Metabolic Activation of 2,6-Xylidine in the Nasal Olfactory Mucosa and the Mucosa of the Upper Alimentary and Respiratory Tracts in Rats

Eva Tydén, Hans Tjälve and Pia Larsson1

Department of Biomedical Sciences and Veterinary Public Health, Division of Pathology, Pharmacology and Toxicology, Swedish University of Agricultural Sciences, 750 07 Uppsala, Sweden

Received April 28, 2004; accepted July 7, 2004


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Whole-body low-temperature radioluminography of 3H-2,6-xylidine in rats indicates that the nonmetabolized substance, which is a volatile and fat-soluble compound, is distributed throughout the body and accumulates in adipose tissues, e.g., in the abdominal and subcutaneous regions. Whole-body autoradiography with freeze-dried or solvent-extracted tissue sections as well as microautoradiography, which were used to trace tissues in the rats accumulating 2,6-xylidine metabolites, showed presence of tissue-bound 2,6-xylidine metabolites in the nasal olfactory mucosa and the mucosa of the upper alimentary and respiratory tracts. These tissues were found to have an in vitro capacity to bioactivate 2,6-xylidine. Our data indicate that 2,6-xylidine in vivo undergoes an in situ bioactivation in these extrahepatic tissues. Our results showed that the nasal olfactory mucosa had a much higher capacity than the other examined tissues to bioactivate 2,6-xylidine. Thus, the carcinogenic effect of 2,6-xylidine toward the nasal mucosa in rats is correlated with a high capacity of this tissue to bioactivate the compound.

Key Words: 2,6-xylidine; metabolic activation; nasal olfactory mucosa; alimentary tract; respiratory tract; nasal cancer.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The aromatic amine 2,6-xylidine (2,6-dimethylaniline) is used as a chemical intermediate in the production of dyes, pesticides, and pharmaceuticals (IARC, 1993Go). It has also been detected in tobacco and tobacco smoke (Irvine and Saxby, 1969Go; Patrinakos and Hoffman, 1979Go).

Several local anesthetics, such as lidocaine (xylocaine), etidocaine, mepivacaine, ropivacaine, and bupivacaine are made with 2,6-xylidine. The substance has been identified as a metabolite in the urine of rats, guinea-pigs, dogs, and humans given lidocaine orally (Keenaghan and Boyes, 1972Go). 2,6-Xylidine has also been shown in samples of milk from bovines given lidocaine during surgery and in breast milk samples of a human donor who received lidocaine during dental work (Puente and Josephy, 2001Go). In addition, the substance has been identified as a major lidocaine metabolite in human liver slices in vitro (Parker et al., 1996Go). 2,6-Xylidine has also been shown to be a metabolite excreted in the urine of humans exposed to etidocaine (Thomas et al., 1976Go).

The alpha-2-adrenergic agonist xylazine, which is used in veterinary medicine as a sedative, analgesic, and muscle relaxant, consists of 2,6-xylidine coupled to a thiazine ring. In cows given a therapeutic im dose of xylazine a metabolite identified as 2,6-xylidine has been detected in urine up to 10 h after administration (Pütter and Sagner, 1973Go). 2,6-Xylidine is an in vivo and in vitro metabolite in rats and humans of the experimental anticonvulsant agent N-(2,6-dimethylphenyl)-5-methyl-3-isoxazolecarboxamide (Martin et al., 1997Go). The substance is also the major degradation product of the hydrolytic reaction of lidamidine, which is an antidiarrheal agent (Zalipsky et al., 1978Go). In addition, 2,6-xylidine has been identified as an impurity of the fungicide metalaxyl (Dureja et al., 2000Go).

2,6-Xylidine is a rat nasal carcinogen. Thus, a high incidence of nasal tumors has been observed in rats fed a diet containing 3000 ppm of 2,6-xylidine for 2 years (Kornreich and Montgomery, 1990Go). There was also an increased incidence of subcutaneous tissue fibromas in the 2,6-xylidine–treated rats. In rats fed a diet containing 3000 ppm of 2,6-xylidine for 4 weeks atrophy of Bowman's glands was observed in the dorsal meatus of the olfactory region of the nasal cavity (Yasuhara et al., 2000Go).

Aromatic amine carcinogens are known to exert their biological effects following cytochrome P-450 (CYP)-mediated metabolic activation to reactive electrophilic intermediates that bind to DNA (Marques et al., 1997Go). In the case of 2,6-xylidine, the nitrogen of the amino group and the para position on the aromatic ring are the two possible sites for the initial CYP-mediated monooxygenation, leading to the production of N-(2,6-dimethylphenyl)hydroxylamine (DMHA) and 4-amino-3,5-dimethylphenol (DMAP), respectively (Fig. 1). DMHA has the potential to react with DNA either directly or after esterification (Gan et al., 2001Go; Marques et al., 2002Go). DMAP is potentially genotoxic by a mechanism involving nonenzymatic oxidation to an iminoquinone, which is a strong electrophile (Gan et al., 2001Go). Indirect evidence for the formation of N-hydroxylated aromatic amines in vivo can be obtained from the detection of hemoglobin adducts, which are formed by covalent binding of the bioactivated amines with cystein residues of the ß-chain of hemoglobin (Ringe et al., 1988Go).



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FIG. 1. Proposed activation pathways of 2,6-xylidine. Oxidation of 2,6-xylidine by CYP2A6 or CYP2E1 or by CYP2A6 forms DMAP and DMHA, respectively. DMHA has the potential to react with DNA or hemoglobin. It may also undergo phase II conjugation to reactive esters that can decompose spontaneously to a reactive nitrenium ion, which can react with DNA and other macromolecules. The nitrenium ion can also react with H2O to form DMAP. The latter can undergo a non-enzymatic oxidation to an iminoquinone, which is a strong electrophile. DMHA can also be rearranged to DMAP by a CYP2E1-catalyzed reactions (modified after Gan et al., 2001Go).

