Testicular Toxicity of Candidate Fuel Additive 1,6-Dimethoxyhexane: Co
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《毒物学科学杂志》
Systemic Toxicology and Pharmacokinetics Section, Environmental and Occupational Toxicology Division, Environmental Health Science Bureau, Health Canada, Ottawa, Ontario, K1A 0L2, Canada
ABSTRACT
Previous studies of the toxicity of candidate fuel additives identified severe testicular toxicity in animals exposed to 1,6-dimethoxyhexane (DMH). A series of studies were conducted to characterize the effects of DMH on spermatogenesis and to compare the effects of DMH with responses to structural similar aliphatic ethers. In the first study, sexually mature male rats were administered a single dose (600 mg/kg) of DMH, and subsets of animals were sampled at intervals post exposure (PE). Both testis and thymus weight declined steadily after DMH exposure, both being significantly lower than control by 7 days PE. Treatment with DMH led, at 24 to 48 h PE, to an increase in dying primary spermatocytes in seminiferous tubule stages I–IV and stages XII–XIV, but not intervening stages. The affected cohort of germ cells was seen progressing through the developmental sequence of spermatogenesis as numbers of dying cells returned to control levels by 7 days PE, coincident with a significant decline in the proportion of round spermatids among germ cells as determined by flow cytometry. Resolution of round spermatids to control levels by day 21 PE coincided with a reduction in condensed spermatids (homogenization-resistant spermatid nuclei) and was followed at 28 days PE by a significant reduction in cauda epididymal sperm counts. Further studies of repeated exposures to DMH (200 mg · kg–1 · day–1, 5 days per week for 4 weeks) confirmed the significant testis toxicity of this material. In contrast, similar treatment with any of a variety of structurally similar aliphatic ethers had little or no impact on testis function. Methoxyacetic acid, previously shown to cause rapid death of some meiotic germ cell stages, was found at high concentrations in the urine of DMH-treated rats but not in rats treated with other ethers, suggesting that DMH exerts its testis toxicity via metabolism to this substance. These results demonstrate that DMH selectively deletes germ cells from the testis At the very early or very late pachytene, diplotene, or M-phase spermatocyte stages, possibly through conversion to MAA.
Key Words: fuel additives; testis toxicity; spermatogenesis; methoxyacetic acid; pachytene spermatocytes.
INTRODUCTION
Diesel fuels are important energy sources in transport and industry. The combustion of diesel, however, produces massive quantities of air pollutants, including carbon monoxide, nitrogen oxides, volatile organics, and exhaust particles and particulates. There is growing realization that these pollutants contribute to death and morbidity due to cardiovascular and respiratory disease, among other health impacts, and that they are therefore very significant health concerns (Burnett et al., 1998; Goldberg et al., 2000; Villeneuve et al., 2002). The addition of oxygenated substances, such as liquid ethers, into diesel has been shown to improve combustion efficiency. Furthermore, the presence of oxygenates has been reported to reduce the emission of exhaust particulates, carbon monoxide, and total hydrocarbons (Zhu et al., 2003; Shen et al., 1993; Heinz, 1993).
Our laboratory has undertaken toxicological studies of various aliphatic ethers in order to identify compounds with low toxicity for use as diesel additives (Poon, et al., 2000; 2004; in press). These studies revealed that repeated oral exposure to 1,6-dimethoxyhexane (DMH) causes significant reduction in testis weight to about 70% of control, among other significant toxic effects (Poon et al., 2004). Histological examination of the testes and epididymis of DMH-exposed animals found many seminiferous tubules lacking normal spermatids, increased numbers of spermatid giant cells, and apparently a reduced density of mature sperm. In contrast, eight other aliphatic ethers examined in these studies had no significant effect on testis. It was postulated that the testicular effects of DMH were caused by its metabolism to methoxyacetic acid (MAA) (Poon, 2004). This hypothesis was supported by a subsequent study wherein MAA was detected in the blood and urine of DMH-treated animals (Poon et al., 2005). In the present study, we extend initial observations of a testicular effect in DMH by examining multiple testicular end points in animals exposed to DMH or to any of eight other structurally similar ethers. These end points include the relative proportion of round spermatids, number of homogenization-resistant sperm nuclei, cauda epididymal sperm morphology, and testis lactate dehydrogenase X (LDH-X) activity. To further understand the mechanisms and time course of action of DMH, acute dosing studies were performed which examined the progression of DMH-induced testis lesions and recovery.
MATERIALS AND METHODS
Chemicals and reagents.
1,6-Dimethoxyhexane was synthesized by Wychem (Suffolk, UK) and had a purity of >99% as measured by gas chromatography. Syntheses of 1,1-dimethoxyoctane (1,1-DMO), 1-ethoxyheptane (EHpE), 1- ethoxyhexane (EHxE), 1-isopropoxyoctane (IPO), 1-methoxyheptane (MHpE), 1-methoxyoctane (MOE), and 1-proxyhexane (PHxE) were carried out by M. S. Kaliaguine (Universite Laval, Quebec, PQ) and all had a purity >97% as measured by gas chromatography. Dibutyl ether (BE) was purchased from Aldrich Chemical (Milwaukee, WI) and had a purity of >99%. Nonyl-acridine orange (NAO) and propidium iodide (PI) were purchased from Molecular Probes (Eugene, OR). In situ apoptosis detection kit was purchased from R&D Systems (Minneapolis, MN). Periodic acid–Schiff histological staining reagents were purchased from Richard Allan Scientific (Kalamazoo, MI). All other chemicals are of reagent grade and were obtained from either Fisher Scientific (Nepean ON, Canada) or Sigma Chemical Co. (St Louis, MO).
Animals and experimental approach.
Sexually mature Sprague-Dawley rats, (8–9 weeks of age for subacute studies (see below) or 11–12 weeks for acute study) were purchased from Charles River Canada (St. Constant, PQ) and were routinely housed in pairs in polycarbonate hanging cages, open to room air through a micro-barrier filter, with free access to water and laboratory rat chow (Purina 5008; Agribrand Purina; Saint-Hubert, Quebec) under controlled photoperiod (12:12 light:dark), humidity (40–70%), and temperature (20–24°C). All animal handling procedures adhered to Canadian Council on Animal Care guidelines and were approved by the Health Canada Animal Care Committee prior to the initiation of the study.
The results of four separate animal studies are described in this article. The first study examined the progression of lesion and recovery of the testis and the reproductive tract after a single oral dose of DMH (acute study). The remaining studies were designed to examine the effects of repeated doses of any of a variety of aliphatic ethers, along with a group receiving an effective dose of DMH, for 4 weeks, on an array of end points indicating reproductive toxicity (subacute studies). The full results of these latter studies, beyond the effects on the reproductive tract described herein, will be reported elsewhere.
For the acute study, animals were administered a single dose of DMH (600 mg/kg in 5 ml/kg corn oil) by gavage, and groups of 7–8 animals were sacrificed 4 h, 1, 2, 3, 7, 14, 21, or 28 days post exposure (PE). An additional control group of 8 animals received 5 ml/kg corn oil and were sacrificed 24 h PE.
Three separate studies examined the general and reproductive consequences of 4 weeks of exposure (200 mg · kg–1 · day–1, 5 days per week for 4 weeks) to BE, EHxE, and MHpE (Experiment 1); EHpE, MOE, and PHxE (Experiment 2); and IPO and 1,1-DMO (Experiment 3). In each of these studies, a separate group of rats treated with DMH were used as the comparative group. The details of animal treatment and urine collection are described elsewhere (Poon et al., 2005). Briefly, animals were randomized into treatment groups of 7 animals each during a 1-week acclimation phase. Animals received, by oral gavage, 200 mg/kg of one of the 9 alkyl ethers in corn oil, 5 days per week for 4 weeks. Control animals received corn oil only. Animals were sacrificed beginning 24 h after the final exposure.
Tissue collection.
Animals were sacrificed by exsanguination under isofluorane anesthesia. Routinely, the testes and epididymis were rapidly removed, dissected clear of fat, and weighed. The tunica albuginia of the right testis was perforated 20 times with a 20-gauge needle and fixed by immersion in modified Davidson's solution (Latendresse et al., 2002) for subsequent analysis of stage-specific germ cell apoptosis. The left testis was decapsulated, and the parenchyma was divided into four roughly equivalent portions by sagittal then transverse cuts through the midline of the testis. One portion was placed in ice-cold medium 199 (M199 containing 0.1% BSA) for analysis of spermatogenic cell populations. Two portions were weighed, separately frozen on dry ice, and stored at –20°C prior for analysis of homogenization-resistant sperm nuclei and testis-specific lactate dehydrogenase (LDH) isoform, LDH-X. Several small incisions were made in the left cauda epididymis, and it was placed into 10 ml of warmed Dulbecco's phosphate buffered saline (PBS) (Gibco BRL, Bethesda, MD) to allow sperm to swim out. One 200-μl sample of sperm was stained with eosin and a portion smeared on a microscope slide, dried, and cover-slipped to examine sperm morphology. The remaining media containing sperm and the cauda were stored at –20°C for analysis of cauda sperm reserves.
