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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
J Toxicol Sci. Author manuscript; available in PMC 2012 December 12.
Published in final edited form as:
PMCID: PMC3520611

Dose-Response Assessment of Four Genotoxic Chemicals in a Combined Mouse and Rat Micronucleus and Comet Assay Protocol


The in vivo micronucleus (MN) assay has proven to be an effective measure of genotoxicity potential. However, sampling a single tissue (bone marrow) for a single indicator of genetic damage using the MN assay provides a limited genotoxicity profile. The in vivo alkaline (pH>13) Comet assay, which detects a broad spectrum of DNA damage, can be applied to a variety of rodent tissues following administration of test agents. To determine if the Comet assay is a useful supplement to the in vivo MN assay, a combined test protocol (MN/Comet assay) was conducted in male B6C3F1 mice and F344/N rats using four model genotoxicants: ethyl methanesulfonate (EMS), acrylamide (ACM), cyclophosphamide (CP), and vincristine sulfate (VS). Test compounds were administered on 4 consecutive days at 24-hour intervals (VS was administered to rats for 3 days); animals were euthanized 4 hours after the last administration. All compounds induced significant increases in micronucleated reticulocytes (MN-RET) in the peripheral blood of mice, and all but ACM induced MN-RET in rats. EMS and ACM induced significant increases in DNA damage, measured by the Comet assay, in multiple tissues of mice and rats. CP-induced DNA damage was detected in leukocytes and duodenum cells. VS, a spindle fiber disrupting agent, was negative in the Comet assay. Based on these results, the MN/Comet assay holds promise for providing more comprehensive assessments of potential genotoxicants, and the National Toxicology Program is presently using this combined protocol in its overall evaluation of the genotoxicity of substances of public health concern.

Keywords: DNA damage, Comet assay, acrylamide, ethyl methanesulfonate, cyclophosphamide, vincristine sulfate


Genotoxicity studies in rodents with defined exposures are useful biological test models for investigative toxicology and mechanistic studies, and they serve as an important element of the regulatory test battery used in pre-clinical safety assessment and the evaluation of environmental agents for genotoxic risk to humans (ICH, 1996; ICH, 1997; U.S. EPA, 2005). Compared with in vitro tests, in vivo tests may provide more relevant data for the assessment of DNA damage potential in humans since they take into account dynamic whole-animal physiological processes such as uptake and systemic distribution by the circulatory system, Phase I and Phase II metabolism, and intact elimination/excretory systems that cannot be entirely recreated in vitro. The rodent erythrocyte micronucleus (MN) assay in peripheral blood or bone marrow is considered to be the primary assay to assess in vivo genotoxic potential (Blakey et al., 2008; Eastmond et al., 2009; ICH, 2008). Micronuclei (MN) are surrogate measures of structural and numerical chromosomal aberrations that are associated with increased cancer risk (Bonassi et al., 2007). MN can also be considered bridging biomarkers of genotoxic exposure, since they can be enumerated across multiple species including humans (Dertinger et al., 2007).

Although the in vivo rodent MN assay is an effective measure of genotoxicity, the assay is not without limitations. The assay can only be conducted in rapidly dividing cells and typically measures chromosomal damage induced in a single tissue (bone marrow), thereby providing a limited assessment of genotoxic potential for a chemical. Since direct measurements of chromosomal aberrations or endogenous gene mutations in most tissues other than blood or bone marrow are not currently technologically feasible, a number of surrogate endpoints are used to assess mutagenicity and genotoxicity in other rodent tissues. These surrogate endpoints permit the evaluation of DNA damage, chromosomal damage, genomic responses to DNA damage, and mutation in marker genes (Guyton et al., 2009; ICH, 2008; Kirkland and Speit, 2008; Lambert et al., 2005). The alkaline (pH>13) Comet assay is being proposed by testing and regulatory agencies as a second (additional) in vivo genotoxicity bioassay, to complement the in vivo MN assay, since it can detect DNA repair and a broad spectrum of DNA damage, including DNA breaks, apurinic sites, alkali-labile DNA adducts, and a spectrum of reactive oxygen/lipid peroxidation species-induced DNA lesions in virtually any tissue (Fortini et al., 1996; Gedik and Collins, 2005). Furthermore, the Comet assay requires small numbers of cells and importantly, it does not require cell division for the evaluation of DNA damage. However, uncertainties with respect to the origin and fate of the detectable transient DNA lesions produced by xenobiotics may limit the use of the Comet assay to hazard identification (Fortini et al., 1996). The Comet assay is recommended as a follow-up to a negative or equivocal in vivo MN assay, as a confirmation to a positive MN assay, and as a means to measure genotoxicity in a target tissue other than bone marrow (e.g., liver) (Eastmond et al., 2009; ICH, 2008). An extensive international effort, led by the Japanese Center for the Validation of Alternative Methods (JaCVAM), is currently underway to conduct an in vivo Comet assay validation study in rats (