 
In rats and dogs DMAP is the major 2,6-xylidine metabolite found in urine (Short et al., 1989aGo). Studies using recombinant human CYPs and human liver microsomes have shown that DMAP is the only metabolic product at micromolar concentrations of 2,6-xylidine, whereas at nanomolar 2,6-xylidine concentrations DMHA was a substantial product (Gan et al., 2001Go). CYP2A6 and CYP2E1 were identified as the major CYPs responsible for the formation of DMAP, whereas CYP2A6 was identified as the CYP responsible for the formation of DMHA (Gan et al., 2001Go). DMHA can be rearranged to DMAP, and CYP2E1 was shown to catalyze this reaction (Gan et al., 2001Go).

2,6-Xylidine-hemoglobin adduct levels have been found to be elevated in human patients receiving lidocaine treatment for local anesthesia or cardiac arrhythmias, which is indicative that DMHA is a metabolite of lidocaine in man (Bryant et al., 1994Go). 2,6-Xylidine-hemoglobin adducts were also found in humans with no known exposure to lidocaine. This was attributed to chronic exposure to iatrogenic or environmental sources of 2,6-xylidine and to the 120-day life span of the erythrocyte (IARC 1993Go; Bryant et al., 1994Go). 2,6-Xylidine-hemoglobin adducts have in addition been found in rats given 2,6-xylidine or lidocaine (Bryant et al., 1994Go).

Covalent binding of 14C-2,6-xylidine to DNA has been shown in ethmoid turbinates of rats given the substance orally (Short et al., 1989bGo). In addition, a 32P-postlabeling assay has shown DNA adducts in the nasal mucosa of rats given 2,6-xylidine orally (Jeffrey et al., 2002Go).

In the present study we have used whole-body autoradiography with freeze-dried or solvent-extracted tissue sections to trace tissues accumulating extractable and nonextractable metabolites after iv or oral administration of 3H-labeled 2,6-xylidine in rats. Low-temperature whole-body radioluminography was used to examine the tissue deposition of the volatile 2,6-xylidine. Based on the whole-body autoradiography the detailed localization of tissue-bound 2,6-xylidine metabolites was examined by microautoradiography. In addition, the ability of subcellular fractions of some tissues to form 2,6-xylidine metabolites, which bind to protein and DNA, were studied in vitro.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals. 3H-2,6-xylidine (generally labeled) with a specific radioactivity of 10 Ci/mmol was purchased from Moravek Biochemicals (Brea, CA). Calf thymus DNA, glucose-6-phosphate dehydrogenase, glucose-6-phosphate, nicotinamide adenine dinucleotide (NADP), and metyrapone were purchased from Sigma-Aldrich Co. (St Louis, MO). Other chemicals used in the study were obtained from regular commercial sources.

Animals. Female non-pregnant Sprague-Dawley rats, body weight about 125 g, were obtained from Bantin and Kingman (Sollentuna, Sweden). The animals were housed at 22°C with a 12 h light/dark cycle with free access to tap water and a standard pellet diet (Lactamin AB, Vadstena, Sweden). The Local Ethics Committee for Animal Research approved the study.

Whole-body autoradiography and whole-body radioluminography. Rats were given 3H-2,6-xylidine (2 mCi [24 µg] /kg body weight, given in 60% ethanol [1.2 ml/kg body weight]) iv or po The rats were killed by CO2-asphyxiation after 30 min (one animal given 3H-2,6-xylidine iv) or 24 h (one animal given 3H-2,6-xylidine iv and one animal given 3H-2,6-xylidine orally). They were then embedded in carboxymethyl cellulose and put in a hexane bath cooled to –78°C with CO2-ice and used for whole-body sectioning according to the method of Ullberg (Ullberg et al., 1982Go). Twenty-µm-thick sagittal sections were taken through the entire bodies. To study the distribution of firmly bound radioactivity, every other freeze-dried tissue-section was extracted successively with 5% trichloroacetic acid, 50% ethanol, 99.5% ethanol, and heptane for 1 min, and then rinsed with tap water for 5 min. These sections, together with the non-extracted tissue-section, were dried and pressed against X-ray film for about 8 weeks (Hyperfilm-3H, Amersham, Great Britain). To study the distribution of the volatile nonmetabolized 3H-2,6-xylidine, the surface of each of the sectioned carboxymethyl cellulose–embedded rats was pressed for 7 days at –20°C against phosphor imaging plates (Storage Phosphor Screen, tritium-sensitive, type: TR). Radioluminographic images were then obtained by developing the plates using the Packard Cyclone Storage Phosphor System (Packard, Canberra Company, Japan).

Microautoradiography. For in vivo microautoradiography, one rat was injected iv with 3H-2,6-xylidine (4 mCi [48 µg]/kg body weight; given in 60% ethanol [2.4 ml/kg body weight]) and killed by CO2-asphyxiation after 24 h. Pieces of the olfactory and respiratory mucosa, trachea, lung, tongue, cheek, soft palate, esophagus, and forestomach were immediately removed and fixed in phosphate buffered 4% formaldehyde (pH 7.4). Following dehydration in an ethanol series the pieces were embedded in Technovit 100 (Heraeus Kulzer GmbN, Wehrheim, Germany). Two-µm-thick sections were cut on glass slides and covered with Kodak NTB-2 emulsion. Exposure was carried out for about 3 months at 4°C, followed by photographic development and fixation and staining of the sections with toluidine blue. The extensive extractions during the fixation and embedding procedures will remove all unbound radioactivity, and the autoradiograms will therefore show only tissue-bound labeling (Larsson, 1994Go).

For in vitro microautoradiography one non-pretreated rat was killed by CO2-asphyxiation. Pieces of the same tissues as those examined for in vivo microautoradiography (see above) were immediately removed and incubated for 1 h at 37°C in O2-atmosphere with 2 µCi 3H-2,6-xylidine (0.1 µM) in 2 ml Tris-HCl buffer pH 7.4, containing 5 mM glucose-6-phosphate. In some incubations the CYP-inhibitor metyrapone (500 µM) was present in the incubation medium. After the incubations the pieces were fixed in phosphate buffered 4% formaldehyde (pH 7.4) and used for microautoradiography, as described above.