Testis histology and analyses of apoptotic germ cells.
Germ cell death was examined in five animals each, from control, 4 h, 1, 2, 3, and 7 days post exposure groups of the first study only. Testes were fixed in modified Davidson's solution for 72 h and washed twice in PBS (48 h total), after which a central transverse section was embedded in paraffin and sectioned at 5 μm. One section was stained with periodic acid–Schiff to highlight acrosome structure to assist in seminiferous tubule staging (Hess, 1990), and the adjacent section was processed for in situ DNA end-labeling (TUNEL labeling) to identify apoptotic cells with the TdT In Situ Apoptosis Detection Kit–DAB (daminobenzidine; R & D Systems, Minneapolis, MN). Stages of seminiferous tubule cross sections were determined by previously described criteria (Hess, 1990).
Testis cell isolation and analysis.
Fresh testes were dispersed using a two-stage enzymatic digestion as previously described (Bellve et al., 1977). Testes were incubated in 2.5 ml M199 1% BSA, containing 56 U/ml collagenase type III (Worthington Biochemical, Freehold, NJ) and 8 U/ml DNAase I (Sigma, St Louis, MO) at 32°C in a shaking water bath for 15 min. Tubes were then transferred to ice and, after settling for 1 min, the supernatant, containing interstitial cells, was aspirated and discarded. The remaining seminiferous tubules were washed twice with 10 ml of M199 0.1% BSA and then incubated as above in 5 ml of M199 containing 110 U/ml trypsin (Worthington) and 10 U/ml DNAase I. Tubes were then placed on ice and gently aspirated repeatedly with a Pasteur pipette to disrupt cell aggregates, after which 10% BSA stock was added (final concentration of 0.5% BSA) and mixed by aspiration through the Pasteur pipette. The cell suspension was then filtered through a 70-μm nylon mesh (Filicon, Dako Canada, Mississauga, ON), centrifuged (400 x g for 8 min at 4°C), washed twice with 10 ml ice-cold DPBS containing 0.5% BSA (PBS-BSA; pH 7.4) and fixed in ice-cold 70% ethanol. Fixed cells were kept at –20°C for no more than 30 days prior to analysis of spermatogenic cell populations.
Relative proportions of spermatogenic cell populations were assessed in fixed cells using a flow cytometric method (Petit et al., 1995; Suter et al., 1997). The principle of the test is that spermatogenic cells, as they differentiate from normal diploid spermatogonial stem cells through to mature spermatozoa with a highly condensed haploid complement of DNA, progress through various intermediate stages with differing nuclear DNA content and cellular content of mitochondria. With the use of fluorescent probes to quantitatively label DNA (PI) and mitochondria (NAO), the relative numbers of cells in each intermediate population can be resolved to determine relative numbers at each germ cell stage. Cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences, Mississauga, ON) fitted with an argon ion laser (488 nm line excitation); fluorescence emission of NAO was reflected by a 550 dichroic longpass filter and quantified after passage through a 530/30 nm bandpass filter. Fluorescence of PI was detected after sequential passage through the 550 dichroic filter and a 670-nm longpass filter. Relative proportions of cells in each population were calculated with WinList software (Verity Software, Topsham, ME). The germ cell types in each population were identified under a fluorescent microscope (Zeiss Axioskop 2, Carl Zeiss, Thornwood, NY) by examination of cells from each population, which were isolated using the cell-sorting capacity of the FACSCalibur flow cytometer.
Cauda epididymal sperm morphology.
Assessment of sperm morphology followed the guidance of an expert consensus statement (Seed et al., 1996). Briefly, eosin-stained sperm collected from the cauda epididymis were smeared onto two glass slides per sample, air-dried, and cover-slipped. At least 200 sperm on each slide were examined for the proportion of sperm with abnormal head (overhooked, blunt hook, banana-shaped, amorphous, or extremely oversized) or tail (twisted, bent, corkscrew, double/multiple) by one individual unaware of animal number or treatment. Values for individual slides were averaged for each animal.
Homogenization-resistant spermatid nuclei (HRSN) and cauda epididymis sperm reserves.
The total number of condensed spermatids per testis and the total number of sperm in the cauda epididymis were estimated according to a previously described method (Blazak et al., 1985). The principle of this method relies on the extreme stability of the nuclei from stage 17–19 spermatids or cauda sperm, which allows counting of these after complete homogenization and sonication of the tissue. To determine testis condensed spermatid number, a weighed portion of the parenchyma from the left testis, as representative of the whole organ as possible, was homogenized in 20 ml of STA solution (0.9 % NaCl, 0.01% Triton X-100, and 0.025% sodium azide) in a 50-ml Waring Ultramicro blender (Fisher Scientific, Nepean, Ont.) on low setting for 2 min. Similarly, whole cauda and associated sperm suspension in DPBS were thawed on ice and, after addition of 200 μl of 1% Triton X-100 (final concentration of 0.01%) and brought up to 20 ml volume as STA, were homogenized as above. Homogenates of either tissue were then disrupted with a VibraCell sonicator (Sonics & Materials, Danbury, CT) using a microprobe on setting 7 for 60 s. Sperm or HRSN densities were calculated from the average number of nuclei in four fields of a Neubaur hemocytometer and were expressed as total or as per gram of epididymis or testis weight.
Analysis LDH and LDH-X.
Testis fragments were thawed slightly and then homogenized in 2 ml/g of ice-cold 12.5 mM TRIS KCl (pH 9.0) with 5 strokes of a manual homogenizer (Kontes Glass Company, Vineland, NJ). The homogenate was centrifuged at 19,000 x g for 10 min at 4°C and the supernatant was diluted fivefold in TRIS KCl. Separate aliquots were used to determine total LDH and LDH-X. Enzyme activity was measured by monitoring extinction of NAD absorbance at 340 nm in a reaction mixture that contained 4.2 mM of NAD in 10.5 mM Tris KCl (pH 9.0) at 30°C. After background NAD conversion was monitored for 30 s, the appropriate substrate (lactic acid for total LDH, alpha-hydroxycaproic acid for LDH-X) was added to a final concentration of 50 mM and, after brief mixing (2 s), 340 nm extinction was monitored for a further 30 s. Enzyme activity was calculated from the linear region of NAD conversion, corrected for background rate and converted to μmoles converted · (min)–1 · (mg homogenate protein)–1. Total protein in testis homogenates was determined with the modified micro-Lowry protein determination kit (Sigma, St Louis, MO).
Analysis of 2-methoxyacetic acid.
After receiving their final gavage dose, animals were transferred to metabolic cages and provided with food and water ad libitum. The entire volume of urine produced over 24 h was collected in a 50-ml conical tube and held on wet ice until the end of the collection period. The sample was then stored at –20°C until analysis. The method for analysis of the 2-methoxyacetic acid (MAA) was set up in this laboratory and will be reported elsewhere in detail (Li et al., manuscript in preparation). Solid-phase extraction of urine was carried out with a Supelco 12-port Visiprep SPE Vacuum Manifold. Before sample application, 500 mg Bakerbond C18 cartridges were conditioned with 3 ml of methanol and 5 ml of water applied sequentially, after which 1 ml of urine sample was acidified with 100 μl of concentrated HCl and applied to the cartridges with a Pasteur pipette and allowed to flow through by gravity. After sample application, the cartridges were centrifuged at 3000 rpm for 5 min, placed on the SPE vacuum manifolds, and air-dried for 15 min with an air stream generated from a vacuum pump. Each of the C18 cartridges was then connected to a Bond ElutT Na2SO4 cartridge (1.4 g; Varian, Palo Alto, CA). The serially connected cartridges were eluted with 2 ml of 2.5% isopropanol in diethyl ether. The residue water was dried while the eluant was flowing through the Na2SO4 cartridge.
