Metabonomic Deconvolution Of Embedded Toxicity - American

Chem. Res. Toxicol. 2005, 18, 639-654

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Metabonomic Deconvolution Of Embedded Toxicity: Application To Thioacetamide Hepato- and Nephrotoxicity Nigel J. Waters,†,‡ Catherine J. Waterfield,§ R. Duncan Farrant,| Elaine Holmes,† and Jeremy K. Nicholson*,† Biological Chemistry, Division of Biomedical Sciences, Sir Alexander Fleming Building, Imperial College, London SW7 2AZ, United Kingdom, Preclinical Safety Sciences, GlaxoSmithKline Research & Development, Ware, Herts. SG12 0DP, United Kingdom, and Analytical Technologies, GlaxoSmithKline Research & Development, Stevenage, Herts., SG1 2NY, United Kingdom Received May 12, 2004

We present here the potential of an integrated metabonomic strategy to deconvolute the biofluid metabolic signatures in experimental animals following multiple organ toxicities, using the well-known hepato- and nephrotoxin, thioacetamide. Male Han-Wistar rats were dosed with thioacetamide (150 mg/kg, n ) 25), and urine, plasma, liver, and kidney samples were collected postdose for conventional NMR and magic angle spinning (MAS) NMR spectroscopy. These data were correlated with histopathology and plasma clinical chemistry collected at all time points. 1H MAS NMR data from liver and kidney were related to sequential 1H NMR measurements in urine and plasma using pattern recognition methods. One-dimensional 1H NMR spectra were data-reduced and analyzed using principal components analysis (PCA) to show the time-dependent biochemical variations induced by thioacetamide toxicity. From the eigenvector loadings of the PCA, those regions of the 1H NMR spectra, and hence the combinations of endogenous metabolites marking the main phase of the toxic episode, were identified. The thioacetamide-induced biochemical manifestations included a renal and hepatic lipidosis accompanied by hypolipidaemia; increased urinary excretion of taurine and creatine concomitant with elevated creatine in liver, kidney, and plasma; a shift in energy metabolism characterized by depleted liver glucose and glycogen; reduced urinary excretion of tricarboxylic acid cycle intermediates and raised plasma ketone bodies; increased levels of tissue and plasma amino acids leading to amino aciduria verifying necrosis-enhanced protein degradation and renal dysfunction; and elevated hepatic and urinary bile acids indicating secondary damage to the biliary system. This integrated metabonomic approach has been able to identify the tissue of origin for biomarkers present in the metabolic profiles of biofluids, following the onset and progression of a multiorgan pathology, and as such highlights its potential in the evaluation of embedded toxicity in novel drug candidates.

Introduction Thioacetamide causes centrilobular hepatic necrosis and nephrotoxic damage following single dose administration, while prolonged exposure produces bile duct proliferation and liver cirrhosis. Hepatocarcinomas in experimental animals have also been reported following the administration of thioacetamide (1). Metabolic activation of thioacetamide is via oxidation to thioacetamideS-oxide by a microsomal monooxygenase requiring NADPH and cytochromes P450. Necrotic effects are induced following the oxidation of thioacetamide-S-oxide to sulfate, acetamide, unidentified polar products, and microsome-bound material (2). The reactive metabolite of thioacetamide becomes covalently bound to liver proteins, * To whom correspondence should be addressed. Tel: +44(0)207 594-3195. Fax: +44(0)207 594-3226. E-mail: [email protected]. † Imperial College. ‡ Current Address: Physical & Metabolic Sciences, AstraZeneca R&D Charnwood, Loughborough, Leics. LE11 5RH, United Kingdom. § Preclinical Safety Sciences, GlaxoSmithKline Research & Development. | Analytical Technologies, GlaxoSmithKline Research & Development.

as a further oxidation product of thioacetamide-S-oxide. The adduct has been identified on lysine residues as N-acetyl-L-lysine (3). Radiolabeling experiments have identified specific protein targets for the unknown reactive metabolite of thioacetamide. One such target belongs to the glutathione-S-transferase (GST) superfamily (4). However, it was shown that inhibition of GST was not due to the physical binding of the metabolite to GST but most likely due to a negative effect of thioacetamide on the expression of class R GST subunits. Therefore, a direct correlation between thioacetamide binding to macromolecules and liver damage has not been proven. Thioacetamide is known to affect the expression of several genes and proteins including albumin, β-actin, RNA polymerase, and cytochrome P450 monooxygenases. Thioacetamide provokes a remarkable increase in the size of the nucleoli of liver cells, as well as increasing the amount and specific activity of total mRNA in liver cells. It also causes structural and functional modification of the endoplasmic reticulum (5). In light of the current evidence, the main pathway by which thioacetamide exerts its deleterious effects appears to be modification

10.1021/tx049869b CCC: $30.25 © 2005 American Chemical Society Published on Web 03/24/2005

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of protein expression patterns in the hepatocyte, although lipid peroxidation and apoptosis have also been implicated in thioacetamide-induced pathology (6-9). As such, the mechanisms of toxic action for thioacetamide remain poorly defined in both the liver and the kidney. Extensive reports have advocated the use of highresolution proton NMR and magic angle spinning (MAS)1 NMR spectroscopy together with pattern recognition (PR) analysis for the study of biofluids and tissues from animals treated with drugs and toxins, providing insight into the site, severity, and mechanisms of toxic damage (10-28). This approach has been termed “metabonomics” and is defined as “the quantitative measurement of the multiparametric metabolic response of living systems to pathophysiological stimuli or genetic modification” (29). We have recently reported on the application of an integrated metabonomic approach to the study of the cholestatic hepatotoxin, R-naphthylisothiocyanate (30), and acetaminophen toxicity (31). In most metabonomic studies to date, the toxicity of model compounds on a single organ has been investigated. Here, we apply a metabonomic strategy to analyze the onset and progression of centrilobular hepatic necrosis and nephrotoxic damage induced by the model toxin thioacetamide, to derive the tissue of origin for biofluid metabolic signatures arising from these multiple organ toxicities.

Materials and Methods Animal Handling Procedure and Tissue Preparation. All animal studies were conducted under U.K. Home Office License according to appropriate national legislation. Thirtyfive male Han-Wistar rats (250 g, Charles River, Cambs., U.K.) were acclimatized for 13 days in communal plastic cages prior to group allocation and treatment. They were then housed individually in metabolism cages in a well-ventilated room at a temperature of 21 ( 2 °C and a relative humidity of 50 ( 10%, with a 12 h light/12 h dark cycle. Food [Rat & Mouse No. 1 Diet (Special Diet Services Ltd., Cambridge, U.K.)] and tap water were provided ad libitum. Body weights were measured daily. Rats received either a single dose of thioacetamide in sterile water (p.o. 150 mg/kg, n ) 25) or sterile water only (p.o. 10 mL/ kg, n ) 10). Animals were sacrificed by exsanguination (n ) 5) from the abdominal aorta under isoflurane anaesthesia at 3, 7, 24, 31, and 79 h after dosing with thioacetamide, and vehicledosed control animals were sacrificed at 24 and 31 h p.d. Triplicate samples of the left lateral lobe of the liver and renal cortex from the left kidney, weighing between 15 and 25 mg, were excised and immediately snap-frozen in liquid nitrogen. These samples were stored at -70 °C until NMR spectroscopic analysis. The remainder of the left lateral liver lobe and the left kidney was fixed in 10.5% phosphate-buffered formalin, embedded in paraffin wax, sectioned (3-4 µm), and stained with hematoxylin and eosin for histopathological assessment. 1H MAS NMR Spectroscopic Analysis of Intact Tissues. Samples were rinsed in 0.9% saline 2H2O (D2O), placed in zirconia 4 mm diameter rotors (Bruker Analytische GmbH, Rheinstetten, Germany), and analyzed by 1H MAS NMR spectroscopy at 600.13 MHz using a Bruker DRX600 spectrometer, typically at a MAS spin rate of 6000 Hz (22). Spectra were acquired at 283 K, as measured by the thermocouple system and maintained by the cooling of the inlet gas pressures responsible for sample spinning. The in situ temperature 1 Abbreviations: AMIX, analysis of mixtures; CPMG, Carr-Purcell-Meiboom-Gill; D2O, 2H2O; FID, free induction decay; FT, Fourier transformation; GSH, glutathione (reduced form); GSSG, glutathione (oxidized form); MAS, magic angle spinning; PCA, principal components analysis; PR, pattern recognition; TMAO, trimethylamine-Noxide; TSP, 3-(trimethylsilyl)[2,2,3,3-2H4]propionate.