Preparation of subcellular fractions. For preparation of S-9-fractions, pooled samples of various tissues were taken from two pools of 10 non-pre-treated rats killed by CO2-asphyxiation. The tissues were homogenized in 50 mM phosphate buffer, pH 7.4, and centrifuged at 9000 x g for 20 min. The obtained supernatants (S-9 fractions) were stored at –70°C until used.

For preparation of microsomal fractions, pooled samples of various tissues were taken from two pools of 10 non-pre-treated rats killed by CO2-asphyxiation. The tissues were homogenized in 50 mM Tris-HCl buffer, pH 7.4, containing 0.15 M KCl. The homogenates were sedimented at 10,000 x g for 25 min. The resulting supernatants were centrifuged at 105,000 x g for 1 h. The obtained microsomal pellets were washed in the Tris-HCl buffer and re-centrifuged at 105,000 x g for 1 h before re-suspension in the same buffer and storage at –70°C until used.

The protein contents in the microsomal pellets and the S-9-fractions were assayed by the method of Smith et al. (1985)Go, adapted for microplate readers.

In vitro formation of protein-bound 2,6-xylidine metabolites. To examine the capacity of the tissues to form protein-bound 2,6-xylidine metabolites, incubations were performed with S9-fractions in a total volume of 1 ml of O2-saturated 50 mM phosphate buffer containing the same NADPH-generating system as described below. The incubations were carried out at 37°C for 30 min in the buffer containing 0.15 mg protein and 2 µCi 3H-xylidine (0.2 µM). The incubations were stopped by adding 1 ml ice-cold methanol and 2 ml chloroform. The protein pellet was extracted according to the method of Baker and Van Dyke (1984)Go, and the amount of protein-bond 3H-xylidine metabolite was analyzed as described below. Samples of the liver, treated with 1 ml ice-cold methanol and 2 ml chloroform prior to the incubations, served as controls.

In vitro formation of DNA-bound 2,6-xylidine metabolites. To examine the capacity of the tissues to form DNA-bound 2,6-xylidine metabolites in vitro, incubations were performed with microsomal tissue preparations, 3H-2, 6-xylidine, and calf thymus DNA. Incubations were carried out at 37°C for 30 min in a total volume of 2 ml O2-saturated 50 mM Tris-HCL buffer, pH 7.4, containing 0.8 mM NADP, 5 mM glucose-6-phosphate, 1 U glucose-6-phosphate dehydrogenase, 3 mM MgCl2, 0.2 mg microsomal protein, 0.5 mg calf-thymus DNA, and 2 µCi 3H-xylidine (0.1 µM). The reaction was started by adding the 3H-2,6-xylidine after a 10-min preincubation. The incubations were stopped by the addition of 0.5 ml 4.5 M NaCl and 0.1 ml 3% sodium dodecylsulfate. The amount of 3H-2,6-xylidine metabolite bound to DNA was determined by extractions and liquid scintillation counting as described elsewhere (Tjälve et al., 1992Go), with the exception that the DNA-quantitation was performed with Pico Green Reagent, according to the method of Singer et al. (1997)Go. Samples of the liver treated with 0.5 ml 4.5 M NaCl and 0.1 ml 3% sodium dodecylsulfate prior to the incubation served as controls.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Whole-Body Radioluminography and Whole-Body Autoradiography
The results of the low-temperature radioluminography—which will retain volatile radioactive materials in the tissues—of the rat killed 30 min after iv injection of 3H-2,6-xylidine showed that the level of labeling in adipose tissues at various sites, such as abdominal and subcutaneous regions, was much higher than in other tissues of the body (Fig. 2A). A high degree of labeling was also present in some nasal glands. Considerable labeling was present in the wall and the adjacent lumen of the glandular part of the stomach. A moderate level of radioactivity was also seen in brain and spinal cord, with a preferential localization in the white matter. The radioluminographic technique makes it possible to detect a wide range of radioactivity levels in various tissues. The radioluminographic image in Figure 2A has been given a threshold that visualizes the most highly labeled structures in the body. Radioluminograms given thresholds that allow visualization of lower levels of radioactivity showed an evenly distribution in most other tissues of the body (data not shown).



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FIG. 2. Whole-body radioluminogram (A) and whole-body autoradiograms (B and C) of a rat killed 30 min after an iv injection of 3H-xylidine. A. A low-temperature radioluminogram B. An autoradiogram of a freeze-dried tissues-section. C. An autoradiogram of an extracted tissue-section. The radioluminography makes it possible to detect a wide range of radioactivity levels in various tissues. The low-temperature radioluminogram (A) has been given a threshold which visualizes the most highly labeled structures in the body, i.e., primarily adipose tissues at various sites. In the autoradiogram of the freeze-dried tissue section (B) the volatile nonmetabolized 3H-xylidine has disappeared, and the autoradiograms shows all the remaining radioactivity. In the autoradiogram of the extracted tissue section (C) only the 3H-xylidine metabolites that bound firmly to tissue constituents are visible. Abbreviations: af = abdominal fat; ct = connective tissues; e = mucosa of the esophagus; fs = forestomach; gs = glandular stomach; hb = heart blood; ic = intestinal contents; ng = nasal glands; o = nasal olfactory mucosa; sf = subcutaneous fat; t = mucosa of the tongue; u = urinary bladder.

 
In autoradiograms of freeze-dried tissue sections from of the rat killed 30 min after iv injection of 3H-2,6-xylidine, the volatile labeling had disappeared. In these autoradiograms a high level of labeling was observed in the nasal olfactory mucosa (Fig. 2B). High levels of radioactivity were also seen in the mucosa in various structures of the upper alimentary tract, such as the tongue, cheek, gingiva, soft palate, esophagus, and forestomach. A somewhat lower degree of labeling was present in the nasal respiratory mucosa and the tracheal, bronchial, and bronchiolar mucosa. In addition, some nasal glands showed a high degree of labeling. In the abdomen, the most highly labeled structures were the kidney (most prominent in the pelvis) and the contents of stomach, small intestine, and urinary bladder. Moderate labeling was present in the blood and—at a somewhat lower level—in the liver. In addition, the autoradiograms showed a considerable degree of labeling in connective tissues at various sites. There was no labeling of adipose tissues or the central nervous system (CNS).

