either behavioral or neurotoxic effects. J Pharmacol Exp Ther 1995;274:614–21. Dunn AJ, Ando T, Brown RF, Berg RD. HPA axis activation and neurochemical responses to bacterial translocation from the gastrointestinal tract Ann N Y Acad Sci 2003;992: 21–9. Eisch AJ, Schmued LC, Marshall JF. Characterizing cortical neuron injury with FluoroJade labeling after a neurotoxic regimen of methamphetamine. Synapse 1998;30:329–33. Fleckenstein AE, Volz TJ, Riddle EL, Gibb JW, Hanson GR. New insights into the mechanism of action of amphetamines. Annu Rev Pharmacol Toxicol 2007;47: 681–98. Goto S, Korematsu K, Oyama T, Yamada K, Hamada J, Inoue N, et al. Neuronal induction of 72-kDa heat shock protein following methamphetamine-induced hyperthermia in the mouse hippocampus. Brain Res 1993;626:351–6. Groebler LK, Liu J, Shanu A, Codd R, Witting PK. Comparing the potential renal protective activity of desferrioxamine B and the novel chelator desferrioxamine B-N-(3-hydroxyadamant-1-yl)carboxamide in a cell model of myoglobinuria. Biochem J 2011;435:669–77. Halpin LE, Yamamoto BK. Peripheral ammonia as a mediator of methamphetamine neurotoxicity. J Neurosci 2012;32:13155–63. Hess A, Desiderio C, McAuliffe WG. Acute neuropathological changes in the caudate nucleus caused by MPTP and methamphetamine: immunohistochemical studies. J Neurocytol 1990;19:338–42. Hirsiger S, Simmen HP, Werner CM, Wanner GA, Rittirsch D. Danger signals activating the immune response after trauma. Mediat Inflamm 2012;2012:315941. Ishigami A, Tokunaga I, Gotohda T, Kubo S. Immunohistochemical study of myoglobin and oxidative injury-related markers in the kidney of methamphetamine abusers. Leg Med (Tokyo) 2003;5(1):42–8. Jayanthi S, Deng X, Noailles PA, Ladenheim B, Cadet JL. Methamphetamine induces neuronal apoptosis via cross-talks between endoplasmic reticulum and mitochondria-dependent death cascades. FASEB J 2004;18:238–51. Kalant H, Kalant OJ. Death in amphetamine users: causes and rates. Can Med Assoc J 1975;112(3):299–304. Kelly KA, Miller DB, Bowyer JF, O’Callaghan JP. Chronic exposure to corticosterone enhances the neuroinflammatory and neurotoxic responses to methamphetamine. J Neurochem 2012;122(5):995–1009. http://dx.doi.org/10.1111/j.1471-4159.2012. 07864.x. Levi MS, Divine B, Hanig JP, Doerge DR, Vanlandingham MM, George NI, et al. A comparison of methylphenidate-, amphetamine-, and methamphetamine-induced hyperthermia and neurotoxicity in male Sprague-Dawley rats during the waking (lights off) cycle. Neurotoxicol Teratol 2012;34:253–62. Lu D, Das DK. Induction of differential heat shock gene expression in heart, lung, liver, brain and kidney by a sympathomimetic drug, amphetamine. Biochem Biophys Res Commun 1993;192:808–12. McGee SM, McGee DN, McGee MB. Spontaneous intracerebral hemorrhage related to methamphetamine abuse: autopsy findings and clinical correlation. Am J Forensic Med Pathol 2004;25(4):334–7. Miller DB, O’Callaghan JP. Environment-, drug- and stress-induced alterations in body temperature affect the neurotoxicity of substituted amphetamines in the C57BL/6J mouse. J Pharmacol Exp Ther 1994;270:752–60. Miller EK, Raese JD, Morrison-Bogorad M. Expression of heat shock protein 70 and heat shock cognate 70 messenger RNAs in rat cortex and cerebellum after heat shock or amphetamine treatment. J Neurochem 1991;56:2060–71. Miller MA, Coon TP. Re: delayed ischemic stroke associated with methamphetamine use. J Emerg Med 2006;31(3):305–6 [author reply 306]. O’Callaghan JP, Miller DB. Neurotoxicity profiles of substituted amphetamines in the C57BL/6J mouse. J Pharmacol Exp Ther 1994;270:741–51. O’Callaghan JP, Miller DB. Neurotoxic effects of substituted amphetamines in rats and mice. In: Massaro EJ, editor. Handbook of neurotoxicology. Totowa, NJ: Humana Press Inc.; 2001, pp. 269–301. Paller MS. Hemoglobin- and myoglobin-induced acute renal failure in rats: role of iron in nephrotoxicity. Am J Physiol 1988;255:F539–44. Quinton MS, Yamamoto BK. Causes and consequences of methamphetamine and MDMA toxicity. AAPS J 2006;8:E337–47. Raghubeer EV, Matches JR. Temperature range for growth of Escherichia coli serotype O157:H7 and selected coliforms in E. coli medium. J Clin Microbiol 1990;28: 803–5. Ramesh G, Reeves WB. Inflammatory cytokines in acute renal failure. Kidney Int 2004;66:S56–61. Ricaurte GA, McCann UD. Recognition and management of complications of new recreational drug use. Lancet 2005;365:2137–45. Richards JR, Johnson EB, Stark RW, Derlet RW. Methamphetamine abuse and rhabdomyolysis in the ED: a 5-year study. Am J Emerg Med 1999;17(7):681–5.