Waters et al. calculated using the chemical shifts of the water resonance (temperature sensitive) and the R-anomeric proton of glucose (temperature insensitive) averaged 288 K (32). To eliminate the severe dynamic range problem caused by the large water signal, all one-dimensional (1D) 600 MHz 1H NMR spectra were acquired using the standard solvent suppression pulse sequence [(relaxation delay-90°-t1-90°-tm-90°-acquire FID); Bruker Analytische GmbH, Rheinstetten, Germany] in which a secondary irradiation field is applied at the water resonance frequency during the relaxation delay of 3 s and during the mixing period tm (100 ms), with t1 fixed at 3 µs. Typically, 128 transients were collected into 32K data points, with a spectral width of 12000 Hz and an acquisition time per scan of 1.36 s. The 1D CarrPurcell-Meiboom-Gill (CPMG) spin-echo pulse sequence [90 - (τ - 180 - τ)n acquisition] (τ ) 200 µs, n ) 200) with standard presaturation of the water resonance, using a fixed total spinspin relaxation delay of 80 ms, was applied to measure spinecho 1H MAS NMR spectra on all tissue samples. 1H NMR Spectroscopic Analysis of Blood Plasma. Blood (2 mL) was collected into heparinized silica microtainers from abdominal aortal exsanguination. The plasma was separated by centrifugation and frozen prior to NMR measurement. For each sample, 250 µL of blood plasma was made up to a total volume of 600 µL by the addition of an appropriate volume of D2O. The samples were centrifuged at 4200g for 20 min at 4 °C. Aliquots (500 µL) were decanted into 5 mm NMR tubes. 1H NMR spectra were acquired on each sample at 600.13 MHz on a Bruker DRX600 spectrometer at ambient probe temperature (298 K). One-dimensional single pulse experiments were carried out using the standard pulse sequence, as described for the acquisition of MAS NMR spectra. For each sample, 64 transients were collected into 64K data points with a relaxation delay of 2 s and a mixing period of 100 ms. A spectral width of 9600 Hz and an acquisition time per scan of 3.41 s were used. The 1D CPMG spin-echo pulse sequence using a fixed total spin-spin relaxation delay of 80 ms was applied to measure spin-echo 1H NMR spectra on all tissue samples. 1H NMR Spectroscopic Analysis of Urine. Urine was collected continuously from rats in the metabolism cages during the following periods: predose (daytime collection; 0-7 h period on days -5, -4, and -1), 0-7, 7-24, 24-31, 31-48, 48-55, 55-72, and 72-79 h following dosing. Sodium azide was added to the collection vessels as an antibacterial agent. Aliquots (600 µL) of urine were mixed with phosphate buffer (300 µL of 0.1 M) and centrifuged at 2230g for 15 min. The supernatant (800 µL) was pipetted into BEST sample vials together with 0.1% 3-(trimethylsilyl)[2,2,3,3-2H4]propionate (TSP) (δ 0.0)/1% azide in D2O. NMR measurements were made on a Bruker AMX600 spectrometer, operating at 600.13 MHz 1H frequency, using a Bruker BEST flow-injection probe setup (Bruker Analyticshe GmbH, Rheinstetten, Germany). NMR spectra were acquired using the standard pulse sequence (as previously described). For each sample, 64 free induction decays (FIDs) were collected into 48K data points using a spectral width of 12000 Hz and an acquisition time per scan of 2.04 s.

Automatic Data Reduction and Principal Components Analysis (PCA) of 1H NMR Spectra. Single pulse and CPMG (MAS) NMR spectra were data-reduced using the program AMIX (analysis of mixtures, Bruker GmbH, Germany). The spectral region δ 0.2-10.0 was segmented into regions of 0.04 ppm width giving a total of 245 integrated regions per NMR spectrum. The area for each segmented region was expressed as an integral value resulting in an intensity distribution description of the whole spectrum with 245 variables prior to PCA. The region of the spectrum, which included water (δ 4.85.4), was removed from the analysis for all groups to eliminate variation in water suppression efficiency. For urine spectra, the region containing urea (δ 5.4-6.0) was also excluded. All remaining spectral segments were scaled to the total integrated area of the spectrum to reduce the effects of variation in concentration (33).

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Table 1. Effect of Thioacetamide Treatment on Various Blood Plasma Parameters as Measured by Clinical Chemistry Assaysa plasma measurement

control

3h

neutrophils 1.16 ( 0.52 platelets (109/L) 858.33 ( 76.6 fibrinogen (g/L) 1.54 ( 0.55 prothrombin time (s) 23.53 ( 1.32 AP (IU/L) 401.2 ( 140.96 ALT (IU/L) 37.62 ( 8.15 AST (IU/L) 53.08 ( 5.72 GDH (IU/L) 3.55 ( 0.19 SDH (IU/L) 9.46 ( 2.33 GST (IU/L) 30.08 ( 14.42 bilirubin (µmol/L) 0.93 ( 0.27 urea (mmol/L) 5.81 ( 0.72 creatinine (µmol/L) 40.67 ( 1.66 bile acids (µmol/L) 11.08 ( 4.89 albumin (g/L) 38.26 ( 1.77 cholesterol (mmol/L) 1.37 ( 0.18 triglycerides (mmol/L) 1.41 ( 0.68 glucose (mmol/L) 10.24 ( 0.5 blood lactate (mmol/L) 1.6 ( 0.5 (109/L)

7h

24 h

31 h

79 h

2.78 ( 1.58* 2.35 ( 0.49** 3.09 ( 0.55*** 3.35 ( 1.7** 1.36 ( 0.37 977 ( 83.57* 910.4 ( 131.04 734 ( 30.54** 626.2 ( 78.17*** 573.5 ( 71.04** 1.91 ( 0.08 1.85 ( 0.21 0.72 ( 0.34* 0.64 ( 0.36* 2.35 ( 0.68 23.08 ( 1.88 28.64 ( 2.87** 58.87 ( 8.01** 74.1 ( 25.51** 25.52 ( 1.19* 390.8 ( 39.24 556.2 ( 54.57* 558.4 ( 184.17* 624.6 ( 120.77** 470.75 ( 53.98 48.22 ( 10.14 85.96 ( 53.96** 1178.26 ( 840.41*** 2252 ( 758.53*** 152.8 ( 84.22*** 66.92 ( 2.93** 221.8 ( 145.73*** 5636 ( 1739.1*** 7418 ( 2379.7*** 260 ( 65.05*** 6.28 ( 2.89** 65.2 ( 22.97** 105 ( 25** 247 ( 102.57** 73.2 ( 44.13** 17.42 ( 5.85** 302.06 ( 221.58*** 11468 ( 1233.3*** 11332 ( 3619.1*** 380.3 ( 120*** 68.82 ( 23.29** 515.14 ( 103.76*** 3705.26 ( 306.05*** 2822.66 ( 623.99*** 286.78 ( 99.68*** 0.94 ( 0.25 2.04 ( 0.8*** 3.5 ( 1.02*** 5.04 ( 1.29*** 5.27 ( 1.24** 4.9 ( 0.8 5.9 ( 0.42 9.78 ( 0.6*** 16.23 ( 3.24*** 16.62 ( 3.62** 38.6 ( 3.44 44.6 ( 2.07** 54.4 ( 3.05*** 81.2 ( 17.2*** 101 ( 18.95*** 26.46 ( 13.36* 119.04 ( 74.68*** 641 ( 91*** 598 ( 189.43*** 176.4 ( 92.93*** 37.78 ( 0.96 36.96 ( 0.71 38.35 ( 1.44 36.1 ( 1.53 35.3 ( 1.35** 1.31 ( 0.34 1.4 ( 0.22 0.94 ( 0.47 0.83 ( 0.43* 2.62 ( 0.43*** 0.69 ( 0.12* 0.25 ( 0.1*** 0.5 ( 0.18** 0.72 ( 0.19* 1.37 ( 0.61 9 ( 0.83** 8.4 ( 1.79 7.56 ( 0.91*** 7.92 ( 1.23** 7.86 ( 1.21** 2.96 ( 1.59 2.29 ( 0.71 1.82 ( 0.71 3.33 ( 1.18** 1.84 ( 1.19

a Values are expressed as means ( SD. Statistics: (*) p < 0.05, (**) p < 0.01, and (***) p < 0.001 (n ) 5 for all groups, except the control where n ) 9). Key: ALT, alanine aminotransferase; AP, alkaline phosphatase; AST, aspartate aminotransferase; GDH, glutamate dehydrogenase; GST, glutathione-S-transferase; and SDH, sorbitol dehydrogenase.