Autoradiography of extracted tissue sections of the rat killed 30 min after iv injection of 3H-2,6-xylidine showed retention of high levels of bound material in the nasal mucosa and in the mucosa of the upper alimentary tract (Fig. 2C). A somewhat lower level of labeling was present in the tracheal, bronchial, and bronchiolar mucosa. A low level of bound radioactivity was also present in the blood. In addition, a low degree of labeling was retained in connective tissues, most markedly in the subcutis. The contents of the glandular stomach was also labeled. There was no detectable labeling of other tissues of the body.

Whole-body radioluminograms and whole-body autoradiograms of rats killed 1 day after iv or oral administration of 3H-2, 6-xylidine showed similar pictures, whether low-temperature radioluminography or autoradiography with freeze-dried tissue sections or extracted tissue sections were used. These pictures showed marked localization of radioactivity in the nasal mucosa and in the mucosa of the tissues of the upper alimentary tract (Fig. 3A). A somewhat lower level of labeling was present in the tracheal, bronchial, and bronchiolar mucosa. An even lower level of radioactivity was present in the blood and the connective tissues, and there was no detectable labeling in other tissues of the body.



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FIG. 3. A. Whole-body autoradiogram of a freeze-dried tissue section of a rat killed 1 day after oral administration of 3H-xylidine. B. The corresponding tissue-section. Abbreviations: e = mucosa of the esophagus; fs = mucosa of the forestomach; g = mucosa of the gingiva; o = nasal olfactory mucosa; sp = mucosa of the soft palate; t = mucosa of the tongue.

 
Microautoradiography
The in vivo microautoradiography, which was performed in a rat killed 1 day after iv injection of 3H-2,6-xylidine, showed labeling over cells of Bowman's glands in the nasal olfactory mucosa, whereas other structures in the olfactory area were unlabeled (Fig. 4A). In the nasal respiratory mucosa, the acini and ducts of some subepithelial serous glands were labeled, whereas the surface epithelium and subepithelial mucous glands were unlabeled (Fig. 4B). In the bronchioli (Fig. 4C), bronchi, and trachea (data not shown) some cells of the surface epithelia were labeled. The lung parenchyma was unlabeled. In the esophagus there was an accumulation of silver grains over the stratum corneum of the surface epithelium (Fig. 4D), and in the tongue, silver grains were localized over the superficial parts of the stratified squamous epithelium (Fig. 4E). Comparable pictures were seen in the gingiva, cheek, and soft palate (data not shown). In the forestomach silver grains were limited to a narrow zone in the deepest part of the stratum corneum (Fig. 4F).



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FIG. 4. In vivo microautoradiography of various tissues of a rat killed 1 day after an iv injection of 3H-xylidine. A. Nasal olfactory mucosa. B. Subepithelial glands of the nasal respiratory mucosa. C. Lung with a bronchiolus. D. Esophagus. E. Tongue. F. Forestomach. In panel A there is a labeling of Bowman's glands; in panel B silver grains are present over the serous gland but not over the mucous gland of the nasal respiratory mucosa; in panel C the silver grains are confined to the bronchiolar epithelium, whereas the lung parenchyma is unlabeled; in panel D the silver grains are localized over stratum corneum of the esophagus; in panel E silver grains are localized over the superficial part of the stratified squamous epithelium of the tongue; and in panel F silver grains are limited to a narrow zone in the deepest part of the stratum corneum of the forestomach. Abbreviations: be = bronchiolar epithelium; Bg = Bowman's gland; mg = mucous gland; n = neuronal cells; s = sustentacular cells; se = stratified squamous epithelium of the esophagus; sf = stratified squamous epithelium of the forestomach; sg = serous gland; st = stratified squamous epithelium of the tongue. Toluidine blue; panels A and D, magnification x200; C, magnification x250; B, E, and F, magnification x300.

 
The in vitro microautoradiography of the nasal olfactory mucosa incubated with 3H-2,6-xylidine showed preferential localization of silver grains over sustentacular cells and a lower degree of labeling of the cells of Bowman's glands (Fig. 5A). Incubations with nasal respiratory mucosa resulted in labeling of the surface epithelium and of the serous subepithelial glands as well (Fig. 5B). In the trachea, silver grains were present over some epithelial cells (Fig. 5C). Comparable pictures were seen in the bronchi and bronchioli (data not shown). In the esophagus, silver grains were localized over the cells of the stratified squamous epithelium, except the cells of the deepest layers and the stratum corneum (Fig. 5D). Comparable pictures were seen in the forestomach (data not shown). In the cheek and tongue, silver grains were localized over the cells of the stratified squamous epithelium, except the cells of the deepest layers (Fig. 5E and 5F). Comparable pictures were seen in the gingiva and soft palate (data not shown). Microautoradiograms obtained from tissues incubated with 3H-2.6-xylidrine together with metyrapone were devoid of silver grains.



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FIG. 5. In vitro microautoradiography of various rat tissues incubated with 3H-xylidine. A. Nasal olfactory mucosa. B. Nasal respiratory mucosa. C. Trachea. D. Esophagus. E. Cheek. F. Tongue. In panel A there is a preferential labeling of sustentacular cells; in panel B silver grains are present over the nasal respiratory epithelium and over the serous gland; in panel C the silver grains are confined to the tracheal epithelium; in panels D, E, and F silver grains are localized over all cells of the stratified squamous epithelia, except the cells of the deepest layers. Abbreviations: Bg = Bowman's gland; n = neuronal cells; re = nasal respiratory epithelium; s = sustentacular cells; se = stratified squamous epithelium of the esophagus; sg = serous gland; st = stratified squamous epithelium of the tongue. Toluidine blue; panel A, magnification x200; B, C, D, and E, magnification x300; F, magnification x150.