50

M.S. Levi et al. / NeuroToxicology 37 (2013) 40–50

Rothrock JF, Rubenstein R, Lyden PD. Ischemic stroke associated with methamphetamine inhalation. Neurology 1988;38(4):589–92. Schmued LC, Bowyer JF. Methamphetamine exposure can produce neuronal degeneration in mouse hippocampal remnants. Brain Res 1997;759:135–40. Seiden LS, Fischman MW, Schuster CR. Long-term methamphetamine induced changes in brain catecholamines in tolerant rhesus monkeys. Drug Alcohol Depend 1976;1:215–9. Seiden LS, Sabol KE. Neurotoxicity of methamphetamine-related drugs and cocaine. In: Chang LWaRSD, editor. Handbook of neurotoxicology.. New York, Basel, Hong Kong: Marcel Dekker, Inc.; 1995, pp. 824–44. Selkirk GA, McLellan TM, Wright HE, Rhind SG. Mild endotoxemia, NF-kappaB translocation, and cytokine increase during exertional heat stress in trained and untrained individuals. Am J Physiol Regul Integr Comp Physiol 2008;295:R611–23. Sriram K, Miller DB, O’Callaghan JP. Minocycline attenuates microglial activation but fails to mitigate striatal dopaminergic neurotoxicity: role of tumor necrosis factoralpha. J Neurochem 2006;96:706–18. Stephans SE, Yamamoto BK. Methamphetamine-induced neurotoxicity: roles for glutamate and dopamine efflux. Synapse 1994;17:203–9. Tang D, Kang R, Coyne CB, Zeh HJ, Lotze MT. PAMPs and DAMPs: signal 0s that spur autophagy and immunity. Immunol Rev 2012;249:158–75. Thomas DM, Francescutti-Verbeem DM, Liu X, Kuhn DM. Identification of differentially regulated transcripts in mouse striatum following methamphetamine treatment – an oligonucleotide microarray approach. J Neurochem 2004a;88:380–93. Thomas DM, Kuhn DM. MK-801 and dextromethorphan block microglial activation and protect against methamphetamine-induced neurotoxicity. Brain Res 2005;1050: 190–8.

Thomas DM, Walker PD, Benjamins JA, Geddes TJ, Kuhn DM. Methamphetamine neurotoxicity in dopamine nerve endings of the striatum is associated with microglial activation. J Pharmacol Exp Ther 2004b;311:1–7. Thomas M, George NI, Patterson TA, Bowyer JF. Amphetamine and environmentally induced hyperthermia differentially alter the expression of genes regulating vascular tone and angiogenesis in the meninges and associated vasculature. Synapse 2009;63:881–94. Thomas M, George NI, Saini UT, Patterson TA, Hanig JP, Bowyer JF. Endoplasmic reticulum stress responses differ in meninges and associated vasculature, striatum, and parietal cortex after a neurotoxic amphetamine exposure. Synapse 2010;64: 579–93. Tokunaga I, Kubo S, Ishigami A, Gotohda T, Kitamura O. Changes in renal function and oxidative damage in methamphetamine-treated rat. Leg Med (Tokyo) 2006;8:16–21. Volkow ND, Chang L, Wang GJ, Fowler JS, Leonido-Yee M, Franceschi D, et al. Association of dopamine transporter reduction with psychomotor impairment in methamphetamine abusers. Am J Psychiatry 2001;158:377–82. White SR. Amphetamine toxicity. Semin Respir Crit Care Med 2002;23:27–36. Winslow BT, Voorhees KI, Pehl KA. Methamphetamine abuse. Am Fam Physician 2007;76(8):1169–74. Yamaguchi T, Kuraishi Y, Minami M, Nakai S, Hirai Y, Satoh M. Methamphetamineinduced expression of interleukin-1 beta mRNA in the rat hypothalamus. Neurosci Lett 1991;128:90–2. Yuan J, Hatzidimitriou G, Suthar P, Mueller M, McCann U, Ricaurte G. Relationship between temperature, dopaminergic neurotoxicity, and plasma drug concentrations in methamphetamine-treated squirrel monkeys. J Pharmacol Exp Ther 2006;316:1210–8.