Table 2. Effect of Thioacetamide Treatment on Body Weighta day

-5

-4

-1

0

1

2

3

body weight (g)

247.52 ( 11.78

250.28 ( 12.92

262.72 ( 14.11

264.85 ( 14.81

240.13 ( 10.31

228.44 ( 13.15

221.62 ( 13.54

a

Values are expressed as means ( SD. Table 3. Urine Data Following Thioacetamide Treatment

urine volume (mL) urine production (mL/h)

control

0-7 h

7-24 h

24-31 h

31-48 h

48-55 h

55-72 h

72-79 h

6.5 ( 2.2 0.93 ( 0.31

6.8 ( 2.9 0.97 ( 0.41

29.1 ( 7.8 1.71 ( 0.46

8.6 ( 2.9 1.23 ( 0.41

18 ( 6.2 1.06 ( 0.36

6.2 ( 1.8 0.89 ( 0.26

12.4 ( 2.7 0.73 ( 0.16

4.8 ( 1.3 0.69 ( 0.19

The single pulse and CPMG 1H NMR spectral data sets were imported into the SIMCA-P8.0 software package (Umetrics AB, Umea, Sweden) separately. The data were then mean-centered prior to PCA. With mean centering, the average value of each variable is calculated and then subtracted from the data, making it possible to directly compare variables (i.e., spectral descriptors or metabolites). PCA methods involve the calculation of linear combinations of the original descriptors, the PCs, such that each PC is orthogonal to all others with the first PC (PC1) containing the largest amount of variance with subsequent PCs containing progressively less variance (34). Thus, a plot of PC1 vs PC2 provides the most efficient two-dimensional representation of the information contained in the data set. Some of the PCA data were presented as mean trajectory plots, whereby the mean coordinate from each group of animals at each time period was calculated and connected in chronological order. Trajectory plots provide an efficient means of simplifying the data, where the magnitude and direction of each point represent the severity and nature of the pathological lesion, respectively. Clinical Chemistry and Hematology. Clinical chemistry analysis of plasma samples was carried out on a Hitachi 917 analyzer using appropriate kits. Blood (0.5 mL) was collected into EDTA anticoagulant to provide plasma for clinical chemistry. A Technicon H* 1 analyzer was used to determine a range of hematological parameters. Also, 1.8 mL of blood was collected into 0.2 mL of 0.106 M trisodium citrate. Prothrombin time and fibrinogen concentration were determined using a Sysmex CA5000 analyzer. Unpaired Mann-Whitney tests were used to compare clinical chemical data between groups.

Results Clinical Chemistry and Hematology. The activity of the plasma enzymes, alkaline phosphatase and alanine aminotransferase, increased at 7-31 and 7-79 h p.d., respectively, while aspartate aminotransferase, glutamate

dehydrogenase, sorbitol dehydrogenase, and GST all increased severalfold over the entire time course (Table 1). A raised neutrophil count at 3-31 h p.d. was accompanied by decreased platelets and fibrinogen at 2479 and 24-31 h, respectively, while the prothrombin time increased at 7-79 h (Table 1). A variety of plasma metabolites also displayed time-dependent alterations as follows: increased bile acids and a decline in glucose over the entire time course; elevated bilirubin and creatinine at 7-79 h; a decrease in triglycerides at 3-31 h; raised levels of urea at 24-79 h; a transient increase in lactate at 31 h; and lowered albumin levels associated with augmented cholesterol at 79 h (Table 1). In addition, Table 2 presents the average daily body weight data from treated animals illustrating a dramatic weight loss postdose, and Table 3 summarizes the urine volumes and excretion in mL/h. The urinary volume excreted by thioacetamide-treated animals was slightly elevated at 7-24 h p.d., which is consistent with a renal effect. All NMR urine spectra were normalized to the total spectral integral in order to partially compensate for differences in urine volume and osmolarity. Histopathological Assessment. In liver at 7 h p.d., all animals showed centrilobular inflammation and multifocal single cell necrosis, together with centrilobular pallor and glycogen depletion (data not shown). By 24 and 31 h p.d., all animals showed moderate to marked centrilobular necrosis with moderate inflammation, periportal inflammatory infiltration, glycogen depletion, and lipid accumulation. At 79 h after treatment, all animals showed slight centrilobular necrosis with inflammation, glycogen depletion, and lipid accumulation. At 24-79 h, there were signs of cells undergoing apoptosis. In kidney,

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Figure 1. Series of 600 MHz single pulse 1H NMR spectra (δ 0.5-4.5) of urine at predose and various time points following the administration of thioacetamide (150 mg/kg). Key: 2OG, 2-oxoglutarate; 3HB, 3-D-hydroxybutyrate; Ace, acetate; Ala, alanine; Bet, betaine; Cho, choline; Cit, citrate; Cre, creatine; Crn, creatinine; DMG, dimethylglycine; Glc, glucose; Gln, glutamine; Glu, glutamate; Gly, glycine; Ile, isoleucine; Lac, lactate; Leu, leucine; Lys, lysine; Met, methionine; m-HPPA, 3-(3-hydroxyphenyl)propionic acid; NAG, N-acetyl glycoproteins; Pro, proline; Suc, succinate; Tau, taurine; ThAcSO, thioacetamide-S-oxide; Thr, threonine; Tyr, tyrosine, U1, unassigned; and Val, valine.

slight vacuolar degeneration was seen in the tubules of all animals at 24 and 31 h p.d. and lipid accumulation was noted. By 79 h p.d., moderate to marked degeneration of the distal tubules and thick ascending loop of Henle´ was present in all individuals. 1 H NMR Spectroscopic and PR Analysis of Urine. NMR-PCA urinalysis highlighted perturbations in a number of endogenous species (Figures 1-3). The Kreb’s cycle intermediates decreased over the entire time course. Signals from citrate and 2-oxoglutarate declined 0-79

h, while succinate increased initially (0-24 h) before decreasing at 24-79 h. Raised lactate over the study time course, together with elevated acetate (0-72 h), was accompanied by reduced creatinine (48-79 h). Other increases in urinary metabolites included the following: betaine (7-24 h), spermidine (7-48 h), dimethylglycine (7-79 h), bile acids (24-72 h), taurine and creatine (2479 h), 3-D-hydroxybutyrate (48-55 h), and formate (5572 h). At 31-79 h p.d., resonances from the amino acids isoleucine, leucine, valine, alanine, lysine, methionine,



Figure 2. Series of 600 MHz single pulse 1H NMR spectra (δ 6.1-8.5) of urine at predose and various time points following the administration of thioacetamide (150 mg/kg). The key is the same as for Figure 1 with the following additions: PAG, phenylacetylglycine; Phe, phenylalanine; and U2, U3, U4, and U5, unassigned.