 
Formation of Protein-Bound 2,6-Xylidine-Metabolites by Various Tissues
Incubations with S-9-fractions showed that all the examined tissues had a capacity to activate 2,6-xylidine to protein-bound metabolites. The nasal olfactory mucosa showed the highest activating capacity. The activating capacity of the other tissues was in the following order: mucosa of cheek {approx} nasal respiratory mucosa > liver {approx} mucosa of tongue > mucosa of esophagus {approx} mucosa of forestomach (Table 1).


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TABLE 1 Formation of Protein-Bound Xylidine-Metabolites by Microsomal Preparations of Various Tissues from Rat

 
Formation of DNA-Bound 2,6-Xylidine-Metabolites by Various Tissues
Incubations with microsomal preparations showed that all the examined tissues had a capacity to form DNA-bound 2, 6-xylidine-metabolites. The nasal olfactory mucosa showed the highest activating capacity. The activating capacity of the other tissues was in the following order: nasal respiratory mucosa > mucosa of esophagus {approx} mucosa of cheek {approx} mucosa of tongue > mucosa of forestomach {approx} liver (Table 2).


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TABLE 2 Formation of DNA-Bound Xylidine-Metabolites by Microsomal Preparations of Various Tissues from Rat

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The results of the low-temperature whole-body radioluminography indicate that the non-metabolized 2,6-xylidine, which is a volatile and fat-soluble compound, is distributed throughout the body and accumulates in adipose tissues, e.g., in the abdominal and subcutaneous regions.

The autoradiography of the extracted whole-body tissue-sections, as well as the in vivo microautoradiography, showed bound 2,6-xylidine metabolites in the nasal olfactory mucosa and the mucosa of the upper alimentary and respiratory tracts. These tissues were found to have an in vitro capacity to bioactivate 2,6-xylidine. Thus, our data indicate that 2,6-xylidine in vivo undergoes an in situ bioactivation in these extrahepatic tissues.

It should be noted that the 2,6-xylidine-bioactivation in the extrahepatic tissues is confined to a few cells, such as Bowman's glands in the nasal olfactory mucosa; the acini and ducts of some serous glands in the nasal respiratory mucosa; some cells of the stratified squamous epithelium of the mouth, esophagus, and forestomach; and some cells of the surface epithelium of the trachea, bronchi, and bronchioli. The bioactivating capacity of these cells is reflected by the strong labeling in the autoradiograms and must be higher than is reflected by the in vitro data based on subcellular preparations of these heterogeneous tissues.

As mentioned in the Introduction, Gan et al., (2001)Go used recombinant human CYPs and human liver microsomes to study the initial CYP-mediated mono-oxygenation of 2,6-xylidine. They identified CYP2A6 and CYP2E1 as the major CYPs responsible for the formation of DMAP, i.e., hydroxylation of 2,6-xylidine in the para position on the aromatic ring, and CYP2A6 as being responsible for the formation of DMHA, i.e., hydroxylation of the nitrogen of 2,6-xylidine. In addition DMHA can be rearranged to DMAP by a CYP2E1-catalyzed reaction (Gan et al., 2001Go; see Fig. 1).

It is well known that high levels of some CYP-enzymes are present in the nasal olfactory mucosa and the mucosa of the upper alimentary and respiratory tracts (Ding and Kaminsky, 2003Go; Thornton-Manning and Dahl, 1997Go). CYP2A3, which is the rat ortholog of human 2A6, is expressed at high concentrations in the rat nasal mucosa (Thornton-Manning and Dahl, 1997Go). CYP2A3 has been shown in the esophagus of rats (Gopalakrishnan et al., 1999Go; Pinto et al., 2001Go). CYP2E1 has been shown in the rat nasal mucosa (Thornton-Manning and Dahl 1997Go). Shimizu et al. (1990)Go showed ethanol-inducible CYP2E1 in the squamous epithelial cells of the esophagus, cheek, tongue, and forestomach in rats. The mucosa cells of the tracheobronchial airways of rats also express CYP2E1 (Watt et al., 1998Go). It appears that relevant CYP-enzymes are present at the sites of 2,6-xylidine bioactivation. The microautoradiographic observation that the covalent in vitro binding of 2,6-xylidine-metabolites in these structures was completely blocked by the CYP inhibitor metyrapone confirms that the bioactivation is CYP dependent. However, further studies are required to determine the detailed role of the individual CYPs in the bioactivation of 2,6-xylidine in these tissues. It is also possible that CYPs other than 2A3/2A6 and 2E1 may be involved in the bioactivation of 2,6-xylidine at these sites.

Our results showed that the nasal olfactory mucosa had a much higher capacity than the other examined tissues to bioactivate 2,6-xylidine. The structures in the olfactory mucosa engaged in the bioactivation of 2,6-xylidine have been shown to contain high levels of CYP enzymes (Brittebo, 1997Go; Thornton-Manning and Dahl, 1997Go). Thus, the carcinogenic effect of 2,6-xylidine on the nasal mucosa is correlated with a high capacity of this tissue to bioactivate the compound. The in vivo microautoradiographs of the nasal mucosa indicate that the bioactivation of 2,6-xylidine is very active in Bowman's glands in the lamina propria, and that a considerable bioactivation also occurs in some serous submucosal glands. The in vivo microautoradiography indicated lack of bioactivation of 2,6-xylidine in the sustentacular cells, whereas the in vitro microautoradiography indicated a bioactivation in these cells. It is possible that under in vivo conditions the high metabolic capacity of Bowman's glands, which are situated close to the capillaries in the lamina propria, may prevent 2,6-xylidine from reaching the sustentacular cells.