The National Toxicology Program (NTP) is evaluating an “acute” genotoxicity testing protocol in rodents that combines the MN and alkaline (pH>13) Comet assays for a more comprehensive assessment of genotoxicity in tissues of mice or rats (MN/Comet assay) than could be achieved with either assay alone. This manuscript reports results from initial studies evaluating a combined MN/Comet assay protocol using dose-response experiments to test four model genotoxic chemicals in male B6C3F1 mice and male Fisher 344/N rats. The four chemicals used in the studies were ethyl methanesulfonate (EMS), acrylamide (ACM), cyclophosphamide (CP), and vincristine sulfate (VS). These four chemicals induce MN by different mechanisms. ACM and EMS are both direct-acting clastogens, CP is clastogenic after metabolic activation, and VS induces aneuploidy (whole chromosome loss). The tissues or cell types examined were selected to: 1) examine DNA damage in the same accessible cell type used in human biomonitoring (leukocytes), 2) assess genotoxicity in a major site of xenobiotic metabolism (liver), and 3) evaluate genotoxicity in the gut region where most drug absorption takes place (duodenum). Additional tissues were assessed in ACM-treated animals based on prior knowledge of specific targets (i.e., testicular tissue). Here, we report the data from the alkaline (pH>13) Comet assay from the same B6C3F1 mice and Fisher 344/N rats used in MN assay studies reported earlier (Witt et al., 2008).

Materials and Methods


Details of chemicals, animal husbandry, and dosing were reported previously (Witt et al., 2008). Briefly, EMS (CASRN: 62-50-0), ACM (79-06-1), CP (50-18-0), and VS (57-22-7) were purchased from Sigma-Aldrich (St. Louis, MO, USA) and assigned code numbers prior to use in the experiments described below. ACM, EMS (both direct-acting clastogens), and CP (a clastogen that requires metabolic activation) were dissolved in phosphate buffered saline (pH 7.4) and administered by oral gavage. VS (an aneugen) was dissolved in phosphate buffered saline (pH 7.4) and administered by intraperitoneal injection due to the limited bioavailability of VS when administered by oral gavage. For these well-characterized genotoxic compounds, dose-setting information was available from previous studies conducted at ILS, Inc. or from published studies.

Animal Husbandry

Male B6C3F1 mice and male Fisher 344/N rats were used for this study. All studies were approved by the ILS, Inc. Institutional Animal Use and Care Committee. Procedures were completed in compliance with the Animal Welfare Act Regulations, 9 CFR 1-4, and animals were handled and treated according to the Guide for the Care and Use of Laboratory Animals (ILAR, 1996). Animals were acclimated for 7 days after receipt from the supplier (Charles River Laboratories, Portage, MI, USA). Animals were 8–16 weeks of age at the beginning of treatment; treatment groups consisted of five animals. Animals were maintained in constant temperature rooms (71 ± 3°F) with a relative humidity of 30–70% on a 12:12 (5:00 AM – 5:00 PM) light:dark cycle. Animals were housed individually in polycarbonate cages with Sanichip Laboratory hardwood bedding (P.J. Murphy Forest Products Corp., Montvale, NJ, USA) and provided food (Purina Certified Rodent Chow 5002, Ralston Purina, St. Louis MO, USA) and tap water ad libitum. With the exception of VS in rats, the animals were administered each test article once daily over four consecutive days (at 24-hr intervals), euthanized by carbon dioxide anesthesia/sedation, and then exsanguinated 4 hr after the last dosing. Due to cage-side observations indicative of toxicity in the rats, VS was administered for only 3 days and rats were euthanized 4 hr after the last exposure.

Peripheral blood samples were processed for flow cytometry evaluation of micronucleated reticulocytes (MN-RET) (Witt et al., 2008), and blood and tissue samples (blood, liver, duodenum, and additional tissues as noted below) were processed for assessment of DNA damage using the Comet assay (Burlinson et al., 2007; Hartmann et al., 2003; Tice et al., 2000).