tyrosine, threonine, proline, glutamate, glutamine, glycine, phenylalanine as well as glucose, choline, and trimethylamine-N-oxide (TMAO) all increased. Hippurate and the chlorogenic acid metabolite, mhydroxypropionic acid (m-HPPA), were present in predose urine samples. The predose group and some of the early time point groups separated in PC3 on the basis of mainly hippurate excretion vs mainly m-HPPA excretion (Figure 3). On administration of thioacetamide, the urinary levels of these two gut microflora-derived species decreased over the entire time course and led to the

converging trajectory shown in PC1 vs PC3 (Figure 3). Phenylacetylglycine also decreased at 0-31 h p.d. Massive levels of a signal corresponding to thioacetamide-S-oxide (δ 2.19 singlet) appeared at 0-7 h with progressively smaller levels being detected at 7-55 h p.d. In addition, acetamide became apparent in urine NMR spectra at 0-48 h. A number of unassigned signals were detected in urine spectra and are outlined in Table 4. 1 H NMR Spectroscopic and PR Analysis of Blood Plasma. 1H NMR spectroscopic and PC analysis of blood plasma from animals following thioacetamide treatment

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Figure 3. PCA scores plots mapping the position of urine 1H NMR spectra obtained from individuals following treatment with sterile water (control) or thioacetamide at various time points postdose. Plots correspond to PC1 vs PC2 (A) and PC1 vs PC3 (B), where the data variance explained in each plot is 76.7 and 71.1%, respectively. Table 4. Unassigned 1H NMR Resonances Observed in Urine from Thioacetamide-Treated Rats unassigned reference U1 U2 U3 U4 U5

chemical change following shift thioacetamide time-course (ppm) multiplicity dosing of change (h) 2.40 7.69 7.06 6.30 7.84

t s d t t

v

31-79

v v v

31-79 48-79 72-79

showed a number of marked changes (Figure 4). These included an increase in choline and creatine at 7-79 h p.d., elevated levels of the amino acids valine and glutamine at 7-31 h p.d., and leucine, isoleucine, glutamate, lysine, and alanine at 24-31 h p.d. associated with a rise in 3-D-hydroxybutyrate and acetoacetate also at 24-31 h p.d. (Figure 5). A drop in citrate levels was detected over the study time course, while low density lipoprotein (LDL) resonances decreased at 3-31 h accompanied by augmented acetate and the appearance of

acetamide and an unassigned singlet at δ 2.09. The signal corresponding to TMAO increased at 24 h p.d. Some individuals exhibited raised lactate at 3-7 h p.d. PCA of the aromatic region of the blood plasma 1H NMR spectra (δ 6-10) revealed a decrease in formate at 3-31 h p.d., an increase in phenylalanine and an unassigned doublet (24-79 h) and tyrosine (24 h), the disappearance of 1-methylhistidine signals at 24-31 h together with the appearance of fumarate and the emergence of a singlet (δ 7.83) tentatively assigned as formiminoglutamate (Figure 6). 1 H MAS NMR Spectroscopic and PR Analysis of Intact Renal Cortex. 1H MAS NMR spectroscopic analysis of intact renal cortex from thioacetamide-treated rats, with the aid of PCA, identified a variety of biochemical changes (Figures 7 and 8). These included a decrease in taurine over the entire time course, elevated lipid resonances at 7-79 h p.d. with increased TMAO and choline at 7 h p.d., and heightened levels of creatine at 24-79 h p.d. These perturbations were associated with



Figure 4. Series of 600 MHz CPMG 1H NMR spectra (δ 0.5-6.0) of blood plasma at various time points following the administration of thioacetamide (150 mg/kg). The key is the same as for Figure 1 with the following additions: Aca, acetoacetate; d-Ala, deuterated alanine; HOD, residual water; LDL1, low density lipoprotein terminal methyl; LDL2, low density lipoprotein (CH2)n; NAC1/NAC2, composite acetyl signals from R1-acid glycoprotein; region A, signals from glycerol, glucose, and amino acid CH protons; and U1, unassigned.

augmented levels of the amino acids, alanine (3-31 h), glycine (7-24 h), isoleucine, leucine, and valine (3-79 h). On closer inspection of the 1H MAS NMR spectra, a decrease in myo-inositol was observed at 3 h p.d. 1 H MAS NMR Spectroscopic and PR Analysis of Intact Liver. 1H MAS NMR spectroscopic analysis of intact liver tissue from thioacetamide-treated rats, with the aid of PCA, identified a variety of biochemical changes (Figures 9 and 10). These included depleted

glucose and glycogen at 7-79 h with glycogen levels below the detection limits of the MAS NMR experiment at 7 h onward. Also at 7-79 h, an augmentation of the lipid, choline, and phosphocholine resonances accompanied a drop in TMAO and betaine signals. Elevated isoleucine, leucine (24-79 h), and creatine (24-31 h) was observed, and closer inspection of the 1H MAS NMR spectra revealed increases in glutathione (GSH) (24-31 h), bile acids, and methionine (24-79 h).

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Figure 5. PCA scores plots mapping the position of plasma CPMG 1H NMR spectra obtained from individuals following treatment with sterile water (control) or thioacetamide at time points postdose. The data variance explained is 60.2%.

In liver at 24-79 h, some individuals exhibited elevated levels of the amino acids histidine, phenylalanine, and tyrosine; the nitrogenous bases inosine, adenine, and uracil as well as formate and N-methylnicotinamide (Figure 11).

Discussion Lesion Specific Biomarkers of Hepatotoxicity and Nephrotoxicity. Elevated activities for the plasma enzymes alkaline phosphatase, alanine aminotransferase, aspartate aminotransferase, glutamate dehydrogenase, sorbitol dehydrogenase, and GST, as well as changes in various plasma metabolites, including bilirubin and bile acids, confirmed the presence of liver damage manifested as centrilobular necrosis. An increased neutrophil count suggested an inflammatory reaction to thioacetamide treatment. Blood clotting parameters, including an extended prothrombin time, also implied a perturbation in normal liver function. Elevated plasma cholesterol at 79 h p.d. is in agreement with studies on the chronic effects of thioacetamide, where the cholesterol is derived from hepatic lysosome membranes making these organelles more susceptible to liberating their lytic contents (35). Raised plasma levels of urea indicated thioacetamide-induced kidney dysfunction. Histopathology was able to corroborate the structural progression of both the liver and the kidney lesions over time. Results from clinical chemistry assays of plasma triglycerides are in good agreement with 1H NMR spectroscopy of plasma. Measurements of plasma glucose and blood lactate, however, appear to differ in that NMRPC analysis showed a rise in lactate at 3-7 h with no change in glucose while conventional assays demonstrated an increase in lactate at 31 h p.d. with a reduction in glucose at 3-79 h p.d. Possible explanations for this discrepancy are currently being investigated. 1H NMR urinalysis identified a rise in creatine and taurine. These are well-known urinary biomarkers of liver injury (36, 37), and this pattern of changes in creatine and taurine is indicative of hepatic necrosis. Creatine is synthesised and metabolized in the liver, with the kidney providing the necessary synthetic precursor. Guanidinoacetate,

formed in the kidney, is transported through the blood, undergoing methylation in the liver to form creatine, which enters the blood for use in peripheral tissues (38). Therefore, the observed increases in liver, kidney, plasma, and urine suggest leakage from necrotic cells or upregulated creatine biosynthesis. Elevations in hepatic bile acids, urinary bile acids, and glucose, as detected by 1H (MAS) NMR, are markers of damage to the biliary system. These changes were accompanied by excretion of a range of amino acids indicating renal dysfunction. The observed glycosuria, amino aciduria, and depleted TCA cycle intermediates are in agreement with previous 1 H NMR urinalysis studies on thioacetamide toxicity (14). NMR studies on perchloric acid kidney extracts have shown reduced concentrations of the osmolytes betaine, taurine, and GPC as a consequence of polycystic kidney disease (39). Such organic osmolytes are accumulated in kidney tissue to maintain cellular osmotic balance thus alleviating the perturbing and toxic effects of high salt or urea concentrations (40, 41). This may explain the decrease in renal taurine and myo-inositol levels, being a prerequisite to the manifestation of thioacetamideinduced nephrotoxic damage. Energy Intermediary Metabolism. A number of metabolites involved in energy generation were perturbed as a result of thioacetamide treatment. Depleted liver glucose and glycogen at 7-79 h p.d. was followed by a drop in citrate with elevated lactate in plasma suggesting extensive glycogenolysis and glycolysis (42). Glycosuria at 31-79 h without any detected perturbation of plasma glucose suggests renal damage as opposed to saturation or inhibition of kidney glucose reabsorption. Reduced urinary excretion of Kreb’s cycle intermediates, citrate, succinate, and 2-oxoglutarate, a common nonspecific effect of toxic injury, indicated increased energy metabolism. Depletion of glycogen is more severe in necrotic centrilobular hepatocytes and correlates with the increase in glycogen phosphorylase (EC 2.4.1.1) activity, probably caused by elevated cytosolic Ca2+ concentrations resulting from damage to cell membranes as observed by Anghileri et al. (43).