As mentioned in the Introduction, administration 3000 ppm of 2,6-xylidine in the diet to rats for 4 weeks has been reported to result in atrophy of Bowman's glands in the dorsal meatus of the olfactory region of the nasal cavity (Yasuhara et al., 2000Go). The present data indicate that this effect can be related to the formation of cytotoxic 2,6-xylidine-metabolites in Bowman's glands. Conceivably Bowman's glands may also be the structures in the olfactory mucosa that undergo the malignant transformations. It has been shown that the herbicide 2,6-dichlorobenzonitrile (dichlobenil) administered parenterally to mice causes necrosis of the nasal olfactory mucosa after an initial bioactivation-related injury of Bowman's glands (Brandt et al., 1990Go). It has been shown that CYPs of the isoforms 2A6 and 2E1—i.e., those assumed to bioactivate 2,6-xylidine—are active in the metabolism of dichlobenil in the nasal mucosa (Ding et al., 1996Go).

There are no reports that the squamous epithelia of the esophagus, the mouth, and the forestomach, which were found to have a capacity to form DNA-binding 2,6-xylidine-metabolites, will undergo malignant transformations in rats treated with this compound. Neither are there any reports of 2,6-xylidine–induced tumors in the tracheo-bronchial mucosa. The reason for this is not known, but one possibility is that the potent bioactivation of 2,6-xylidine in the nasal mucosa leads to tumor development at this site before malignancies will develop at the other sites. With regard to the squamous epithelium of the upper alimentary tract, there is also a possibility that, because of the cellular turnover in the stratum corneum, the cells will be expelled before any malignant transformation occurs. Our results showed that the in vivo labeling of the stratified squamous epithelium of the esophagus, gingiva, tongue, soft palate, and forestomach—in contrast to the in vitro situation—was confined to the superficial layers of the epithelium. It is possible that under the in vivo conditions the cells in the squamous epithelium will bioactivate the 2,6-xylidine during a relatively short period when nonmetabolized substance is available for metabolism. At the 1-day survival interval, which was used for the microautoradiography, the labeled 2,6-xylidine–bioactivating cells may have been transferred toward the surface of the epithelium and replaced by basally located nonlabeled cells.

In humans, in contrast to rodents, the nasal mucosa is a very unusual site of tumorigenesis. Thus, chemical carcinogens, such as 2,6-xylidine, which in humans may be bioactivated in the nasal olfactory mucosa, as well as the mucosa of the upper alimentary and respiratory tissues, might potentially be expected to be more prone to induce tumors at the latter sites than in the nasal olfactory mucosa. Of special concern in this respect is the squamous epithelium of the upper alimentary tract, in which CYP2E1 may be induced by chronic ethanol intake, leading to xenobiotic activation and potentially ethanol-related tumorigenesis.

The 2-year carcinogenicity study of 2,6-xylidine showed a slightly increased incidence of subcutaneous fibromas in treated rats (Kornreich and Montgomery, 1990Go). It can be assumed that these tumors develop from fibroblasts in the subcutaneous connective tissue. Our whole-body autoradiography of the dried and extracted tissue sections showed uptake of labeled material in the subcutaneous connective tissues of the 3H-2,6-xylidine–treated rats. To our knowledge there are no reports that fibroblasts in connective tissues contain CYP-enzymes, which potentially might bioactivate 2,6-xylidine. Alternatively, a non-enzymatic bioactivation might occur in the fibroblasts. Thus, it is possible that hydrophilic DMHA-esters, formed in tissues with bioactivating capacity, will be translocated to the connective tissues, which may be reached via the fenestrated capillaries at these sites, and that they then may decompose non-enzymatically to an electrophilic nitrenium ion, as shown in Figure 1.

The low-temperature whole-body radioluminography of the rat killed 30 min after 3H-2,6-xylidine administration showed a considerable degree of labeling in the wall and the adjacent lumen of the glandular stomach. We assume that this represents protonated 2,6-xylidine. In the acidic milieu of the glandular stomach, the substance may undergo protonization at the nitrogen atom and therefore accumulate at this site as a result of ion-trapping. The whole-body autoradiography of extracted sections of this rat showed high levels of radioactivity bound to the contents of the glandular stomach. In contrast, the radioactivity in the other parts of the alimentary tract was extractable. The mechanism underlying the retention of bound labeling in the contents of the stomach is not known. However, one possibility is that nitrite ions reaching the stomach lumen via the saliva or feed react with the protonated 2,6-xylidine, which in the acidic milieu may form a positively charged electrophilic diazonium ion that reacts with feed components.

The whole-body autoradiographs showed bound labeling in the blood. We assume that this represents 2,6-xylidine hemoglobin adducts.

The nasal olfactory mucosa and the mucosa of the upper respiratory and alimentary tracts have previously been shown to have the capacity to perform CYP-related bioactivation of other compounds, such as the haloalkanes chloroform, carbon tetrachloride, and halothane (Tjälve and Löfberg, 1983Go; Löfberg and Tjälve, 1986Go; Ghantous et al., 1988Go), several N-nitrosamines (Tjälve 1991Go), the mycotoxin aflatoxin B1 (Larsson 1994Go), the sweet potato furanoterpenoid ipomeanol (Larsson and Tjälve, 1988Go), the herbicides dichlobenil and chlorthiamide (Brandt et al., 1990Go; Brittebo et al., 1991Go), and the 2,6-dichlorobenzene-metabolite methylsulfonyl-2,6-dichlorobenzene (Bahrami et al., 1999Go; Franzén et al. 2003Go). The CYP enzymes active in these tissues, which are portals of entry for foreign compounds, may normally play a role in defending the body from unrestrained uptake of xenobiotics. However, for chemicals that are bioactivated by CYP-dependent reactions, cellular injuries, or transformations leading to malignancies may instead be induced.


    ACKNOWLEDGMENTS
 
The technical assistance of Agneta Boström is gratefully acknowledged. This study was supported by the Swedish Research Council for Environmental, Agricultural Sciences and Spatial Planning (FORMAS).