Alkaline (pH>13) Comet Assay

The alkaline (pH>13) Comet assay was conducted according to published recommendations (Burlinson et al., 2007; Hartmann et al., 2003; Tice et al., 2000). At necropsy, samples (1 cm) of liver, duodenum, and blood (20μL) were collected from each animal and maintained in cold mincing buffer [Mg++ and Ca++ free Hanks' Balanced Salt Solution (Gibco, Carlsbad, CA, USA)) with 20 mM Na2EDTA (EDTA) (Sigma, St. Louis MO, USA) and 10% v/v dimethylsulfoxide (DMSO) (Fisher, Waltham, MA, USA)]. Liver and duodenum samples were minced in cold mincing buffer. For ACM-treated animals, additional tissues were collected as follows. A testis was removed from each animal and placed into mincing solution. Tubules were collected from the testis by dissection, and the contents of the tubules were flushed out into mincing solution. In ACM-treated rats only, thyroid (1 cm) was collected and placed into cold mincing solution and minced.

Aliquots of blood or minced tissue samples were immediately placed into microcentrifuge tubes and flash-frozen in liquid nitrogen; tubes were stored frozen below −60°C until the cells could be processed further. This process of cryopreservation allowed for control of a uniform time interval between tissue collection and electrophoresis (Witt et al., 2009). Slides for the Comet assay were prepared from blood and minced tissue as follows. Blood and tissue samples were thawed at room temperature and allowed to sit momentarily to permit large tissue pieces to settle. A 10 μL aliquot of suspension containing approximately 10,000 cells was mixed with 0.5% low melting point agarose (Sigma) and spread on standard microscope slides (Fisher) pre-dipped in agarose. Slides were then allowed to harden on a cold surface prior to further processing. All slides were placed in cold lysis solution (2.5 M NaCl, 100 mM EDTA, and 10 mM Trizma base, with 1% Triton X-100 (Sigma) and 10% DMSO). Following at least 1-2hr of incubation in lysis solution, two slides per sample were rinsed with 0.4 M Trizma base and incubated in alkaline conditions (300 mM NaOH (Fisher), 1 mM EDTA, pH>13) for 20 minutes, followed by electrophoresis in the same buffer for 20 minutes at 0.7 V/cm (electrode to electrode) and 300 mA. After electrophoresis, slides were immersed in an excess amount of 0.4 M Trizma base to neutralize the alkali and then fixed in 100% ethanol (McCormick, Weston, MO, USA). Following fixation, the slides were air dried and stored at room temperature in a desiccator until they were scored. Prior to scoring slides, the DNA was stained with SYBR Gold™, following the supplier's directions (Molecular Probes, Inc., Eugene, OR, USA). The slides were scored without knowledge of the dose group. The extent of DNA migration was determined for each sample by simultaneous image capture and scoring of 100 cells (50 cells on each of two slides) at 200× magnification using the Kinetic Imaging, Ltd., Komet© 5.5 image analysis system.

During the scoring of testes cell preparations from rats and mice, two distinct sizes of nuclei were noted and these were scored independently for each slide. The larger nuclei, similar in size to somatic cell nuclei in leukocytes, liver, and duodenum, were presumed to be from somatic cells. The smaller nuclei were presumed to contain one-half the DNA content of the somatic cells, reflecting sperm cell DNA. No further cell purification or characterization of cell origin was done on these mixed cell samples. These assumptions were used to classify each data set as DNA damage in presumptive somatic cells or sperm cells.

The extent of DNA migration for all samples was evaluated according to the following endpoint measurements:

% Tail DNA: intensity of all tail pixels divided by the total intensity of all pixels in the Comet, expressed as a percentage.

Tail Length: the horizontal distance from the center of the head (start of tail) to the end of the tail.

Olive Tail Moment (OTM): the distance between the center of gravity of the DNA distribution in the tail and the center of gravity of the DNA distribution in the head, multiplied by the fraction of DNA in the tail.

Statistical Analysis

Data from 100 cells per animal were collected and assessed for significant (P < 0.05) increases in DNA migration endpoints by statistical analysis using Analyse-it® Standard Edition software ( Using individual animal data, the Shapiro-Wilk test was used to assess normality of the negative control group. Data that were not normally distributed were analyzed by the Mann-Whitney test (Mann and Whitney, 1947) to compare each dose level to the concurrent control, and by the Kendall rank correlation test (Kendall, 1938) to determine the presence of a dose response. Data that were normally distributed were analyzed using the F test to determine homogeneity of variances, an independent one-tailed t-test to compare each dose level to the concurrent control, and linear regression to determine the presence of a dose response.