Figure 6. Series of 600 MHz CPMG 1H NMR spectra (δ 6.0-8.5) of blood plasma at various time points following the administration of thioacetamide (150 mg/kg). The key is the same as for Figure 1 with the following additions: 1-MeHis, 1-methylhistidine; Phe, phenylalanine; and U2, unassigned.

Raised levels of plasma 3-D-hydroxybutyrate, acetoacetate, and acetate led to enhanced levels of lactate, acetate, and 3-D-hydroxybutyrate in the urine, verifying a shift in energy metabolism toward ketone body formation and utilization. This is supported by the progressive loss in body weight observed over the 3 days after dosing, likely to result from reduced feeding. Lipid Metabolism. The raised lipid levels observed in the 1H MAS NMR spectra of both liver and kidney represent drug-induced lipid accumulation, which may

be caused by a number of mechanisms. The accumulation of triglycerides and/or phospholipids is a common cellular response to toxic compounds, which is normally reversible. This can occur by increased synthesis or uptake of lipid, decreased lipid metabolism, or inhibition of lipid transport from the cell. The latter of these is the most likely case as the observed hepatic lipidosis was concomitant with hypolipidaemia at 3-31 h p.d. Inhibition of protein synthesis, at the transcription, translation, or assembly level, will block the manufacture of the lipid

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Figure 7. Series of 600 MHz single pulse 1H MAS NMR spectra (δ 0.5-6.0) of renal cortex at various time points following the administration of thioacetamide (150 mg/kg). The key is the same as for Figure 1 with the following addition: GPC, glycerophosphorylcholine.

acceptor protein (apoprotein) required for the transport of lipid out of the hepatocyte as LDL. Thioacetamide is known to impair the synthesis of messenger and transfer RNA or their transport from the nucleus (44) as well as causing structural and functional modification of the endoplasmic reticulum (5). Thioacetamide affects the expression of albumin (45), explaining the lowered plasma albumin levels at 79 h p.d.. The lowered plasma LDL levels observed at 3-31 h are in accordance with published data on the chronic effects of thioacetamide (46, 47).

The manifestation of lipid accumulation in the kidney may be the result of a sequence of distinct toxic incidents. Unlike liver, the kidney does not intrinsically control lipid levels and so the observed renal lipidosis is likely to arise by a different mechanism. This toxin-induced effect has been observed with carbon tetrachloride (Waterfield, personal communication) and chromium (48). It has been suggested that this could be caused by increased uptake of free fatty acids or inhibition of esterases (48). It is known that myo-inositol deficiency leads to perturbed LDL and lipid secretion and a subsequent hepatic lipid



Figure 8. PCA metabolic mean trajectory plot mapping the average position of renal cortex single pulse 1H MAS NMR spectra for each time point following thioacetamide treatment. Principal components 1 and 2 explain 96.5% of the data variance.

accumulation, as inositol is a precursor of phosphatidylinositol, which promotes lipid secretion (49, 50). Therefore, this may explain the observed decrease in renal myoinositol and the ensuing MAS NMR detected kidney lipid accumulation. A block in lipid degradation is not the likely cause of the observed steatoses as elevated phosphocholine was detected in the liver and increased choline was detected in liver, kidney, and plasma. Therefore, this suggests the removal of the excess lipid by enzymatic conversion to intermediate species. Choline and phosphocholine are products of lipid catabolism mediated by the enzymes, phospholipase C and D (EC 3.1.4.3/4). It appears that in kidney, this choline is a substrate for the enzymes involved in N-oxidation (51-53) leading to the rise in TMAO at 7 h p.d. However, in liver, the drop in TMAO levels suggests a thioacetamide-induced block in this N-oxidation pathway. The lipid intermediary metabolites, choline, betaine, and TMAO, were present in the blood plasma before excretion into the urine. Dimethylglycine, an intermediate in the conversion of choline and betaine to glycine, was also excreted into the urine. Amino Acid and Nucleotide Metabolism. The elevated levels of amino acids observed in both liver and kidney precede the increases in plasma amino acids. It is reasonable to consider such increases arising from the necrosis of the liver parenchyma as well as muscle proteolysis inherent to secondary malnutrition, as has been shown in serum of rats with thioacetamide-induced liver cirrhosis (54). In support of muscle proteolysis, lysine, glutamate, and glutamine were only seen to increase in the biofluids and not the tissues studied by MAS NMR. Disturbed protein synthesis as well as necrosis-induced protein degradation would lead to augmented concentrations of free amino acids in the target organs and blood. The fact that these amino acids subsequently appeared in the urine indicates altered amino acid reabsorption and damage to the proximal renal tubule. These changes in amino acids were accompanied by increases in inosine, adenine, and uracil in liver suggesting nucleic acid catabolism. Studies on the activity and expression of rat liver GST during thioacetamide intoxication have illustrated that

thioacetamide metabolites have a negative effect on GST expression (4). GST catalyzes the formation of a thioether linkage between GSH and a second substrate with an electrophilic center, and such an inhibitory effect may explain the rise in GSH signal intensities in MAS NMR spectra of liver. This may also be explained by the induction of glucose-6-phosphate dehydrogenase (G6PDH) in response to thioacetamide-induced hepatic injury and oxidative stress, leading to an increase in the reaction product, NADPH (6). Increases in this reducing equivalent would then lead to an increase in the GSH:GSSG (oxidized form) ratio. However, in the absence of GSSG detection due to overlapping signals, this argument remains tentative and warrants further investigation. Thioacetamide is known to induce and stabilize the enzyme ornithine decarboxylase (EC 4.1.1.17) (55-57). This enzyme is the first and rate-limiting step in mammalian polyamine biosynthesis, which decarboxylates ornithine to putrescine, the immediate precursor of spermidine (58-60). This led to spermidine excretion in the urine at 7-48 h. Urinary spermidine is also known to increase in patients with hepatocellular carcinoma (61). An intermediate in the histidine catabolism pathway, formiminoglutamate, was detected in plasma at 24-31 h p.d. This was accompanied by reduced plasma 1-methylhistidine. Formiminoglutamate is known to accumulate in body fluids in cobalamin and folate deficiency (6264) and thioacetamide may interfere with the availability of these vitamin cofactors. The accumulation in formiminoglutamate may also occur as a result of thioacetamide-induced inhibition of the enzyme, glutamate formimino transferase that converts formiminoglutamate into glutamate. This enzyme utilizes tetrahydrofolate as a cofactor. Folate deficiency also leads to the appearance of formate in the urine (62), and this was the case at 5572 h p.d. Formaldehyde dehydrogenase catalyzes the reversible conversion of formaldehyde, GSH, and NAD+ to NADH, H+, and S-formyl-GSH (65). The latter is cleaved by a hydrolase to formate and GSH. Activities of these enzymes have been found in many animal tissues and, although their function is not yet clear, could explain

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Figure 9. Series of 600 MHz CPMG 1H MAS NMR spectra (δ 0.5-6.0) of the left lateral lobe of the liver at various time points following the administration of thioacetamide (150 mg/kg). The key is the same as for Figure 1 with the following additions: d-Ala, deuterated alanine; and PCho, phosphocholine.

the perturbations in formate seen in liver, plasma, and urine. N-Methylnicotinamide resonances increased in liver at 24-79 h p.d. Human liver cirrhotic patients show raised nicotinamide methylation, and it has been postulated that this methylating pathway may play a protective role against the toxic effect of intracellular accumulation of nicotinamide arising from the catabolic state of cirrhotic tissue (66). This may also be the case with thioacetamide toxicity. N-Methylnicotinamide may arise in the damaged liver because further metabolism (to 2-pyridone-5-car-

boxamide) is not possible because the enzymes responsible (xanthine oxidase and aldehyde oxidase) are inundated with high levels of purine and pyrimidine substrates resulting from necrosis-enhanced nucleotide catabolism. Alterations in the levels of nicotinamide intermediates are likely to lead to perturbations in the availability of coenzymes such as NADH and NADPH and thus a wide variety of energy transfer processes. Furthermore, variation in nicotinamide may interfere with oxidative balance leading to lipid peroxidation and greater consumption of antioxidants such as vitamin E and GSH (67).