An internal standard of 2-bromopropoinic acid (BrPA; 99%, Supelco, Bellefonte, PA), prepared in methyl-tert-butyl ether (MTBE) at 0.25 mg/ml was added (100 μl) to the eluant, and the solution was concentrated at ambient temperature to a final volume of 50 μl with a gentle stream of nitrogen. The concentrates were reconstituted with hexane to a volume of 200 μl. Derivatization with diazomethane was conducted using a modification of USEPA method 515.1 Briefly, the diazomethane generated from diazald and 37% KOH was bubbled through 1/16'' polytetrafluoroethylene (PTFE) tubing into the hexane solution with the aid of a gentle stream of nitrogen at a rate of 10 ml/min until the hexane solution turned yellowish. Despite the very small volumes of diazomethane used, great care was taken to avoid human exposure to this toxic substance. All the work was conducted in a well-ventilated fume hood, and a long tweezers was used to manipulate the outlet of PTFE tubing into the sample solution. The outlet of the PTFE tubing was rinsed with hexane between samples to avoid cross contamination. After 30 min, the diazomethane in sample was destroyed by addition of silicic acid. The GC/MS analysis was performed with a Thermo Finnigan Trace 2000 gas chromatograph coupled to a PolarisQ mass spectrometer. The instrument was equipped with an AS 2000 liquid autosampler, a Supelco OMEGAWAX 250 fused silica capillary column (30 m x 0.25 mm x 0.25 μm), and the Xcalibur data system. The oven temperature was programmed at the initial temperature of 50°C, held for 2 min, and then increased at a rate of 2°C/min to 120°C. The oven was held at this temperature for 1 min, then increased at 30°C/min to the temperature of 220°C, which was held for 3 min. Helium was the carrier gas; it was used with a flow rate of 0.8 ml/min and the split flow was 32 ml/min. A 2-μl volume of sample solution was injected with a splitless injection mode and 1.0-min purge time. The temperatures of the injection port, transfer line, and MS ion source were set at 220°C, 250°C, and 200°C, respectively. The selected ion monitor (SIM) was used for quantification. The method is validated and can be employed to detect the MAA for a concentration as low as 2 ng/ml (ppb).
Statistical analyses.
After successful testing for assumptions of normal distribution (Shapiro-Wilks Goodness of Fit Test) and homoscedacity (Bartlett's test) all data were analyzed by analysis of variance (ANOVA). Proportional data (testis cell data, percent abnormal sperm cells) were arcsine transformed prior to analysis to normalize. Where normality or equal variance were not indicated, data were retested after log transformation and, if normality or equal variance assumptions were not satisfied, non-transformed data were analyzed for ether treatment effects using ANOVA on ranks (Wilcoxon-Kruskal-Wallis test). Data for which ANOVA identified a significant effect of ether treatment (p < 0.05) were subjected to Dunnett's or Dunn's post hoc analysis, for parametric or non-parametric ANOVA results, respectively, to identify effective treatments. All statistical analyses were conducted with JMP software, release 4.0.2 (SAS Institute, Cary, NC) except for the nonparametric ANOVA and Dunn's tests, which were performed using SigmaStat, release 2.03 (SPSS Inc., Chicago, IL).
RESULTS
DMH Effects on Tissue Weights and Germ Cell Death
To examine the progression of DMH-induced testis toxicity, male rats were administered a single dose of DMH (600 mg/kg) and were sacrificed at various times post-exposure. Seven days after DMH exposure, relative testis and thymus gland weights were significantly reduced and continued to decline up until the final sampling 28 days PE. In contrast, epididymis weights were not significantly influenced by DMH exposure (Table 1).
The reduction in testis weight 7 days PE, as well as previous observations of DMH-induced atrophy of the seminiferous epithelium (Poon et al., 2004), suggested that DMH exposure caused disruption of spermatogenesis. To evaluate the impact of DMH on germ cell survival, testis sections were stained for DNA fragmentation indicative of cell death using the TUNEL reaction (terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling: Gavrieli et al., 1992). Seminiferous tubules of control animals had very few TUNEL-positive cells, indicating very low level germ cell attrition in normal testes. The number of TUNEL-positive cells was similar to control levels at 4 h PE, but it increased dramatically at 1 and 2 days PE (Fig. 1, Fig. 2A). Positive cells were still significantly elevated at 3 days PE, to a lesser extent than at 1 and 2 days PE, but they returned to control levels by 7 days PE. Stage-specific analyses of TUNEL-positive cells in seminiferous tubules at 1 and 2 days PE demonstrated that DMH-induced cell death is highly specific to pachytene spermatocytes in stages I–IV and to late pachytene, diplotene, and meiotic primary spermatocytes in stages XII–XIV, whereas pachytene spermatocytes intermediate to these stages are unaffected (Fig. 1, Fig. 2B). The early pachytene spermatocytes (in stage I–IV tubules) seemed to be initially more susceptible to DMH-toxicity, as there was a significantly greater number of TUNEL-positive cells in this group at 24 h PE versus the same group at 48 h PE or stage XI–XIV stage tubules at 24 h PE (Fig. 2B). At 48 h PE, there were more TUNEL-positive cells in the stage XI–XIV group than in stage I–IV tubules at the same time PE or than were seen in stage XI–XIV tubules 24 h PE.
DMH Effects on Spermatogenesis
To further evaluate the effects of DMH on spermatogenesis, the relative proportion of round spermatids was evaluated using a flow cytometric method that identified populations of testicular cells based on PI (indicating DNA) and NAO (mitochondrial) content. Representative scattergrams of testis cells isolated from a control animal (Fig. 3A) and 14 day PE animal (Fig. 3B) are provided, demonstrating the pattern characteristic of these analyses. Although six separate populations, identified by fluorescent microscopic analyses of sorted cells, could be evaluated using this method, only data for round spermatids are presented. The data for other populations tended to be highly variable, with coefficients of variability in excess of 40% (data not shown). As expected, round spermatids constituted the most prominent population of germ cells, representing 53.9 ± 3.5% of all cells in preparations from testes of control animals (Fig. 4). Exposure to DMH caused a latent reduction in the relative proportion of round spermatids that was significant by 7 days PE, and this population reached a minimum level 14 days PE (37.8 ± 8.7%) but returned to levels similar to control thereafter.
The progression of the affected cohort of germ cells through the successive stages of spermatogenesis was also demonstrated by the number of condensed spermatids (homogenization-resistant sperm nuclei [HRSN]; Table 2). The DMH-induced reduction in HRSN appeared 1 week after the DMH-induced reduction of round spermatid levels. Mean HRSN, expressed either as total for the left testis or per gram of testis parenchyma, were significantly reduced by 21 days PE or remained at this lower level at 28 days PE (Table 2). Cauda epididymal sperm reserves per gram of cauda weight were also significantly reduced 28 days PE, but not at earlier times, further indicating the progression of the effected cohort of germ cells through the reproductive tract.
Effects of Repeated Doses of DMH versus Other Ethers
A further series of experiments was conducted to compare the toxicity of DMH with that of several structurally similar types of ether. The effects of these ethers on spermatogenesis and cauda sperm are presented below. The effects of oral exposure to various aliphatic ethers for 5 days per week for 4 weeks on body weight, weights and histopathological analyses of major organs, measures of serum biochemistry, hematology, and some liver enzymes from these studies will be described elsewhere (Poon et al., 2005; Poon and Chu, unpublished). In all of these studies, only DMH caused a significant reduction in relative testis weight (i.e., g testis weight/100 g body weight) with a 21.4%, 16.7%, and 29.6% reduction from relative control testis weight in the first, second, and third studies, respectively (data not shown).
To determine the impact on germ cell survival, the effect of ether treatment on the proportion of round spermatids among testis cells was examined for all animals (see Table 4). In all three of the 4-week studies, DMH treatment led to a significant reduction in the proportion of round spermatids to 63%, 64%, and 58% of control levels in Exeriments 1, 2, and 3, respectively (Table 3). Of all other ethers examined, only MOE had any effect on round spermatid number, causing a significant reduction of round spermatids to 83.8% of control levels (p < 0.05).
Testis sperm production, as indicated by HRSN, was significantly reduced in animals exposed to DMH, yielding HRSN numbers that were 49%, 45%, and 71% of total left testis HRSN of control animals in the three experiments (Table 4). Like testis sperm production, epididymal sperm reserves were reduced by DMH treatment, causing a 37%, 43%, and 31% reduction in absolute numbers per left cauda epididymis for the first, second, and third studies, respectively, although the effect in the third experiment did not reach statistical significance (Table 4). In contrast to DMH, most of the other ethers were without effect on either testis HRSN or cauda sperm counts. The exceptions to this was BE, which caused a slight but significant reduction in HRSN (p < 0.05).
Sperm morphology was examined as a measure of spermatid final maturation (Table 5). Treatment with DMH led to a significant reduction in the proportion of morphologically normal sperm (p < 0.05) in all three studies, although the degree of effect differed between studies ranging from roughly a 50% decline in normal in the first study (42.5 ± 17.6% normal sperm in DMH versus 94.5 ± 2.3% in control) to roughly 13% for the other two studies. In all three studies there was a marked increase in the proportion of morphologically abnormal sperm heads, indicating a possible effect of DMH on spermatid nuclear condensation. In contrast with the effects of DMH, sperm morphology was unaffected by treatment with any other ether tested in these studies (Table 5: p > 0.05).