    NOTES
 

1 To whom correspondence should be addressed at Department of Biomedical Sciences and Veterinary Public Health, Division of Pathology, Pharmacology and Toxicology, Swedish University of Agricultural Sciences, Box 7028, SE-750 07 Uppsala, Sweden. Fax: + 46 18 504144. E-mail: Pia.Larsson{at}farmtox.slu.se.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Bahrami, F., Brittebo, E. B., Bergman, A., Larsson, C., and Brandt, I. (1999). Localization and comparative toxicity of methylsulfonyl-2,5- and 2,6-dichlorobenzene in the olfactory mucosa of mice. Toxicol. Sci. 49, 116–123.[Abstract]

Baker, M. T., and Van Dyke, R. A. (1984). Metabolism-dependent binding of the chlorinated insecticide DDT and its metabolite, DDD, to microsomal protein and lipids. Biochem. Pharmacol. 33, 255–260.[CrossRef][ISI][Medline]

Brandt, I., Brittebo, E. B., Feil, V. J., and Bakke, J. E. (1990). Irreversible binding and toxicity of the herbicide dichlobenil (2,6-dichlorobenzonitrile) in the olfactory mucosa of mice. Toxicol. Appl. Pharmacol. 103, 491–501.[ISI][Medline]

Brittebo, E. B. (1997). Metabolism-dependent activation and toxicity of chemicals in nasal glands. Mutat/Res. 380, 61–75.[ISI][Medline]

Brittebo, E. B., Eriksson, C., Feil, V., Bakke, J., and Brandt, I. (1991). Toxicity of 2,6-dichlorothiobenzamide (chlorthiamid) and 2,6-dichlorobenzamide in the olfactory nasal mucosa of mice. Fundam. Appl. Toxicol. 17, 92–102.[ISI][Medline]

Bryant, M. S., Simmons, H. F., Harrell, R. E., and Hinson, J. A. (1994). 2,6-Dimethylaniline–hemoglobin adducts from lidocaine in humans. Carcinogenesis 15, 2287–2290.[Abstract]

Ding, X., and Kaminsky, L. S. (2003). Human extrahepatic cytochromes P450: Function in xenobiotic metabolism and tissue-selective chemical toxicity in the respiratory and gastrointestinal tracts. Annu. Rev. Pharmacol. Toxicol. 43, 149–73.[CrossRef][ISI][Medline]

Ding, X., Spink, D. C., Bhama, J. K., Sheng, J. J., Vaz, A. D., and Coon, M. J. (1996). Metabolic activation of 2,6-dichlorobenzonitrile, an olfactory-specific toxicant, by rat, rabbit, and human cytochromes P450. Mol. Pharmacol. 49, 1113–1121.[Abstract]

Dureja, P., Tanwar, R. S., and Choudhary, P. P. (2000). Identification of impurities in technical metalaxyl. Chemosphere 41, 1407–1410.[CrossRef][ISI][Medline]

Franzén, A., Carlsson, C., Brandt, I., and Brittebo, E. B. (2003). Isomer-specific bioactivation and toxicity of dichlorophenyl methylsulphone in rat olfactory mucosa. Toxicol. Pathol. 31, 364–372.[CrossRef][ISI][Medline]

Gan, J., Skipper, P. L., and Tannenbaum, S. R. (2001). Oxidation of 2,6-dimethylaniline by recombinant human cytochrome P450s and human liver microsomes. Chem. Res. Toxicol. 14, 672–677.[ISI][Medline]

Ghantous, H., Lofberg, B., Tjälve, H., Danielsson, B. R., and Dencker, L. (1988). Extrahepatic sites of metabolism of halothane in the rat. Pharmacol. Toxicol. 62, 135–141.[ISI][Medline]

Gopalakrishnan, R., Morse, M. A., Lu, J., Weghorst, C. M., Sabourin, C. L., Stoner, G. D., and Murphy, S. E. (1999). Expression of cytochrome P450 2A3 in rat esophagus: Relevance to N-nitrosobenzylmethylamine. Carcinogenesis 20, 885–891.[Abstract/Free Full Text]

IARC (International Agency for Research on Cancer). (1993) IARC Monographs on the Evaluation of the Carcinogenic Risks of Chemicals to Humans Vol.57, 2,6-dimethylaniline (2,6-xylidine), pp. 323–335, IARC Press, Lyons, France.

Irvine, W. J., and Saxby, M. J. (1969). Steam volatile amines of Lataki tobacco leaf. Phytochemistry 8, 473–476.[CrossRef][ISI]

Jeffrey, A. M., Luo, F. Q., Amin, S., Krzeminski, J., Zech, K., and Williams, G. M. (2002). Lack of DNA binding in the rat nasal mucosa and other tissues of the nasal toxicants roflumilast, a phosphodiesterase 4 inhibitor, and a metabolite, 4-amino-3,5-dichloropyridine, in contrast to the nasal carcinogen 2,6-dimethylaniline. Drug Chem. Toxicol. 25, 93–107.[CrossRef][ISI][Medline]

Keenaghan, J. B., and Boyes, R. N. (1972). The tissue distribution, metabolism and excretion of lidocaine in rats, guinea pigs, dogs and man. J. Pharmacol. Exp. Ther. 180, 454–463.[Medline]

Kornreich, M., and Montgomery, C. A., Jr. (1990). Toxicology and Carcinogenesis Studies of 2,6-xylidine (2,6-dimethylaniline) in Charles River CD Rats. National Toxicology Program (NTP). NTP TR 278, NIH Publication No. 90–2534, pp. 1–138.

Larsson, P. (1994). Biotransformation and retention of aflatoxin B1 in extrahepatic tissues of various animal species. Thesis from the Swedish University of Agricultural Sciences, Uppsala, Sweden (ISBN 91–576–4805–0).