Figure 10. PCA scores plots showing the mapping position of liver CPMG 1H MAS NMR spectra obtained from individuals treated with sterile water (control) or thioacetamide at various time points postdose. Plot of PC 1 vs PC 2 where the data variance explained is 94.1%.

Figure 11. Series of 600 MHz CPMG 1H MAS NMR spectra (δ 6.0-9.0) of the left lateral lobe of the liver at 24 h following the administration of sterile water vehicle or thioacetamide (150 mg/kg). The key is the same as for Figure 1 with the following additions: His, histidine; Phe, phenylalanine; and U1 and U2, unassigned.

Thioacetamide Metabolism. 1H NMR spectroscopic analysis of urine detected both the primary metabolite of thioacetamide, thioacetamide-S-oxide, and one of the products of further metabolism, acetamide. 1H NMRPCA of blood plasma detected acetamide and an unassigned singlet at δ 2.09, which may correspond to the parent compound or an N-acetylated species derived from thioacetamide biotransformation. Thioacetamide is known to stimulate spermidine acetylase (EC 2.3.1.57) (68). The available evidence suggests that the development of hepatic necrosis after the administration of thioacetamide occurs as follows: first, thioacetamide is metabolized by cytochrome P450 containing mixed function oxidases in the liver to thioacetamide-S-oxide. Second, thioacetamide-S-oxide is converted in a cytochrome P450 monooxygenase-catalyzed reaction to an intermediate capable of either reacting with tissue macromolecules or decomposing to acetamide. Finally, the formation of

irreversibly bound products of thioacetamide-S-oxide metabolism may initiate changes in hepatic cellular function resulting in cell death (2, 44, 69, 70). Urinary Excretion of Gut Microflora-Derived Metabolites. PC analysis of 1H NMR urine spectra highlighted separation in the predose group and early time points up to 31 h p.d., based on the differential urinary excretion of hippurate, m-HPPA, and phenylacetylglycine. On inspection of spectra, individuals were found to predominantly excrete either hippurate or m-HPPA, as reported previously (71). These species are present in urine as products of intestinal microfloral metabolism of dietary-derived plant phenolics (e.g., chlorogenic acid) and aromatic amino acids (72-77). As the toxin-induced lesion developed, the excretion of the three bacterially derived species decreased while the thioacetamide-induced response by all individuals became uniform, leading to the converging metabolic PCA trajec-

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Figure 12. Integrated metabonome diagram describing the NMR-detected biochemical changes observed in urine, blood plasma, renal cortex, and liver in the Han-Wistar rat following thioacetamide treatment. The key is the same as for Figure 1 with the following additions: v, above control levels; V, below control levels; Aca, acetoacetate; FIGlu, formiminoglutamate; His, histidine; LDL, low density lipoprotein; MeHis, 1-methylhistidine; NMN, N-methylnicotinamide; and Phe, phenylalanine. Note: L, P, K, and U refer to samplings of liver, plasma, kidney, and urine, respectively.

tory. This implies that thioacetamide was not only affecting the host but also the symbiotic gut microflora and the enzyme systems responsible for phenolic metabolism. Integrated Metabonomics: Deconvolution of Embedded Toxicity. The thioacetamide-induced perturbations in endogenous metabolites observed in the 1H NMR-PR analysis of intact liver, intact kidney, blood

plasma, and urine are summarized in Figure 12, highlighting the correlations between time-dependent changes in each biomatrix. With such a vast and complex number of toxin-induced changes in the endogenous metabolite profiles of urine and plasma, it is necessary to determine the tissue origin of these biomarkers in order to deconvolute the series of biochemical events in the onset and progression of toxicities in more than one tissue. This


study has utilized high resolution 1H MAS NMR spectroscopy of intact tissues, which provides the essential link between the metabolite profiles obtained from biofluid NMR and the structural progression of the lesion observed by histopathological techniques. The ability of integrated metabonomic studies to delineate and define the tissue origin of biomarkers present in biofluids lends itself to novel drug candidate safety investigations in the pharmaceutical discovery setting, where embedded toxicity is not uncommon. Such an approach allows the deconvolution of embedded pathologies, identifying and locating sites of toxin-induced damage, and is thus able to direct histopathology.


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References (1) Fitzhugh, O. G., and Nelson, A. A. (1948) Liver tumors in rats fed thiourea or thioacetamide. Science 108, 626-628. (2) Porter, W. R., and Neal, R. A. (1978) Metabolism of thioacetamide and thioacetamide-S-oxide by rat liver microsomes. Drug Metab. Dispos. 6, 379-388. (3) Dyroff, M. C., and Neal, R. A. (1981) Identification of the major protein adduct formed in rat liver after thioacetamide administration. Cancer Res. 41, 3430-3435. (4) Spira, B., and Raw, I. (2000) The effect of thioacetamide on the activity and expression of cytosolic rat liver glutathione-Stransferase. Mol. Cell. Biochem. 211, 103-110. (5) Barker, E. A., and Smuckler, E. A. (1972) Altered microsome function during acute thioacetamide poisoning. Mol. Pharmacol. 8, 318-326. (6) Diez-Fernandez, C., Sanz, N., and Cascales, M. (1996) Changes in glucose-6-phosphate dehydrogenase and malic enzyme gene expression in acute hepatic injury induced by thioacetamide. Biochem. Pharmacol. 51, 1159-1163. (7) Sun, F., Hayami, S., Ogiri, Y., Haruna, S., Tanaka, K., Yamada, Y., Tokumaru, S., and Kojo, S. (2000) Evaluation of oxidative stress based on lipid hydroperoxide, vitamin C and vitamin E during apoptosis and necrosis caused by thioacetamide in rat liver. Biochim. Biophys. Acta 1500, 181-185. (8) Ledda-Columbano, G. M., Coni, P., Curto, M., Giacomini, L., Faa, G., Oliverio, S., Piacentini, M., and Columbano, A. (1991) Induction of two different modes of cell death, apoptosis and necrosis, in rat liver after a single dose of thioacetamide. Am. J. Pathol. 139, 1099-1109. (9) Hayami, S., Ikeda, K., Sun, F., Tanaka, K., and Kojo, S. (1999) Increase of caspase-3 activity in rat liver and plasma by thioacetamide. Biochem. Pharmacol. 58, 1941-1943. (10) Holmes, E., Nicholls, A. W., Lindon, J. C., Connor, S. C., Connelly, J. C., Haselden, J. N., Damment, S. J. P., Spraul, M., Neidig, P., and Nicholson, J. K. (2000) Chemometric models for toxicity classification based on NMR spectra of biofluids. Chem. Res. Toxicol. 13, 471-478. (11) Griffin, J. L., Walker, L. A., Troke, J., Osborn, D., Shore, R. F., and Nicholson, J. K. (2000). The initial pathogenesis of cadmium induced renal toxicity. FEBS Lett. 23962, 1-4. (12) Nicholls, A. W., Nicholson, J. K., Haselden, J. N., and Waterfield, C. J. (2001) A metabonomic approach to the investigation of druginduced phospholipidosis: A NMR spectroscopy and pattern recognition study. Biomarkers 5, 410-423. (13) Garrod, S., Humpfer, E., Connor, S. C., Connelly, J. C., Spraul, M., Nicholson, J. K., and Holmes, E. (2000) High resolution 1H NMR and magic angle spinning NMR spectroscopic investigation of the biochemical effects of 2-bromoethanamine in intact renal and hepatic tissue. Magn. Reson. Med. 45, 781-790. (14) Holmes, E., Nicholls, A. W., Lindon, J. C., Ramos, S., Spraul, M., Neidig, P., Connor, S. C., Connelly, J., Damment, S. J. P., Haselden, J., and Nicholson, J. K. (1998) Development of a model for classification of toxin-induced lesions using 1H NMR spectroscopy of urine combined with pattern recognition. NMR Biomed. 11, 235-244. (15) Beckwith-Hall, B. M., Nicholson, J. K., Nicholls, A. W., Foxall, P. J. D., Lindon, J. C., Connor, S. C., Abdi, M., Connelly, J., and Holmes, E. (1998) Nuclear magnetic resonance spectroscopic and principal components analysis investigations into biochemical effects of three model hepatotoxins. Chem. Res. Toxicol. 11, 260272. (16) Holmes, E., Bonner, F. W., and Nicholson, J. K. (1995) Comparative studies on the nephrotoxicity of 2-bromoethanamine hydro-