To further evaluate testicular toxicity, the activity of a germ-cell–specific LDH isoform, LDH-X was evaluated. Activity of total LDH and LDH-X specific activity were measured in testis homogenate using lactic acid and -hydroxycaproic acid, as the substrate. Although total testicular LDH activity was not influenced by any ether treatment, the activity of LDH-X was reduced by DMH treatment to 77.0%, 75.3%, and 85.9% of control values for the first, second and third studies, respectively, although the effect in the third experiment did not reach statistical significance (Table 6). These results are consistent with the observed reduction in round spermatids where this isoform is expressed (Hintz and Goldberg, 1977; Meistrich et al., 1977). Activity of LDH-X was not significantly influenced by any other ether treatment.
Urinary MAA Levels in DMH-Exposed Animals
As the pattern of germ cell death observed in DMH-treated animals 1 and 2 days PE was remarkably similar to that observed in animals exposed to 2-methoxyacetic acid (Foster et al., 1987), we hypothesized that DMH-induced germ cell toxicity may be consequent to metabolism of this compound. To test this hypothesis, we compared MAA residue levels in urine from DMH-treated animals from experiment 1 of the subacute studies with those in the urine collected from control animals or animals exposed to ethers that caused no testis toxicity. Methoxyacetic acid was slightly above the limit of detection in all samples, including those from vehicle control animals, suggesting some transfer of the volatile parent compound from dosed animals. However, urine collected from DMH-treated animals contained MAA levels that were, on average, 230-fold greater than that found in control animals (Table 7). Levels of MAA detected in urine from MHpE-treated animals appeared to be higher than controls, but they did not reached a statistically significant level.
DISCUSSION
The present study demonstrates that subacute oral exposure to the aliphatic diether DMH causes significant disruption of spermatogenesis and impaired maturation of sperm. This confirms and extends the findings of a previous study showing that subacute oral exposure to DMH caused a dose-related decrease in testis weight, degeneration of seminiferous epithelium, among other histopathological lesions in the testis, and increased presence of abnormal sperm in the epididymal tubules (Poon et al., 2004). Here we have demonstrated that DMH exposure results in death of pachytene spermatocytes at very precise stages of differentiation, with consequent reduction in relative numbers of round spermatids, numbers of HRSN, and epididymal sperm reserves. In conjunction with the observed reduction in sperm production, repeated DMH exposure caused an increased proportion of sperm with abnormal morphology, demonstrating a toxicant-induced impairment in the production of normal spermatozoa.
Our data demonstrate that a single oral exposure to DMH causes a sharp increase in germ cell death at 24 and 48 h PE, with a subsequent reduction in germ cell death to control levels by 7 days PE. We also demonstrate that DMH toxicity is highly specific to early pachytene spermatocytes in tubule stages I–IV and to late pachytene spermatocytes, diplotene spermatocytes, or meiotically dividing spermatocytes in tubule stages X–XIV. Other than these cells, there was no increase in TUNEL-positive cells in the intervening stages, nor was there any evidence of dying spermatids or spermatogonia in any tubule at any stage. The reduction in germ cell death to control levels by 7 days PE, suggesting that DMH-induced effects on spermatogenesis are transient, is supported by the transient reduction in the number of round spermatids observed on PE days 7 and 14 but not on PE days 21 or 28. The effects seen in the single-dose study are largely confirmed by the results of the repeated dose studies, with one exception: The significant (p < 0.05) increase in sperm with abnormal head morphology was consistently observed in DMH-treated animals in the subacute studies but was very minimal in samples from the single-dose study, suggesting that repeated exposure to DMH has a greater impact on events in spermiogenesis, spermiation, or cauda epididymal function than occurs in response to a single exposure.
In addition to reducing sperm production and cauda epididymal sperm reserves, exposure to multiple doses of DMH for 4 weeks also caused an increase in the proportion of sperm with abnormal morphology. As abnormal morphology of sperm head has been shown to be a good indicator of reduced fertility in both animal studies (e.g., Ballachey et al., 1987) and human studies (e.g., Bonde et al., 1998; Kruger et al., 1988), such observations suggest that DMH causes a reduction in both epididymal sperm numbers and quality.
The above results, together with earlier observations, provide a close glimpse into the mechanism by which DMH induces germ cell death. The lack of effect on serum testosterone levels in the current study, coupled with the lack of interstitial cell histopathology previously reported (Poon et al., 2004), argues that this effect is not due to impaired androgen secretion. A further argument against the notion that DMH impairs the endocrine control of spermatogenesis is the rapid increase in germ cells with TUNEL-detectable DNA fragmentation (i.e., 1 day PE) versus the more extended latency for a similar effect in studies of impaired follicle-stimulating hormone (FSH), luteinizing hormone (LH), or androgen secretion (e.g., Sinha Hikim et al., 1995; Shetty et al., 1996; Woolveridge et al., 1999). The pattern of germ cell death that rapidly follows DMH treatment is virtually identical to the pattern generated by treatment with a group of glycol ethers that are notable as testicular toxicants. Ethylene glycol monomethyl ether (EGME; Creasy et al., 1985) or its metabolite methoxyacetic acid (MAA: Tirado et al., 2004), cause marked increase in the frequency of DNA degradation, visualized by TUNEL labeling, in exactly the same germ cell populations as seen in the present study. Degeneration in late pachytene to meiotic spermatocytes in stage XI–XIV tubules and early pachytene spermatocytes in stage I–V tubules is visible morphologically within 12–24 h (Creasy et al., 1986; Moss et al., 1985) after a single oral dose of EGME. In addition, repeated exposure to either EGME or MAA results in atrophied testes and impaired spermatogenesis (Foster et al., 1983) and causes cortical atrophy of the thymus gland (House et al., 1985; Smialowicz et al., 1992; Williams et al., 1995), both of which have been noted in DMH-exposed animals (Poon et al., 2004). The similarity of response between EGME and DMH suggests that both substances are metabolized to a common, toxic metabolite such as MAA. In support of this contention, analysis of urine of animals in Experiment 1 (Table 7) showed that only the DMH-treated animals had a significant, 230-fold increase in MAA over the control, suggesting that metabolic conversion of DMH to MAA leads to the both the antispermatogenic and antithymic effects of this substance. In contrast, no significant elevation in MAA was observed in urine from BE- or EHxE-treated animals, whereas MHpE treatment led to a slight, albeit insignificant (p > 0.05), increase in MAA. It should be noted, however, that MHpE treatment resulted in a 16% decrease in HRSN (n.s.), a 20% reduction in cauda sperm reserves (n.s.), and a greater than fourfold increase in sperm heads with abnormal morphology; it also caused a significant decrease in the relative thymic weight (Poon et al., 2005), suggesting that this slight increase in MAA may have some impact on the testis and thymus gland. In addition, treatment with MOE, which bears a single methyl ether moiety (Fig. 5C), resulted in significantly elevated MAA levels in the urine, although at levels much lower than in DMH-treated animals (Poon and Chu unpublished), and in a slight but significant reduction in the proportion of round spermatids among testis cells. Taken together, it appears that the monomethoxy ethers do produce a very low level of MAA, and that level is reflected in small but observable toxic effects. It is notable that exposure to a methyl diether, 1,1-DMHp (Fig. 5E), with both methoxy moieties conjugated to a single apical carbon on the aliphatic chain, caused no evidence of testis toxicity, whereas DMH, with 2 methoxy moieties occurring at opposite ends of the carbon chain, is a potent testis toxicant. Although urine MAA levels have not been determined for 1,1-DMHp-treated animals, the dissimilarity in testis response from DMH-treated male rats argues for a lack of metabolism to MAA. It is noteworthy that urinary levels of MAA excreted by DMH-treated rats (roughly 3.6–6.0 mg over 24 h) are comparable to urinary MAA levels (0.95 mg over 18 h) observed in female rats exposed by inhalation to EGME at levels that produced embryo toxicity (Gargas et al., 2000) and in male rats exposed to EGME via drinking water to approximately 110 mg/kg (Medinsky et al., 1990). Although testicular effects were not examined in this latter study, daily oral administration of 100 mg/kg for 5 days caused pronounced atrophy of the seminiferous epithelium and a significant reduction in testicular LDH-X activity by 3 weeks after the first dose (Chapin et al., 1985). These observations support our contention that DMH toxicity is indeed mediated by its conversion to MAA.
In conclusion, our results indicate that the presence of two methoxy residues at opposite ends of an aliphatic chain is likely a critical feature that determines the dramatic toxicity of DMH to the seminiferous epithelium, as these structures predispose this molecule to be converted to MAA. Consequently, the use of ether molecules containing methoxy-residues, particularly with multiple methoxy residues at disparate points across an aliphatic chain, are inappropriately toxic for use as cetane-enhancing diesel additives. These insights into the critical molecular attributes that mark diethers with potential for testis toxicity will assist in the selection of environmentally benign oxygenated additives to improve diesel fuel combustion.
ACKNOWLEDGMENTS
The authors thank Dominic Patry, Gail Merrikin and the staff of the Animal Resources Division, and Lorraine Casavant of the Environmental & Occupational Toxicology Division, Health Canada, for their technical assistance. This work was supported by a grant from the Canadian Programme on Energy Research and Development (NRCan) to R.P. and I.C. and funding from the Environmental Health Science Bureau to M.G.W.