Ethyl Methanesulfonate

The alkaline (pH>13) Comet assay was conducted in blood leukocytes and in liver and duodenum cells of B6C3F1 male mice (Table 1) and F344/N male rats (Table 2) treated with 50, 100, 200, or 300 mg/kg EMS. One hundred cells were scored for each tissue examined per animal, and three measures of DNA damage were made for each sample: % Tail DNA, Tail Length, and OTM. The mean values from all animals per treatment group per tissue are presented in the data tables along with the results of the statistical analyses.

Table 1
Comet Assay Data for Mice Exposed to Ethyl Methanesulfonate
Table 2
Comet Assay Data for Rats Exposed to Ethyl Methanesulfonate

Based on the OTM endpoint, EMS induced significant increases in DNA damage in mice at all dose levels in all of the tissues examined. In rats, EMS also induced a significant, dose-related increase in DNA damage in all tissues examined. In leukocytes and in liver cells of rats, significant increases in DNA damage were detected at all dose levels. However, in contrast to the response seen in mice, significant increases in OTM in the rat duodenum were observed only at the highest administered dose of EMS (300 mg/kg).


The Comet assay was conducted in blood leukocytes and in cells of the liver, duodenum, and testes of B6C3F1 mice (Table 3) and F344/N rats (Table 4) administered 12.5, 25, 37.5 or 50 mg/kg ACM. For rats, cells from the thyroid were also examined (Table 4). In mice, ACM induced significant increases in DNA damage based on OTM in all of the tissues examined at all dose levels tested, with the single exception of testicular somatic cells of animals administered the lowest dose of ACM (12.5 mg/kg). In rats, the DNA damage response was more variable than in mice. ACM induced significant increases in DNA damage based on the OTM measurement in blood leukocytes, testicular somatic cells, and cells of the thyroid and duodenum; no increases in DNA damage were detected in rat liver cells or in presumptive sperm cells. Although statistically significant increases in DNA damage were observed in multiple tissues of rats, the overall magnitude of the increases was lower than that observed for the corresponding tissues in mice. In blood leukocytes from ACM-treated rats, low variability within the vehicle control group allowed the detection of small but statistically significant increases in DNA damage. In duodenum, testicular somatic cells, and thyroid cells, clear statistically significant increases in DNA damage were observed.

Table 3
Comet Assay Data for Mice Exposed to Acrylamide
Table 4
Comet Assay Data for Rats Exposed to Acrylamide


The Comet assay was conducted in blood leukocytes and in cells of the liver and duodenum of B6C3F1 mice and F344/N rats administered CP at doses of 25, 50, 75, or 100 mg/kg (Table 5) and 2.5, 5.0, 10 or 20 mg/kg (Table 6), respectively. In mice, CP induced significant (P < 0.05) increases in DNA damage in blood leukocytes, as indicated by OTM measurements, at all dose levels. No increase in DNA damage was seen in liver of CP-treated mice. In duodenum, a statistically significant increase was measured only in the highest dose group (100 mg/kg); the trend test was not significant. In rats, CP induced significant increases in DNA damage in blood leukocytes at the highest dose tested (20 mg/kg). No increase in DNA damage was detected in liver of CP-treated rats; in duodenum, significant increases were detected, but the magnitude of the increases was similar across doses.

Table 5
Comet Assay Data for Mice Exposed to Cyclophosphamide
Table 6
Comet Assay Data for Rats Exposed to Cyclophosphamide

Vincristine Sulfate

The Comet assay was conducted in blood leukocytes and in cells of the liver and duodenum of B6C3F1 mice and F344/N rats administered VS by intraperitoneal injection at doses of 0.0125, 0.0250, 0.0500 or 0.0750 mg/kg (Table 7) and 0.00625, 0.01250, 0.02500 or 0.03125 mg/kg (Table 8), respectively. Under the conditions used in this study, VS did not induce significantly increased levels of DNA damage in any of the tissues examined in mice or rats.

Table 7
Comet Assay Data for Mice Treated with Vincristine Sulfate
Table 8
Comet Assay Data for Rats Exposed to Vincristine Sulfate

Summary of Combined MN/Comet Assay in Mice and Rats

As indicated previously, the MN assay was conducted in the same animals used in the studies conducted by Witt et al. (2008). The data for the MN assays, along with the corresponding Comet assay results, are summarized in Table 9.