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(22)

(23)

(24)

(25)

(26)

(27)

(28)

(29)

(30)

(31)

(32)

(33)

(34) (35)

bromide in the Fischer 344 rat and the multimammate desert mouse (Mastomys natalensis). Arch. Toxicol. 70 (2), 89-95. Anthony, M. L., Sweatman, B. C., Beddell, C. R., Lindon, J. C., and Nicholson, J. K. (1994) Pattern recognition classification of the site of nephrotoxicity based on metabolic data derived from proton nuclear magnetic resonance spectra of urine. Mol. Pharmacol. 46, 199-211. Brindle, J. T., Antti, H., Holmes, E., Tranter, G., Nicholson, J. K., Bethell, H. W. L., Clarke, S., Schofield, P. M., McKilligin, E., Mosedale, D. E., and Grainger, D. J. (2002) Rapid and noninvasive diagnosis of the presence and severity of coronary heart disease using 1H NMR based metabonomics. Nat. Med. 8 (12), 1439-1445. Gartland, K. P. R., Bonner, F. W., and Nicholson, J. K. (1989) Investigations into the biochemical effects of region-specific nephrotoxins. Mol. Pharmacol. 35, 242-250. Waters, N. J., Holmes, E., Waterfield, C. J., Farrant, R. D., Nicholson, J. K. (2001) NMR and pattern recognition studies on liver extracts and intact livers from rats treated with R-naphthylisothiocyanate. Biochem. Pharmacol. 64 (1), 67-77. Humpfer, E., Spraul, M., Nicholls, A. W., Nicholson, J. K., and Lindon, J. C. (1997). Direct observation of resolved intracellular and extracellular water signals in intact human red blood cells using 1H MAS NMR spectroscopy. Magn. Reson. Med. 38, 334336. Waters, N. J., Garrod, S., Farrant, R. D., Haselden, J. N., Connor, S. C., Connelly, J., Lindon, J. C., Holmes, E., and Nicholson, J. K. (2000) High-resolution magic angle spinning 1H NMR spectroscopy of intact liver and kidney: Optimization of sample preparation procedures and biochemical stability of tissue during spectral acquisition. Anal. Biochem. 282, 16-23. Garrod, S., Humpfer, E., Spraul, M., Connor, S. C., Polley, S., Connelly, J., Lindon, J. C., Nicholson, J. K., and Holmes, E. (1999) High-resolution magic angle spinning 1H NMR spectroscopic studies on intact rat renal cortex and medulla. Magn. Reson. Med. 41, 1108-1118. Bollard, M. E., Garrod, S., Holmes, E., Lindon, J. C., Humpfer, E., Spraul, M., and Nicholson, J. K. (2000) High resolution 1H and 1H-13C magic angle spinning NMR spectroscopy of rat liver. Magn. Reson. Med. 44, 201-207. Millis, K. K., Maas, W. E., Cory, D. G., and Singer, S. (1997) Gradient high-resolution magic angle spinning NMR spectroscopy of human adipocyte tissue. Magn. Reson. Med. 38, 399-403. Tomlins, A. M., Foxall, P. J. D., Lindon, J. C., Lynch, M. J., Spraul, M., Everett, J. R., and Nicholson, J. K. (1998) High-resolution magic angle spinning 1H NMR analysis of intact prostatic hyperplastic and tumour tissues. Anal. Commun. 35, 113-115. Griffin, J. L., Troke, J., Walker, L. A., Shore, R. F., Lindon, J. C., and Nicholson, J. K. (2000) The biochemical profile of rat testicular tissue as measured by magic angle spinning 1H NMR spectroscopy. FEBS Lett. 486, 225-229. Cheng, L. L., Ma, M. J., Becerra, L., Ptak, T., Tracey, I., Lackner, A., and Gonzalez, R. G. (1997) Quantitative neuropathology by high-resolution magic angle spinning proton magnetic resonance spectroscopy. Proc. Natl. Acad. Sci. U.S.A. 94, 6408-6413. Nicholson, J. K., Lindon, J. C., and Holmes, E. (1999) ‘Metabonomics’: understanding the metabolic responses of living systems to pathophysiological stimuli via multivariate statistical analysis of biological NMR spectroscopic data. Xenobiotica 29, 1181-1189. Waters, N. J., Holmes, E., Williams, A., Waterfield, C. J., Farrant, R. D., and Nicholson, J. K. (2001) NMR and pattern recognition studies on the time-related metabolic effects of R-naphthylisothiocyanate on liver, urine and plasma in the rat: An integrative metabonomic approach. Chem. Res. Toxicol. 14, 1401-1412. Coen, M., Lenz, E. M., Nicholson, J. K., Wilson, I. D., Pognan, F., and Lindon, J. C. (2002) An integrated metabonomic investigation of acetaminophen toxicity in the mouse using NMR spectroscopy. Chem. Res. Toxicol. 16 (3), 295-303. Farrant, R. D., Lindon, J. C., and Nicholson, J. K. (1994) Internal temperature calibration for 1H NMR spectroscopy of blood plasma and other biofluids. NMR Biomed. 7, 243-247. Holmes, E., Foxall, P. J. D., Nicholson, J. K., Neild, G. H., Brown, S. M., Beddell, C. R., Sweatman, B. C., Rahr, E., Lindon, J. C., Spraul, M., and Neidig, P. (1994) Automatic data reduction and pattern recognition methods for analysis of 1H nuclear magnetic resonance spectra of human urine from normal and pathological states. Anal. Biochem. 220, 284-296. Beebe, K. R., Pell, R. J., and Seahsholt, M. B. (1998) Chemometrics: A Practical Guide, John Wiley & Sons, New York. Osada, J., Aylagas, H., Miro-Obradors, M. J., and Palacios-Alaiz, E. (1993) Alterations in lipid characteristics of lysosomes are involved in liver necrosis induced by thioacetamide. Biol. Chem. 374, 129-132.

