Evaluating the weak in vivo micronucleus response of a genotoxic carcinogen, Aristolochic acids

Mutation Research 753 (2013) 82–92

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Evaluating the weak in vivo micronucleus response of a genotoxic carcinogen, Aristolochic acids夽 Javed A. Bhalli 1, Wei Ding, Joseph G. Shaddock, Mason G. Pearce, Vasily N. Dobrovolsky, Robert H. Heflich ∗ Division of Genetic and Molecular Toxicology, National Center for Toxicological Research, U.S. Food and Drug Administration, Jefferson, AR 72079, United States

a r t i c l e

i n f o

Article history: Received 21 November 2012 Received in revised form 1 March 2013 Accepted 3 March 2013 Available online 14 March 2013 Keywords: Bone marrow Red blood cells Pig-a assay Comet assay Hprt lymphocyte assay

a b s t r a c t Aristolochic acids (AAs) are carcinogenic plant toxins that are relatively strong gene mutagens, both in vitro and in vivo, but weak inducers of micronuclei in vivo. In order to clarify the reasons for these disparate responses, we evaluated the genotoxicity of AAs in F344 rats using several assays that respond to DNA damage in bone marrow. Groups of 7- to 8-week-old male rats (n = 6) were gavaged with 0, 2.75, 5.5, and 11 mg/kg AAs for 28 days or with 0, 11, 22, and 30 mg/kg AAs for 3 days. Day 1 being the first day of treatment, Pig-a mutant frequencies (MFs) were assayed in peripheral blood erythrocytes up to Day 56 for the 28-day treatment or Day 42 for the 3-day treatment; micronuclei were assayed in peripheral blood reticulocytes on Day 4 (both treatment protocols) and on Day 29 of the 28-day treatment protocol; and at the final sampling times (Day 59 or Day 42), the animals were sacrificed and Hprt mutant lymphocytes were measured. In a separate study, the Comet assay was performed on liver, kidney, and bone marrow of animals gavaged with 0, 11, 22, and 30 mg/kg AAs for 4 days and sacrificed 3 h after the last treatment. While only weak increases in micronucleated reticulocyte frequency were observed in treated animals, Pig-a MFs increased in a dose- and time-dependent manner with both treatment schedules. Lymphocyte Hprt mutant frequencies also increased dose dependently in treated animals, and the Comet assay detected elevated levels of DNA damage in all the tissues evaluated. These findings indicate that the DNA damage produced by AAs in rat bone marrow is a weak inducer of micronuclei but a relatively strong inducer of gene mutation. Published by Elsevier B.V.

1. Introduction Aristolochic acids (AAs) are a family of naturally occurring mutagenic and carcinogenic toxins produced by plants of the genus Aristolochia [1]. 8-Methoxy-6-nitro-phenanthro-(3,4-d)-1,3dioxolo-5-carboxylic acid (AAI) and 6-nitro-phenanthro-(3,4-d)1,3-dioxolo-5-carboxylic acid (AAII) are two major AAs occurring in plants as a mixture. AAI and AAII are metabolized similarly, to aristolactams, and form similar DNA adducts [2] (Fig. 1), although their structural differences appear to affect their genotoxicity [3]. Aristolochia species and their extracts have been used in a number of dietary supplements and traditional medicines. However, beginning in 1981, pharmaceutical preparations containing AAs

夽 Disclaimer: The views presented in this article are not necessarily those of the U.S. Food and Drug Administration. ∗ Corresponding author at: Division of Genetic and Molecular Toxicology, U.S. FDA/NCTR, 3900 NCTR Rd., HFT-120, Jefferson, AR 72079, United States. Tel.: +1 870 543 7493; fax: +1 870 543 7393. E-mail address: robert.hefl[email protected] (R.H. Heflich). 1 Present address: Covance, Inc., Greenfield, IN 46140, United States. 1383-5718/$ – see front matter. Published by Elsevier B.V. http://dx.doi.org/10.1016/j.mrgentox.2013.03.002

were withdrawn from the market in many countries [4]. Regulatory authorities in the United Kingdom, Canada, Australia, and Germany banned the use of products containing AAs for human consumption [5], and the U.S. Food and Drug Administration issued a warning advisory to consumers using AAs-based herbal medicines [6]. Herbal remedies containing Aristolochia are classified as carcinogenic to humans (Group 1) and in 2008 purified AAs were upgraded from Group 2A to Group 1 carcinogens by the International Agency for Research on Cancer [7]. However, despite a general appreciation of their adverse health effects, a wide variety of products containing extracts of Aristolochia are still in use in many regions worldwide [5]. AAs are positive in most in vitro genetic toxicity tests, including tests for gene mutation in bacteria [8,9], and tests measuring micronuclei, chromosomal aberration [10,11], and gene mutation in mammalian cell cultures [12,13]. In addition, in vivo studies indicate that AAs are relatively potent gene mutagens in various tissues of transgenic mice and rats [14–17]. These uniformly positive responses contrast with weak or poorly reproducible responses in the in vivo erythrocyte micronucleus (MN) assay [14,18]. AAs were classified as in vivo gene mutation positive, MN negative in an extensive review of in vivo genotoxicity assays by Lambert et al.