Table 9
Summary of Combined MN/Comet Assay Data in B6C3F1 Mice and F344/N Rats


There is a clear need to identify compounds having the potential to permanently alter the human genome. The genetic toxicology test battery addresses this need by evaluating genotoxicity resulting from chemical exposures (ICH, 1996; ICH, 1997; U.S. EPA, 2005). However, this test battery is used in the hazard identification step in risk assessment and is not designed to be a quantitative predictor of organ- or tissue-specific tumor induction in rodents (Elespuru et al., 2009). As the test battery continues to undergo revision and refinement, the Comet assay is being considered for use as a second in vivo genotoxicity assay (Eastmond et al., 2009; ICH, 2008). The Comet assay was recently shown to detect nearly 90% of carcinogens that were negative or equivocal in the MN assay and therefore, a combined MN/Comet assay has been recommended to broadly assess in vivo genotoxic potential (Kirkland and Speit, 2008; Pfuhler et al., 2007).

The purpose of the studies reported here was to evaluate a combined in vivo genetic toxicity testing protocol using the MN/Comet assay. The four model genotoxic compounds chosen—EMS, ACM, CP, and VS—are well described in the literature and dose-response studies were conducted to provide causal links to exposure. EMS, ACM, CP, and VS have all been previously tested in the Comet assay in vivo or in vitro, and they represent a variety of genotoxic modes of action (Anderson et al., 1995; Dearfield et al., 1988; Eastmond and Tucker, 1989; Gocke and Muller, 2009; Tice et al., 2000).

Using this combined MN/Comet assay protocol in mice and rats, a dose response for EMS genotoxicity was detected in all of the mouse tissues analyzed and in rat blood leukocytes and liver cells. In rat duodenum, EMS was positive only at the top dose of 300 mg/kg. Although EMS-induced DNA damage in the rat duodenum was not detectable with the Comet assay at doses up to 200 mg/kg, the increases in DNA damage seen in leukocytes and liver cells at these doses suggests that EMS absorption and entry into systemic circulation may occur proximal to the duodenum and reflects EMS-induced “first-pass” DNA damage in blood leukocytes and liver. If so, then the increased DNA damage observed in the rat duodenum at the top dose of EMS (300 mg/kg) may be due to incomplete absorption of EMS in the stomach, thus allowing some EMS to reach the duodenum at biologically effective levels detectable by the Comet assay.

Recent studies have examined the dose response for genotoxicity and mutagenicity of EMS administered for 28 days by oral gavage in the Muta™ mouse using the in vivo MN assay and the lacZ mutation assay (Gocke and Muller, 2009). No increases in lacZ mutations were observed with EMS doses of 25 – 50 mg/kg/day, and doses up to 80 mg EMS/kg/day did not induce a significant increase in MN-RET in bone marrow. In the present study, the Comet assay detected EMS-induced DNA damage in multiple tissues of mice and rats after four daily exposures of 50 mg/kg/day in the absence of an increased frequency of MN-RET. These results indicate that the Comet assay detects EMS-induced DNA lesions at lower levels in these tissues than are required for the MN assay to detect increases in MN-PCE due to bone marrow damage (Table 9; Witt et al., 2008). This difference in sensitivity may be due to the efficient repair of “low dose” EMS lesions prior to the cell division required for the formation of a chromosomal aberration detectable as a MN in reticulocytes. Higher EMS exposure levels (>50 mg/kg) may produce greater numbers of DNA lesions that are not as efficiently repaired, translating into chromosomal damage detectable as MN.