654


(36) Waterfield, C. J., Turton, J. A., Scales, M. D. C., and Timbrell, J. A. (1993) Investigations into the effects of various hepatotoxic compounds on urinary and liver taurine levels in rats. Arch. Toxicol. 67, 244-254. (37) Timbrell, J. A., Waterfield, C. J., and Draper, R. P. (1995) Use of urinary taurine and creatine as biomarkers of organ dysfunction and metabolic perturbations. Comput. Haematol. Int. 5, 112-119. (38) Wyss, M., and Kaddurah-Daouk, R. (2000) Creatine and creatinine metabolism. Physiol. Rev. 80, 1107-1209. (39) Ogborn, M. R., Sareen, S., Prychitko, J., Buist, R., and Peeling, J. (1997) Altered organic anion and osmolyte content and excretion in rat polycystic kidney disease: An NMR study. Am. J. Physiol. 272, F63-F69. (40) Bagnasco, S., Balaban, R., Fales, H. M., Yang, Y.-M., and Burg, M. (1986) Predominant osmotically active organic solutes in rat and rabbit renal medullas. J. Biol. Chem. 261, 5872-5877. (41) Burg, M. B. (1997) Renal osmoregulatory transport of compatible organic osmolytes. Curr. Opin. Nephrol. Hypertens. 6, 430-433. (42) Cascales, M., Martin-Sanz, P., Alvarez, A., Sanchez-Perez, M., Diez-Fernandez, C., and Bosca, L. (1992) Isoenzymes of carbohydrate metabolism in primary cultures of hepatocytes from thioacetamide-induced rat liver necrosis: Responses to growth factors. Hepatology 16, 232-240. (43) Anghileri, L. J. (1976) Metabolism of calcium and magnesium in liver during acute thioacetamide intoxication. Int. J. Clin. Pharmacol. 14, 101-108. (44) Hunter, A. L., Holscher, M. A., and Neal, R. A. (1977) Thioacetamide-induced hepatic necrosis I: Involvement of the mixedfunction oxidase enzyme system. J. Pharmacol. Exp. Ther. 200, 439-448. (45) Mesa, M. L., Carrizosa, R., Martinez-Honduvilla, C., Benito, M., and Fabregat, I. (1996) Changes in rat liver gene expression induced by thioacetamide: Protective role of S-adenosyl-L-methionine by a glutathione-dependent mechanism. Hepatology 23, 600-606. (46) Zimmermann, T., Franke, H., and Dargel, R. (1986) Studies on lipid and lipoprotein metabolism in rat liver cirrhosis induced by different regimens of thioacetamide administration. Exp. Pathol. 30, 109-117. (47) Moreira, E., Fontana, L., Periago, J. L., De Medina, F. S., and Gil, A. (1995) Changes in fatty acid composition of plasma, liver microsomes and erythrocytes in liver cirrhosis induced by oral intake of thioacetamide in rats. Hepatology 21, 199-206. (48) Kumar, A., and Rana, S. V. S. (1982) Lipid accumulation in chromium poisoned rats. Int. J. Tissue React. 4, 291-295. (49) Holub, B. J. (1982) The nutritional significance, metabolism and function of myo-inositol and phosphatidylinositol in health and disease. Adv. Nutr. Res. 4, 107-141. (50) Holub, B. J. (1986) Metabolism and function of myo-inositol and inositol phospholipids. Annu. Rev. Nutr. 6, 563-597. (51) Zeisel, S. H., Gettner, S., and Youssef, M. (1989) Formation of aliphatic amine precursors of N-nitrosodimethylamine after oral administration of choline analogues in the rat. Food Chem. Toxicol. 27, 31-34. (52) Tjoa, S., and Fennessey, P. (1991) The identification of trimethylamine excess in man: Quantitative analysis and biochemical origins. Anal. Biochem. 197, 77-82. (53) Smith, J. L., Wishnok, J. S., and Deen, W. M. (1994) Metabolism and excretion of methylamines in rats. Toxicol. Appl. Pharmacol. 125, 296-308. (54) Fontana, L., Moreira, E., Torres, M. I., Fernandez, M. I., Rios, A., De Medina, F. S., and Gil, A. (1996) Serum amino acid changes in rats with thioacetamide-induced liver cirrhosis. Toxicology 106, 197-206. (55) Schanbacher, F. L., Willett, L. B., and Moorhead, P. D. (1981) Ornithine decarboxylase, serum isocitrate dehydrogenase and clinical chemistry changes during thioacetamide-induced hepatotoxicity in a calf. J. Anim. Sci. 53, 1658-1670. (56) Kameji, T., Murakami, Y., Fujita, K., and Hayashi, S.-I. (1982) Purification and some properties of ornithine decarboxylase from

Waters et al. rat liver. Biochim. Biophys. Acta 717, 111-117. (57) Yurdaydin, C., Swain, M. G., Mininberg, E. D., Vergalla, J., Kleiner, D., Paul, S. M., and Jones, E. A. (1998) Effect of inhibition of ornithine decarboxylase activity in a model of acute hepatocellular necrosis. Eur. J. Gastroenterol. Hepatol. 10, 503-507. (58) Domschke, S., and Domschke, W. (1972) Polyamines and the liver. Acta Hepato-Gastroenterol. 19, 212-217. (59) Aigner-Held, R., and Daves, G. D. (1980) Polyamine metabolites and conjugates in man and higher animals: A review of the literature. Physiol. Chem. Phys. 12, 389-400. (60) Grillo, M. A. (1985) Metabolism and function of polyamines. Int. J. Biochem. 17, 943-948. (61) Antoniello, S., Auletta, M., Magri, P., and Pardo, F. (1998) Urinary excretion of free and acetylated polyamines in hepatocellular carcinoma. Int. J. Biol. Markers 13, 92-97. (62) Krebs, H. A., Hems, R., and Tyler, B. (1976) The regulation of folate and methionine metabolism. Biochem. J. 158, 341-353. (63) Chanarin, I. (1985) Folate and cobalamin. Clinics Haematol. 14, 629-641. (64) Davis, R. E. (1986) Clinical chemistry of folic acid. Adv. Clin. Chem. 25, 233-294. (65) Meister, A., and Anderson, M. E. (1983) Glutathione. Annu. Rev. Biochem. 52, 711-760. (66) Pumpo, R., Sarnelli, G., Spinella, A., Budillon, G., and Cuomo, R. (2001) The metabolism of nicotinamide in human liver cirrhosis: A study on N-methylnicotinamide and 2-pyridone-5carboxamide production. Am. J. Gastroenterol. 96, 1183-1187. (67) Soares Melo, S. S., Meirelles, M. S., Jordao, A. A., and Vannucchi, H. (2000) Lipid peroxidation in nicotinamide-deficient and nicotinamide-supplemented rats. Int. J. Vitam. Nutr. Res. 70, 321323. (68) Matsui, I., and Pegg, A. E. (1980) Effect of thioacetamide, growth hormone or partial hepatectomy on spermidine acetylase activity of rat liver cytosol. Biochim. Biophys. Acta 633, 87-94. (69) Porter, W. R., Gudzinowicz, M. J., and Neal, R. A. (1979) Thioacetamide-induced hepatic necrosis II: Pharmacokinetics of thioacetamide and thioacetamide-S-oxide in the rat. J. Pharmacol. Exp. Ther. 208, 386-391. (70) Dyroff, M. C., and Neal, R. A. (1982) Studies of the mechanism of metabolism of thioacetamide-S-oxide by rat liver microsomes. Mol. Pharmacol. 23, 219-227. (71) Gavaghan, C. L., Nicholson, J. K., Connor, S. C., Wilson, I. D., Wright, B., and Holmes, E. (2001) Directly coupled high performance liquid chromatography and nuclear magnetic resonance spectroscopic with chemometric studies on metabolic variation in Sprague-Dawley rats. Anal. Biochem. 291, 245-252. (72) Martin, M. E., Karoum, F., and Wyatt, R. J. (1979) Phenylacetic acid excretion in man. Anal. Biochem. 99, 283-287. (73) Jones, A. R. (1982) Some observations on the urinary excretion of glycine conjugates by laboratory animals. Xenobiotica 12, 387395. (74) Linhart, I., and Smejkal, J. (1987) Changes in the excretion of endogenous glycine conjugate as a possible artifact in toxicological experiments. Arch. Toxicol. 61, 83-85. (75) Phipps, A. N., Wright, B., Stewart, J., and Wilson, I. D. (1997) Use of proton NMR for determining changes in metabolite excretion profiles induced by dietary changes in the rat. Pharm. Sci. 3, 143-146. (76) Phipps, A. N., Stewart, J., Wright, B., and Wilson, I. D. (1998) Effect of diet on the urinary excretion of hippuric acid and other dietary-derived aromatics in rat. A complex interaction between diet, gut microflora and substrate specificity. Xenobiotica 28, 527537. (77) Goodwin, B. L., Ruthven, C. R. J., and Sandler, M. (1994) Gut flora and the origin of some urinary aromatic phenolic compounds. Biochem. Pharmacol. 47, 2294-2297.

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Metabonomic Deconvolution Of Embedded Toxicity - American

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