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ABSTRACT
Previous studies of the toxicity of candidate fuel additives identified severe testicular toxicity in animals exposed to 1,6-dimethoxyhexane (DMH). A series of studies were conducted to characterize the effects of DMH on spermatogenesis and to compare the effects of DMH with responses to structural similar aliphatic ethers. In the first study, sexually mature male rats were administered a single dose (600 mg/kg) of DMH, and subsets of animals were sampled at intervals post exposure (PE). Both testis and thymus weight declined steadily after DMH exposure, both being significantly lower than control by 7 days PE. Treatment with DMH led, at 24 to 48 h PE, to an increase in dying primary spermatocytes in seminiferous tubule stages I–IV and stages XII–XIV, but not intervening stages. The affected cohort of germ cells was seen progressing through the developmental sequence of spermatogenesis as numbers of dying cells returned to control levels by 7 days PE, coincident with a significant decline in the proportion of round spermatids among germ cells as determined by flow cytometry. Resolution of round spermatids to control levels by day 21 PE coincided with a reduction in condensed spermatids (homogenization-resistant spermatid nuclei) and was followed at 28 days PE by a significant reduction in cauda epididymal sperm counts. Further studies of repeated exposures to DMH (200 mg · kg–1 · day–1, 5 days per week for 4 weeks) confirmed the significant testis toxicity of this material. In contrast, similar treatment with any of a variety of structurally similar aliphatic ethers had little or no impact on testis function. Methoxyacetic acid, previously shown to cause rapid death of some meiotic germ cell stages, was found at high concentrations in the urine of DMH-treated rats but not in rats treated with other ethers, suggesting that DMH exerts its testis toxicity via metabolism to this substance. These results demonstrate that DMH selectively deletes germ cells from the testis At the very early or very late pachytene, diplotene, or M-phase spermatocyte stages, possibly through conversion to MAA.
Key Words: fuel additives; testis toxicity; spermatogenesis; methoxyacetic acid; pachytene spermatocytes.
INTRODUCTION
Diesel fuels are important energy sources in transport and industry. The combustion of diesel, however, produces massive quantities of air pollutants, including carbon monoxide, nitrogen oxides, volatile organics, and exhaust particles and particulates. There is growing realization that these pollutants contribute to death and morbidity due to cardiovascular and respiratory disease, among other health impacts, and that they are therefore very significant health concerns (Burnett et al., 1998; Goldberg et al., 2000; Villeneuve et al., 2002). The addition of oxygenated substances, such as liquid ethers, into diesel has been shown to improve combustion efficiency. Furthermore, the presence of oxygenates has been reported to reduce the emission of exhaust particulates, carbon monoxide, and total hydrocarbons (Zhu et al., 2003; Shen et al., 1993; Heinz, 1993).
Our laboratory has undertaken toxicological studies of various aliphatic ethers in order to identify compounds with low toxicity for use as diesel additives (Poon, et al., 2000; 2004; in press). These studies revealed that repeated oral exposure to 1,6-dimethoxyhexane (DMH) causes significant reduction in testis weight to about 70% of control, among other significant toxic effects (Poon et al., 2004). Histological examination of the testes and epididymis of DMH-exposed animals found many seminiferous tubules lacking normal spermatids, increased numbers of spermatid giant cells, and apparently a reduced density of mature sperm. In contrast, eight other aliphatic ethers examined in these studies had no significant effect on testis. It was postulated that the testicular effects of DMH were caused by its metabolism to methoxyacetic acid (MAA) (Poon, 2004). This hypothesis was supported by a subsequent study wherein MAA was detected in the blood and urine of DMH-treated animals (Poon et al., 2005). In the present study, we extend initial observations of a testicular effect in DMH by examining multiple testicular end points in animals exposed to DMH or to any of eight other structurally similar ethers. These end points include the relative proportion of round spermatids, number of homogenization-resistant sperm nuclei, cauda epididymal sperm morphology, and testis lactate dehydrogenase X (LDH-X) activity. To further understand the mechanisms and time course of action of DMH, acute dosing studies were performed which examined the progression of DMH-induced testis lesions and recovery.
MATERIALS AND METHODS
Chemicals and reagents.
1,6-Dimethoxyhexane was synthesized by Wychem (Suffolk, UK) and had a purity of >99% as measured by gas chromatography. Syntheses of 1,1-dimethoxyoctane (1,1-DMO), 1-ethoxyheptane (EHpE), 1- ethoxyhexane (EHxE), 1-isopropoxyoctane (IPO), 1-methoxyheptane (MHpE), 1-methoxyoctane (MOE), and 1-proxyhexane (PHxE) were carried out by M. S. Kaliaguine (Universite Laval, Quebec, PQ) and all had a purity >97% as measured by gas chromatography. Dibutyl ether (BE) was purchased from Aldrich Chemical (Milwaukee, WI) and had a purity of >99%. Nonyl-acridine orange (NAO) and propidium iodide (PI) were purchased from Molecular Probes (Eugene, OR). In situ apoptosis detection kit was purchased from R&D Systems (Minneapolis, MN). Periodic acid–Schiff histological staining reagents were purchased from Richard Allan Scientific (Kalamazoo, MI). All other chemicals are of reagent grade and were obtained from either Fisher Scientific (Nepean ON, Canada) or Sigma Chemical Co. (St Louis, MO).
Animals and experimental approach.
Sexually mature Sprague-Dawley rats, (8–9 weeks of age for subacute studies (see below) or 11–12 weeks for acute study) were purchased from Charles River Canada (St. Constant, PQ) and were routinely housed in pairs in polycarbonate hanging cages, open to room air through a micro-barrier filter, with free access to water and laboratory rat chow (Purina 5008; Agribrand Purina; Saint-Hubert, Quebec) under controlled photoperiod (12:12 light:dark), humidity (40–70%), and temperature (20–24°C). All animal handling procedures adhered to Canadian Council on Animal Care guidelines and were approved by the Health Canada Animal Care Committee prior to the initiation of the study.
The results of four separate animal studies are described in this article. The first study examined the progression of lesion and recovery of the testis and the reproductive tract after a single oral dose of DMH (acute study). The remaining studies were designed to examine the effects of repeated doses of any of a variety of aliphatic ethers, along with a group receiving an effective dose of DMH, for 4 weeks, on an array of end points indicating reproductive toxicity (subacute studies). The full results of these latter studies, beyond the effects on the reproductive tract described herein, will be reported elsewhere.
For the acute study, animals were administered a single dose of DMH (600 mg/kg in 5 ml/kg corn oil) by gavage, and groups of 7–8 animals were sacrificed 4 h, 1, 2, 3, 7, 14, 21, or 28 days post exposure (PE). An additional control group of 8 animals received 5 ml/kg corn oil and were sacrificed 24 h PE.
Three separate studies examined the general and reproductive consequences of 4 weeks of exposure (200 mg · kg–1 · day–1, 5 days per week for 4 weeks) to BE, EHxE, and MHpE (Experiment 1); EHpE, MOE, and PHxE (Experiment 2); and IPO and 1,1-DMO (Experiment 3). In each of these studies, a separate group of rats treated with DMH were used as the comparative group. The details of animal treatment and urine collection are described elsewhere (Poon et al., 2005). Briefly, animals were randomized into treatment groups of 7 animals each during a 1-week acclimation phase. Animals received, by oral gavage, 200 mg/kg of one of the 9 alkyl ethers in corn oil, 5 days per week for 4 weeks. Control animals received corn oil only. Animals were sacrificed beginning 24 h after the final exposure.
Tissue collection.
Animals were sacrificed by exsanguination under isofluorane anesthesia. Routinely, the testes and epididymis were rapidly removed, dissected clear of fat, and weighed. The tunica albuginia of the right testis was perforated 20 times with a 20-gauge needle and fixed by immersion in modified Davidson's solution (Latendresse et al., 2002) for subsequent analysis of stage-specific germ cell apoptosis. The left testis was decapsulated, and the parenchyma was divided into four roughly equivalent portions by sagittal then transverse cuts through the midline of the testis. One portion was placed in ice-cold medium 199 (M199 containing 0.1% BSA) for analysis of spermatogenic cell populations. Two portions were weighed, separately frozen on dry ice, and stored at –20°C prior for analysis of homogenization-resistant sperm nuclei and testis-specific lactate dehydrogenase (LDH) isoform, LDH-X. Several small incisions were made in the left cauda epididymis, and it was placed into 10 ml of warmed Dulbecco's phosphate buffered saline (PBS) (Gibco BRL, Bethesda, MD) to allow sperm to swim out. One 200-μl sample of sperm was stained with eosin and a portion smeared on a microscope slide, dried, and cover-slipped to examine sperm morphology. The remaining media containing sperm and the cauda were stored at –20°C for analysis of cauda sperm reserves.