DNA damage, detected by the Comet assay, in multiple tissues of mice and rats administered ACM has been reported previously (Dobrzynska, 2007; Maniere et al., 2005). In the present study, ACM induced a significant dose-dependent increase in DNA damage in all of the tissues examined in mice (blood leukocytes, liver, duodenum, and gonadal cells). In ACM-treated rats, a dose-dependent increase in DNA damage detected by the Comet assay was observed in thyroid and presumptive testicular tubule somatic cells, and in blood leukocytes; no DNA damage was seen in liver or presumptive germ cells. The DNA damage seen in testes of mice and rats exposed to ACM is consistent with the well-documented germ cell genotoxicity and testicular toxicity of ACM in rodents (Dearfield et al., 1988; Yang et al., 2005). A variable but dose-responsive increase in DNA damage was seen in rat duodenum, with a clear increase above the background detected at the top dose of 50 mg/kg/day. ACM-induced genetic damage in somatic and germ cells is dependent upon the extent of metabolism of the parent compound to the genotoxic metabolite glycidamide, mediated by CYP2E1 (Dearfield et al., 1988; Dobrzynska, 2007; Ghanayem et al., 2005; Maniere et al., 2005; Witt et al., 2008). Compared to rats, mice produce higher levels of hemoglobin adducts derived from the bioactivation of ACM to glycidamide, likely accounting for the more extensive distribution of DNA damage observed in ACM-treated mice compared to rats (Doerge et al., 2005; Ghanayem et al., 2005; Sumner et al., 2003). Consistent with what is known about metabolism of ACM, MN-RET frequencies were elevated in mice, but not rats, treated with ACM (Table 9).

The antitumor agents CP and VS induced genotoxicity and bone marrow toxicity in mice and rats based on evaluations of MN-RET frequencies and percent RET data reported previously (Witt et al., 2008); rats were more sensitive to these effects than mice, necessitating the use of lower top doses in rats. These two antitumor drugs operate through two distinct modes of action: CP is bioactivated to reactive metabolites that produce a spectrum of DNA lesions resulting in chromosomal breakage and formation of MN. In contrast, VS-induced chromosomal damage is primarily numerical in nature (chromosome loss) and results from impaired microtubule assembly and subsequent chromosome malsegregation and loss (Anderson et al., 1995; Cushnir et al., 1990; Eastmond and Tucker, 1989). Thus, VS does not induce DNA damage (negative results in the Comet assay) but does, through non-DNA reactive mechanisms, induce aberrant mitoses, resulting in chromosome loss (aneuploidy) and production of MN. As predicted by their known modes of action, evaluation of the nature of the CP-induced MN in mice and rats revealed that the MN were primarily due to breakage events and contained chromosomal fragments, while VS-induced MN were likely due to aneuploidy events, since they contained larger amounts of chromosomal material, pointing to the presence of whole chromosomes, rather than fragments (Witt et al., 2008).

A dose response for CP-induced DNA damage was detectable in mice by the Comet assay in blood leukocytes, and at the top dose in duodenum cells; DNA damage was not observed in liver cells of CP-treated mice. In rats, significant increases in DNA damage were observed in duodenum cells and leukocytes at the top dose of CP. The absence of CP-induced DNA damage in liver of mice and rats suggests efficient detoxication of reactive metabolites. In the case of VS, there was no indication of DNA damage in blood leukocytes, liver, or duodenum cells under the conditions used in this study. The lack of detectable DNA damage using the Comet assay in mice or rats administered VS is consistent with microtubulin, rather than DNA, as a primary cellular target of VS.

The overall objective of this study was to initiate development of a database from which to evaluate the usefulness of integrating the Comet assay into the subacute dosing regimens used by the NTP in the evaluation of the genotoxicity of environmental agents of concern. The three-day exposure regimen adopted by the NTP for the in vivo MN assay was extended to include a fourth day of dosing, with final dosing occurring 4 hr prior to euthanasia, to meet the sample time requirements of the Comet assay (Tice et al., 1998). With this design, sample times for the assessment of these two genotoxicity endpoints comply with regulatory requirements stipulated for the MN assay (OECD 474) and recent recommendations for the conduct of the Comet assay (Burlinson et al., 2007). This combined MN/Comet assay protocol permits the evaluation of two distinct genotoxicity endpoints in the same animal, thereby reducing animal usage and cost, and providing a basis for integrating genotoxicity endpoints into traditional subacute toxicological studies in animals. The MN/Comet assay appears to be a useful combination of in vivo genotoxicity endpoints for hazard identification in preclinical safety assessment and the evaluation of environmental agents. Based on these results, the NTP is presently using this combined protocol as part of its efforts to evaluate the genetic toxicity of substances of public health concern.


The authors appreciate the skilled assistance of Cathy Baldetti and John Winters at ILS, Inc. in conducting the Comet assay. The authors are grateful to Drs. David DeMarini (US EPA), Daniel Shaughnessy (NIEHS), and Raymond Tice (NTP) for helpful comments and useful discussions during the preparation of this manuscript, and Claudine Gregorio at ILS, Inc. for critical review and editing of this manuscript.

Funding: This work was supported by the National Institute of Environmental Health Sciences/National Toxicology Program [contract number NO1-ES-35514].


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