Larsson, P., and Tjälve, H. (1988). Tracing tissues with 4-ipomeanol-metabolizing capacity in rats. Chem. Biol. Interact. 67, 1–24.[CrossRef][ISI][Medline]

Löfberg, B., and Tjälve, H. (1986). Tracing tissues with chloroform-metabolizing capacity in rats. Toxicology 39, 13–35.[CrossRef][ISI][Medline]

Marques, M. M., Mourato, L. L., Amorim, M. T., Santos, M. A., Melchior, W. B., Jr., and Beland, F. A. (1997). Effect of substitution site upon the oxidation potentials of alkylanilines, the mutagenicities of N-hydroxyalkylanilines, and the conformations of alkylaniline-DNA adducts. Chem. Res. Toxicol. 10, 1266–1274.[CrossRef][ISI][Medline]

Marques, M., Gamboa da Costa, G., Blankenship, L. R., Culp, S. J., and Beland, F. A. (2002). The effect of deuterium and fluorine substitution upon the mutagenicity of N-hydroxy-2,6-dimethylaniline. Mutat. Res. 506–507, 41–48.[ISI]

Martin, S. W., Bishop, F. E., Kerr, B. M., Moor, M., Moore, M., Sheffels, P., Rashed, M., Slatter, J. G., Berthon-Cedille, L., Lepage, F., Descombe, J. J., Picard, M., Baillie, T. A., and Levy, R. H. (1997). Pharmacokinetics and metabolism of the novel anticonvulsant agent N-(2,6-dimethylphenyl)-5-methyl-3-isoxazolecarboxamide (D2624) in rats and humans. Drug Metab. Dispos. 25, 40–46.[Abstract/Free Full Text]

Parker, R. J., Collins, J. M., and Strong, J. M. (1996). Identification of 2,6-xylidine as a major lidocaine metabolite in human liver slices. Drug Metab. Dispos. 24, 1167–1173.[Abstract]

Patrinakos, C., and Hoffman, D. (1979). Chemical studies of tobacco smoke. LXIV. On analysis of aromatic amines in cigarette smoke. J. Anal. Toxicol. 3, 150–154.[ISI]

Pinto, L. F., Moraes, E., Albano, R. M., Silva, M. C., Godoy, W., Glisovic, T., and Lang, M. A. (2001). Rat oesophageal cytochrome P450 (CYP) monooxygenase system: comparison to the liver and relevance in N-nitrosodiethylamine carcinogenesis. Carcinogenesis 22, 1877–1883.[Abstract/Free Full Text]

Puente, N. W., and Josephy, P. D. (2001). Analysis of the lidocaine metabolite 2,6-dimethylaniline in bovine and human milk. J. Anal. Toxicol. 25, 711–715.[ISI][Medline]

Pütter, J., and Sagner, G. (1973). Chemical studies to detect residues of xylazine hydrochloride. Vet. Med. Rev. 2, 145–159.

Ringe, D., Turesky, R. J., Skipper, P. L., and Tannenbaum, S. R. (1988). Structure of the single stable hemoglobin adduct formed by 4-aminobiphenyl in vivo. Chem. Res. Toxicol. 1, 22–24.[ISI][Medline]

Shimizu, M., Lasker, J. M., Tsutsumi, M., and Lieber, C. S. (1990). Immunohistochemical localization of ethanol-inducible P450IIE1 in the rat alimentary tract. Gastroenterology 99, 1044–1053.[ISI][Medline]

Short, C. R., Hardy, M. L., and Barker, S. A. (1989a). The in vivo oxidative metabolism of 2,4- and 2,6-dimethylaniline in the dog and rat. Toxicology 57, 45–58.[CrossRef][ISI][Medline]

Short, C. R., Joseph, M., and Hardy, M. L. (1989b). Covalent binding of [14C]-2,6-dimethylaniline to DNA of rat liver and ethmoid turbinate. J. Toxicol. Environ. Health 27, 85–94.[ISI][Medline]

Singer, V. L., Jones, L. J., Yue, S. T., and Haugland, R. P. (1997). Characterization of PicoGreen reagent and development of a fluorescence-based solution assay for double-stranded DNA quantitation. Anal. Biochem. 249, 228–238.[CrossRef][ISI][Medline]

Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985). Measurement of protein using bicinchoninic acid. Anal. Biochem. 150, 76–85.[ISI][Medline]

Thomas, J., Morgan, D., and Vine, J. (1976). Metabolism of etidocaine in man. Xenobiotica 6, 39–48.[ISI][Medline]

Thornton-Manning, J. R., and Dahl, A. R. (1997). Metabolic capacity of nasal tissue interspecies comparisons of xenobiotic-metabolizing enzymes. Mutat. Res. 380, 43–59.[ISI][Medline]

Tjälve, H., and Löfberg, B. (1983). Extrahepatic sites of metabolism of carbon tetrachloride in rats. Chem. Biol. Interact. 46, 299–316.[CrossRef][ISI][Medline]

Tjälve, H. (1991). The tissue distribution and the tissue specificity of bioactivation of some tobacco-specific and some other N-nitrosamines. Crit. Rev. Toxicol. 21, 265–294.[ISI][Medline]

Tjälve, H., Larsson, P., Andersson, C., and Busk, L. (1992). Bioactivation of aflatoxin B1 in the bovine olfactory mucosa: DNA binding, mutagenicity and induction of sister chromatid exchanges. Carcinogenesis 13, 1345–1350.[Abstract]

Ullberg, S., Larsson, B., and Tjälve, H. (1982). Autoradiography. In Biologic Application of Radiotracers (H.J. Glenn, ed.), pp. 56–108. CRC Press, Inc., Boca Raton, FL.

Watt, K. C., Plopper, C. G., Weir, A. J., Tarkington, B., and Buckpitt, A. R. (1998). Cytochrome P450 2E1 in rat tracheobronchial airways: Response to ozone exposure. Toxicol. Appl. Pharmacol. 149, 195–202.[CrossRef][ISI][Medline]

Yasuhara, K., Kobayashi, H., Shimamura, Y., Koujitani, T., Onodera, H., Takagi, H., Hirose, M., and Mitsumori, K. (2000). Toxicity and blood concentrations of xylazine and its metabolite, 2,6-dimethylaniline, in rats after single or continuous oral administrations. J. Toxicol. Sci. 25, 105–113.[Medline]

Zalipsky, J. J., Won, C. M., and Patel, D. M. (1978). Analytical-physical profile of lidamidine hydrochloride (WHR-1142A), a novel antidiarrheal agent. Arzneimittelforschung 28, 1441–1447.[Medline]





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