Testis histology and analyses of apoptotic germ cells.
Germ cell death was examined in five animals each, from control, 4 h, 1, 2, 3, and 7 days post exposure groups of the first study only. Testes were fixed in modified Davidson's solution for 72 h and washed twice in PBS (48 h total), after which a central transverse section was embedded in paraffin and sectioned at 5 μm. One section was stained with periodic acid–Schiff to highlight acrosome structure to assist in seminiferous tubule staging (Hess, 1990), and the adjacent section was processed for in situ DNA end-labeling (TUNEL labeling) to identify apoptotic cells with the TdT In Situ Apoptosis Detection Kit–DAB (daminobenzidine; R & D Systems, Minneapolis, MN). Stages of seminiferous tubule cross sections were determined by previously described criteria (Hess, 1990).
Testis cell isolation and analysis.
Fresh testes were dispersed using a two-stage enzymatic digestion as previously described (Bellve et al., 1977). Testes were incubated in 2.5 ml M199 1% BSA, containing 56 U/ml collagenase type III (Worthington Biochemical, Freehold, NJ) and 8 U/ml DNAase I (Sigma, St Louis, MO) at 32°C in a shaking water bath for 15 min. Tubes were then transferred to ice and, after settling for 1 min, the supernatant, containing interstitial cells, was aspirated and discarded. The remaining seminiferous tubules were washed twice with 10 ml of M199 0.1% BSA and then incubated as above in 5 ml of M199 containing 110 U/ml trypsin (Worthington) and 10 U/ml DNAase I. Tubes were then placed on ice and gently aspirated repeatedly with a Pasteur pipette to disrupt cell aggregates, after which 10% BSA stock was added (final concentration of 0.5% BSA) and mixed by aspiration through the Pasteur pipette. The cell suspension was then filtered through a 70-μm nylon mesh (Filicon, Dako Canada, Mississauga, ON), centrifuged (400 x g for 8 min at 4°C), washed twice with 10 ml ice-cold DPBS containing 0.5% BSA (PBS-BSA; pH 7.4) and fixed in ice-cold 70% ethanol. Fixed cells were kept at –20°C for no more than 30 days prior to analysis of spermatogenic cell populations.
Relative proportions of spermatogenic cell populations were assessed in fixed cells using a flow cytometric method (Petit et al., 1995; Suter et al., 1997). The principle of the test is that spermatogenic cells, as they differentiate from normal diploid spermatogonial stem cells through to mature spermatozoa with a highly condensed haploid complement of DNA, progress through various intermediate stages with differing nuclear DNA content and cellular content of mitochondria. With the use of fluorescent probes to quantitatively label DNA (PI) and mitochondria (NAO), the relative numbers of cells in each intermediate population can be resolved to determine relative numbers at each germ cell stage. Cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences, Mississauga, ON) fitted with an argon ion laser (488 nm line excitation); fluorescence emission of NAO was reflected by a 550 dichroic longpass filter and quantified after passage through a 530/30 nm bandpass filter. Fluorescence of PI was detected after sequential passage through the 550 dichroic filter and a 670-nm longpass filter. Relative proportions of cells in each population were calculated with WinList software (Verity Software, Topsham, ME). The germ cell types in each population were identified under a fluorescent microscope (Zeiss Axioskop 2, Carl Zeiss, Thornwood, NY) by examination of cells from each population, which were isolated using the cell-sorting capacity of the FACSCalibur flow cytometer.
Cauda epididymal sperm morphology.
Assessment of sperm morphology followed the guidance of an expert consensus statement (Seed et al., 1996). Briefly, eosin-stained sperm collected from the cauda epididymis were smeared onto two glass slides per sample, air-dried, and cover-slipped. At least 200 sperm on each slide were examined for the proportion of sperm with abnormal head (overhooked, blunt hook, banana-shaped, amorphous, or extremely oversized) or tail (twisted, bent, corkscrew, double/multiple) by one individual unaware of animal number or treatment. Values for individual slides were averaged for each animal.
Homogenization-resistant spermatid nuclei (HRSN) and cauda epididymis sperm reserves.
The total number of condensed spermatids per testis and the total number of sperm in the cauda epididymis were estimated according to a previously described method (Blazak et al., 1985). The principle of this method relies on the extreme stability of the nuclei from stage 17–19 spermatids or cauda sperm, which allows counting of these after complete homogenization and sonication of the tissue. To determine testis condensed spermatid number, a weighed portion of the parenchyma from the left testis, as representative of the whole organ as possible, was homogenized in 20 ml of STA solution (0.9 % NaCl, 0.01% Triton X-100, and 0.025% sodium azide) in a 50-ml Waring Ultramicro blender (Fisher Scientific, Nepean, Ont.) on low setting for 2 min. Similarly, whole cauda and associated sperm suspension in DPBS were thawed on ice and, after addition of 200 μl of 1% Triton X-100 (final concentration of 0.01%) and brought up to 20 ml volume as STA, were homogenized as above. Homogenates of either tissue were then disrupted with a VibraCell sonicator (Sonics & Materials, Danbury, CT) using a microprobe on setting 7 for 60 s. Sperm or HRSN densities were calculated from the average number of nuclei in four fields of a Neubaur hemocytometer and were expressed as total or as per gram of epididymis or testis weight.
Analysis LDH and LDH-X.
Testis fragments were thawed slightly and then homogenized in 2 ml/g of ice-cold 12.5 mM TRIS KCl (pH 9.0) with 5 strokes of a manual homogenizer (Kontes Glass Company, Vineland, NJ). The homogenate was centrifuged at 19,000 x g for 10 min at 4°C and the supernatant was diluted fivefold in TRIS KCl. Separate aliquots were used to determine total LDH and LDH-X. Enzyme activity was measured by monitoring extinction of NAD absorbance at 340 nm in a reaction mixture that contained 4.2 mM of NAD in 10.5 mM Tris KCl (pH 9.0) at 30°C. After background NAD conversion was monitored for 30 s, the appropriate substrate (lactic acid for total LDH, alpha-hydroxycaproic acid for LDH-X) was added to a final concentration of 50 mM and, after brief mixing (2 s), 340 nm extinction was monitored for a further 30 s. Enzyme activity was calculated from the linear region of NAD conversion, corrected for background rate and converted to μmoles converted · (min)–1 · (mg homogenate protein)–1. Total protein in testis homogenates was determined with the modified micro-Lowry protein determination kit (Sigma, St Louis, MO).
Analysis of 2-methoxyacetic acid.
After receiving their final gavage dose, animals were transferred to metabolic cages and provided with food and water ad libitum. The entire volume of urine produced over 24 h was collected in a 50-ml conical tube and held on wet ice until the end of the collection period. The sample was then stored at –20°C until analysis. The method for analysis of the 2-methoxyacetic acid (MAA) was set up in this laboratory and will be reported elsewhere in detail (Li et al., manuscript in preparation). Solid-phase extraction of urine was carried out with a Supelco 12-port Visiprep SPE Vacuum Manifold. Before sample application, 500 mg Bakerbond C18 cartridges were conditioned with 3 ml of methanol and 5 ml of water applied sequentially, after which 1 ml of urine sample was acidified with 100 μl of concentrated HCl and applied to the cartridges with a Pasteur pipette and allowed to flow through by gravity. After sample application, the cartridges were centrifuged at 3000 rpm for 5 min, placed on the SPE vacuum manifolds, and air-dried for 15 min with an air stream generated from a vacuum pump. Each of the C18 cartridges was then connected to a Bond ElutT Na2SO4 cartridge (1.4 g; Varian, Palo Alto, CA). The serially connected cartridges were eluted with 2 ml of 2.5% isopropanol in diethyl ether. The residue water was dried while the eluant was flowing through the Na2SO4 cartridge.
An internal standard of 2-bromopropoinic acid (BrPA; 99%, Supelco, Bellefonte, PA), prepared in methyl-tert-butyl ether (MTBE) at 0.25 mg/ml was added (100 μl) to the eluant, and the solution was concentrated at ambient temperature to a final volume of 50 μl with a gentle stream of nitrogen. The concentrates were reconstituted with hexane to a volume of 200 μl. Derivatization with diazomethane was conducted using a modification of USEPA method 515.1 Briefly, the diazomethane generated from diazald and 37% KOH was bubbled through 1/16'' polytetrafluoroethylene (PTFE) tubing into the hexane solution with the aid of a gentle stream of nitrogen at a rate of 10 ml/min until the hexane solution turned yellowish. Despite the very small volumes of diazomethane used, great care was taken to avoid human exposure to this toxic substance. All the work was conducted in a well-ventilated fume hood, and a long tweezers was used to manipulate the outlet of PTFE tubing into the sample solution. The outlet of the PTFE tubing was rinsed with hexane between samples to avoid cross contamination. After 30 min, the diazomethane in sample was destroyed by addition of silicic acid. The GC/MS analysis was performed with a Thermo Finnigan Trace 2000 gas chromatograph coupled to a PolarisQ mass spectrometer. The instrument was equipped with an AS 2000 liquid autosampler, a Supelco OMEGAWAX 250 fused silica capillary column (30 m x 0.25 mm x 0.25 μm), and the Xcalibur data system. The oven temperature was programmed at the initial temperature of 50°C, held for 2 min, and then increased at a rate of 2°C/min to 120°C. The oven was held at this temperature for 1 min, then increased at 30°C/min to the temperature of 220°C, which was held for 3 min. Helium was the carrier gas; it was used with a flow rate of 0.8 ml/min and the split flow was 32 ml/min. A 2-μl volume of sample solution was injected with a splitless injection mode and 1.0-min purge time. The temperatures of the injection port, transfer line, and MS ion source were set at 220°C, 250°C, and 200°C, respectively. The selected ion monitor (SIM) was used for quantification. The method is validated and can be employed to detect the MAA for a concentration as low as 2 ng/ml (ppb).
Statistical analyses.
After successful testing for assumptions of normal distribution (Shapiro-Wilks Goodness of Fit Test) and homoscedacity (Bartlett's test) all data were analyzed by analysis of variance (ANOVA). Proportional data (testis cell data, percent abnormal sperm cells) were arcsine transformed prior to analysis to normalize. Where normality or equal variance were not indicated, data were retested after log transformation and, if normality or equal variance assumptions were not satisfied, non-transformed data were analyzed for ether treatment effects using ANOVA on ranks (Wilcoxon-Kruskal-Wallis test). Data for which ANOVA identified a significant effect of ether treatment (p < 0.05) were subjected to Dunnett's or Dunn's post hoc analysis, for parametric or non-parametric ANOVA results, respectively, to identify effective treatments. All statistical analyses were conducted with JMP software, release 4.0.2 (SAS Institute, Cary, NC) except for the nonparametric ANOVA and Dunn's tests, which were performed using SigmaStat, release 2.03 (SPSS Inc., Chicago, IL).
RESULTS
DMH Effects on Tissue Weights and Germ Cell Death
To examine the progression of DMH-induced testis toxicity, male rats were administered a single dose of DMH (600 mg/kg) and were sacrificed at various times post-exposure. Seven days after DMH exposure, relative testis and thymus gland weights were significantly reduced and continued to decline up until the final sampling 28 days PE. In contrast, epididymis weights were not significantly influenced by DMH exposure (Table 1).
The reduction in testis weight 7 days PE, as well as previous observations of DMH-induced atrophy of the seminiferous epithelium (Poon et al., 2004), suggested that DMH exposure caused disruption of spermatogenesis. To evaluate the impact of DMH on germ cell survival, testis sections were stained for DNA fragmentation indicative of cell death using the TUNEL reaction (terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling: Gavrieli et al., 1992). Seminiferous tubules of control animals had very few TUNEL-positive cells, indicating very low level germ cell attrition in normal testes. The number of TUNEL-positive cells was similar to control levels at 4 h PE, but it increased dramatically at 1 and 2 days PE (Fig. 1, Fig. 2A). Positive cells were still significantly elevated at 3 days PE, to a lesser extent than at 1 and 2 days PE, but they returned to control levels by 7 days PE. Stage-specific analyses of TUNEL-positive cells in seminiferous tubules at 1 and 2 days PE demonstrated that DMH-induced cell death is highly specific to pachytene spermatocytes in stages I–IV and to late pachytene, diplotene, and meiotic primary spermatocytes in stages XII–XIV, whereas pachytene spermatocytes intermediate to these stages are unaffected (Fig. 1, Fig. 2B). The early pachytene spermatocytes (in stage I–IV tubules) seemed to be initially more susceptible to DMH-toxicity, as there was a significantly greater number of TUNEL-positive cells in this group at 24 h PE versus the same group at 48 h PE or stage XI–XIV stage tubules at 24 h PE (Fig. 2B). At 48 h PE, there were more TUNEL-positive cells in the stage XI–XIV group than in stage I–IV tubules at the same time PE or than were seen in stage XI–XIV tubules 24 h PE.
DMH Effects on Spermatogenesis
To further evaluate the effects of DMH on spermatogenesis, the relative proportion of round spermatids was evaluated using a flow cytometric method that identified populations of testicular cells based on PI (indicating DNA) and NAO (mitochondrial) content. Representative scattergrams of testis cells isolated from a control animal (Fig. 3A) and 14 day PE animal (Fig. 3B) are provided, demonstrating the pattern characteristic of these analyses. Although six separate populations, identified by fluorescent microscopic analyses of sorted cells, could be evaluated using this method, only data for round spermatids are presented. The data for other populations tended to be highly variable, with coefficients of variability in excess of 40% (data not shown). As expected, round spermatids constituted the most prominent population of germ cells, representing 53.9 ± 3.5% of all cells in preparations from testes of control animals (Fig. 4). Exposure to DMH caused a latent reduction in the relative proportion of round spermatids that was significant by 7 days PE, and this population reached a minimum level 14 days PE (37.8 ± 8.7%) but returned to levels similar to control thereafter.
The progression of the affected cohort of germ cells through the successive stages of spermatogenesis was also demonstrated by the number of condensed spermatids (homogenization-resistant sperm nuclei [HRSN]; Table 2). The DMH-induced reduction in HRSN appeared 1 week after the DMH-induced reduction of round spermatid levels. Mean HRSN, expressed either as total for the left testis or per gram of testis parenchyma, were significantly reduced by 21 days PE or remained at this lower level at 28 days PE (Table 2). Cauda epididymal sperm reserves per gram of cauda weight were also significantly reduced 28 days PE, but not at earlier times, further indicating the progression of the effected cohort of germ cells through the reproductive tract.
Effects of Repeated Doses of DMH versus Other Ethers
A further series of experiments was conducted to compare the toxicity of DMH with that of several structurally similar types of ether. The effects of these ethers on spermatogenesis and cauda sperm are presented below. The effects of oral exposure to various aliphatic ethers for 5 days per week for 4 weeks on body weight, weights and histopathological analyses of major organs, measures of serum biochemistry, hematology, and some liver enzymes from these studies will be described elsewhere (Poon et al., 2005; Poon and Chu, unpublished). In all of these studies, only DMH caused a significant reduction in relative testis weight (i.e., g testis weight/100 g body weight) with a 21.4%, 16.7%, and 29.6% reduction from relative control testis weight in the first, second, and third studies, respectively (data not shown).
To determine the impact on germ cell survival, the effect of ether treatment on the proportion of round spermatids among testis cells was examined for all animals (see Table 4). In all three of the 4-week studies, DMH treatment led to a significant reduction in the proportion of round spermatids to 63%, 64%, and 58% of control levels in Exeriments 1, 2, and 3, respectively (Table 3). Of all other ethers examined, only MOE had any effect on round spermatid number, causing a significant reduction of round spermatids to 83.8% of control levels (p < 0.05).
Testis sperm production, as indicated by HRSN, was significantly reduced in animals exposed to DMH, yielding HRSN numbers that were 49%, 45%, and 71% of total left testis HRSN of control animals in the three experiments (Table 4). Like testis sperm production, epididymal sperm reserves were reduced by DMH treatment, causing a 37%, 43%, and 31% reduction in absolute numbers per left cauda epididymis for the first, second, and third studies, respectively, although the effect in the third experiment did not reach statistical significance (Table 4). In contrast to DMH, most of the other ethers were without effect on either testis HRSN or cauda sperm counts. The exceptions to this was BE, which caused a slight but significant reduction in HRSN (p < 0.05).
Sperm morphology was examined as a measure of spermatid final maturation (Table 5). Treatment with DMH led to a significant reduction in the proportion of morphologically normal sperm (p < 0.05) in all three studies, although the degree of effect differed between studies ranging from roughly a 50% decline in normal in the first study (42.5 ± 17.6% normal sperm in DMH versus 94.5 ± 2.3% in control) to roughly 13% for the other two studies. In all three studies there was a marked increase in the proportion of morphologically abnormal sperm heads, indicating a possible effect of DMH on spermatid nuclear condensation. In contrast with the effects of DMH, sperm morphology was unaffected by treatment with any other ether tested in these studies (Table 5: p > 0.05).
To further evaluate testicular toxicity, the activity of a germ-cell–specific LDH isoform, LDH-X was evaluated. Activity of total LDH and LDH-X specific activity were measured in testis homogenate using lactic acid and -hydroxycaproic acid, as the substrate. Although total testicular LDH activity was not influenced by any ether treatment, the activity of LDH-X was reduced by DMH treatment to 77.0%, 75.3%, and 85.9% of control values for the first, second and third studies, respectively, although the effect in the third experiment did not reach statistical significance (Table 6). These results are consistent with the observed reduction in round spermatids where this isoform is expressed (Hintz and Goldberg, 1977; Meistrich et al., 1977). Activity of LDH-X was not significantly influenced by any other ether treatment.
Urinary MAA Levels in DMH-Exposed Animals
As the pattern of germ cell death observed in DMH-treated animals 1 and 2 days PE was remarkably similar to that observed in animals exposed to 2-methoxyacetic acid (Foster et al., 1987), we hypothesized that DMH-induced germ cell toxicity may be consequent to metabolism of this compound. To test this hypothesis, we compared MAA residue levels in urine from DMH-treated animals from experiment 1 of the subacute studies with those in the urine collected from control animals or animals exposed to ethers that caused no testis toxicity. Methoxyacetic acid was slightly above the limit of detection in all samples, including those from vehicle control animals, suggesting some transfer of the volatile parent compound from dosed animals. However, urine collected from DMH-treated animals contained MAA levels that were, on average, 230-fold greater than that found in control animals (Table 7). Levels of MAA detected in urine from MHpE-treated animals appeared to be higher than controls, but they did not reached a statistically significant level.
DISCUSSION
The present study demonstrates that subacute oral exposure to the aliphatic diether DMH causes significant disruption of spermatogenesis and impaired maturation of sperm. This confirms and extends the findings of a previous study showing that subacute oral exposure to DMH caused a dose-related decrease in testis weight, degeneration of seminiferous epithelium, among other histopathological lesions in the testis, and increased presence of abnormal sperm in the epididymal tubules (Poon et al., 2004). Here we have demonstrated that DMH exposure results in death of pachytene spermatocytes at very precise stages of differentiation, with consequent reduction in relative numbers of round spermatids, numbers of HRSN, and epididymal sperm reserves. In conjunction with the observed reduction in sperm production, repeated DMH exposure caused an increased proportion of sperm with abnormal morphology, demonstrating a toxicant-induced impairment in the production of normal spermatozoa.
Our data demonstrate that a single oral exposure to DMH causes a sharp increase in germ cell death at 24 and 48 h PE, with a subsequent reduction in germ cell death to control levels by 7 days PE. We also demonstrate that DMH toxicity is highly specific to early pachytene spermatocytes in tubule stages I–IV and to late pachytene spermatocytes, diplotene spermatocytes, or meiotically dividing spermatocytes in tubule stages X–XIV. Other than these cells, there was no increase in TUNEL-positive cells in the intervening stages, nor was there any evidence of dying spermatids or spermatogonia in any tubule at any stage. The reduction in germ cell death to control levels by 7 days PE, suggesting that DMH-induced effects on spermatogenesis are transient, is supported by the transient reduction in the number of round spermatids observed on PE days 7 and 14 but not on PE days 21 or 28. The effects seen in the single-dose study are largely confirmed by the results of the repeated dose studies, with one exception: The significant (p < 0.05) increase in sperm with abnormal head morphology was consistently observed in DMH-treated animals in the subacute studies but was very minimal in samples from the single-dose study, suggesting that repeated exposure to DMH has a greater impact on events in spermiogenesis, spermiation, or cauda epididymal function than occurs in response to a single exposure.
In addition to reducing sperm production and cauda epididymal sperm reserves, exposure to multiple doses of DMH for 4 weeks also caused an increase in the proportion of sperm with abnormal morphology. As abnormal morphology of sperm head has been shown to be a good indicator of reduced fertility in both animal studies (e.g., Ballachey et al., 1987) and human studies (e.g., Bonde et al., 1998; Kruger et al., 1988), such observations suggest that DMH causes a reduction in both epididymal sperm numbers and quality.
The above results, together with earlier observations, provide a close glimpse into the mechanism by which DMH induces germ cell death. The lack of effect on serum testosterone levels in the current study, coupled with the lack of interstitial cell histopathology previously reported (Poon et al., 2004), argues that this effect is not due to impaired androgen secretion. A further argument against the notion that DMH impairs the endocrine control of spermatogenesis is the rapid increase in germ cells with TUNEL-detectable DNA fragmentation (i.e., 1 day PE) versus the more extended latency for a similar effect in studies of impaired follicle-stimulating hormone (FSH), luteinizing hormone (LH), or androgen secretion (e.g., Sinha Hikim et al., 1995; Shetty et al., 1996; Woolveridge et al., 1999). The pattern of germ cell death that rapidly follows DMH treatment is virtually identical to the pattern generated by treatment with a group of glycol ethers that are notable as testicular toxicants. Ethylene glycol monomethyl ether (EGME; Creasy et al., 1985) or its metabolite methoxyacetic acid (MAA: Tirado et al., 2004), cause marked increase in the frequency of DNA degradation, visualized by TUNEL labeling, in exactly the same germ cell populations as seen in the present study. Degeneration in late pachytene to meiotic spermatocytes in stage XI–XIV tubules and early pachytene spermatocytes in stage I–V tubules is visible morphologically within 12–24 h (Creasy et al., 1986; Moss et al., 1985) after a single oral dose of EGME. In addition, repeated exposure to either EGME or MAA results in atrophied testes and impaired spermatogenesis (Foster et al., 1983) and causes cortical atrophy of the thymus gland (House et al., 1985; Smialowicz et al., 1992; Williams et al., 1995), both of which have been noted in DMH-exposed animals (Poon et al., 2004). The similarity of response between EGME and DMH suggests that both substances are metabolized to a common, toxic metabolite such as MAA. In support of this contention, analysis of urine of animals in Experiment 1 (Table 7) showed that only the DMH-treated animals had a significant, 230-fold increase in MAA over the control, suggesting that metabolic conversion of DMH to MAA leads to the both the antispermatogenic and antithymic effects of this substance. In contrast, no significant elevation in MAA was observed in urine from BE- or EHxE-treated animals, whereas MHpE treatment led to a slight, albeit insignificant (p > 0.05), increase in MAA. It should be noted, however, that MHpE treatment resulted in a 16% decrease in HRSN (n.s.), a 20% reduction in cauda sperm reserves (n.s.), and a greater than fourfold increase in sperm heads with abnormal morphology; it also caused a significant decrease in the relative thymic weight (Poon et al., 2005), suggesting that this slight increase in MAA may have some impact on the testis and thymus gland. In addition, treatment with MOE, which bears a single methyl ether moiety (Fig. 5C), resulted in significantly elevated MAA levels in the urine, although at levels much lower than in DMH-treated animals (Poon and Chu unpublished), and in a slight but significant reduction in the proportion of round spermatids among testis cells. Taken together, it appears that the monomethoxy ethers do produce a very low level of MAA, and that level is reflected in small but observable toxic effects. It is notable that exposure to a methyl diether, 1,1-DMHp (Fig. 5E), with both methoxy moieties conjugated to a single apical carbon on the aliphatic chain, caused no evidence of testis toxicity, whereas DMH, with 2 methoxy moieties occurring at opposite ends of the carbon chain, is a potent testis toxicant. Although urine MAA levels have not been determined for 1,1-DMHp-treated animals, the dissimilarity in testis response from DMH-treated male rats argues for a lack of metabolism to MAA. It is noteworthy that urinary levels of MAA excreted by DMH-treated rats (roughly 3.6–6.0 mg over 24 h) are comparable to urinary MAA levels (0.95 mg over 18 h) observed in female rats exposed by inhalation to EGME at levels that produced embryo toxicity (Gargas et al., 2000) and in male rats exposed to EGME via drinking water to approximately 110 mg/kg (Medinsky et al., 1990). Although testicular effects were not examined in this latter study, daily oral administration of 100 mg/kg for 5 days caused pronounced atrophy of the seminiferous epithelium and a significant reduction in testicular LDH-X activity by 3 weeks after the first dose (Chapin et al., 1985). These observations support our contention that DMH toxicity is indeed mediated by its conversion to MAA.
In conclusion, our results indicate that the presence of two methoxy residues at opposite ends of an aliphatic chain is likely a critical feature that determines the dramatic toxicity of DMH to the seminiferous epithelium, as these structures predispose this molecule to be converted to MAA. Consequently, the use of ether molecules containing methoxy-residues, particularly with multiple methoxy residues at disparate points across an aliphatic chain, are inappropriately toxic for use as cetane-enhancing diesel additives. These insights into the critical molecular attributes that mark diethers with potential for testis toxicity will assist in the selection of environmentally benign oxygenated additives to improve diesel fuel combustion.
ACKNOWLEDGMENTS
The authors thank Dominic Patry, Gail Merrikin and the staff of the Animal Resources Division, and Lorraine Casavant of the Environmental & Occupational Toxicology Division, Health Canada, for their technical assistance. This work was supported by a grant from the Canadian Programme on Energy Research and Development (NRCan) to R.P. and I.C. and funding from the Environmental Health Science Bureau to M.G.W.
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