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Fig. 1. Aristolochic acids (AAs) structure, metabolic activation, and DNA adduct formation. (Adopted from Arlt et al. [2]).

[19], and equivocal for in vivo MN induction in the review by Kirkland and Speit [20]. Kohara et al. [14] reasoned that the weak MN responses in vivo could be due to AAs or their metabolites not reaching the bone marrow where MN formation takes place. An alternative hypothesis is that AAs are gene mutagens that are weak clastogens and poor inducers of micronuclei. This distinction is important, since the in vivo MN assay is the major in vivo genotoxicity assay used for regulatory safety assessments [21], and in vivo assays are often given more weight than in vitro assays when evaluating the potential hazards of human exposure (e.g., [22]). Ideally any battery for detecting genotoxic hazards should be able to identify Group I genotoxic carcinogens with a high degree of certainty. The Comet assay and transgenic gene mutation assays previously have been conducted on various tissues of AAs-treated mice and rats [14–17,23]; however, other than MN assays, no assays have measured AAs genotoxicity in the bone marrow. In order to examine if AAs damage bone marrow DNA and to determine the consequences of that damage, we have measured the DNA damage, erythrocyte MN induction, and gene mutation induced in the hematopoietic tissues of treated rats. Gene mutation was

measured using the erythrocyte Pig-a assay (reviewed in [24]) and the lymphocyte Hprt assay [25], assays that primarily detect the consequences of DNA damage to bone marrow cells [26–28]. We used the Comet assay to compare the relative levels of DNA damage formed in the bone marrow with that formed in the liver and kidney. 2. Materials and methods 2.1. Reagents A 1:1 mixture of AAI and AAII (CAS no. 313-67-7) was purchased from Acros Organics (Janssen Pharmaceuticalaan, Geel, Belgium). Methyl methanesulfonate (MMS) was obtained from Sigma (St. Louis, MO). Heparin anticoagulant solution and all reagents necessary to perform the MN analysis were from In Vivo Rat MicroFlow® PLUS kits (v090203; Litron Laboratories, Rochester, NY). Many of the reagents necessary for conducting the erythrocyte Pig-a assay (e.g., anti-CD59-PE and anti-CD61-PE antibodies, and SYTO 13 nucleic acid stain) were provided in Prototype In Vivo Rat MutaFlow® kits supplied by Litron. Anti-PE Magnetic Beads, LS Columns, and QuadroMACSTM separators were from Miltenyi Biotec (Auburn, CA). Additional reagents used for the Pig-a assay included CountBrightTM Absolute Count Beads, fetal bovine serum (FBS), and Ca++ and Mg++ free phosphate buffered saline (PBS) from Invitrogen (Carlsbad, CA), and Lympholyte® -Mammal cell separation reagent from Cedarlane Laboratories (Burlington, NC).

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2.2. Experimental design: animal treatments, blood and tissue collection

2.5. Lymphocyte Hprt assay

All experimental procedures involving animals were approved by the Institutional Animal Care and Use Committee (IACUC) of the National Center for Toxicological Research (NCTR). F344 male rats from the NCTR breeding colony were housed in conventional animal rooms, 2–3 rats per cage; individual animals were identified by ear clipping. Animals were allowed to acclimate for two weeks before starting the treatment. Water and food were available ad libitum throughout the acclimation and experimental period. The study was conducted in four experiments. All treatments were administered by gavage in a volume of 10 mL/kg body weight using PBS (pH 7.6) as the vehicle. In Experiment 1, a dose range-finding study was conducted by treating groups of 7- to 8-week-old male rats (n = 2) with 0, 5, 10, 15, 20, 25, and 30 mg/kg/day of the AAs mixture for 28 consecutive days and monitoring the animals’ weight and physical signs of distress. The dose resulting in a weight gain over the treatment period of approximately 0% to 20% that of the control rats was used as the high dose for the subsequent experiments. In Experiment 2, groups of 7- to 8-week-old male rats (n = 6) were treated with 0, 2.75, 5.5, 11, and 22 mg/kg/day of the AAs mixture for 28 consecutive days. Due to excessive toxicity, the 22 mg/kg dosing was discontinued after 7 days. On Days -1, 4, 15, and 56, with Day 1 being the first day of dosing (there was no Day 0), 80 ␮L of peripheral blood were collected from the tail vein of each animal into a tube containing 100 ␮L of heparin solution. On Day 29, 160 ␮L of blood were collected into 200 ␮L of heparin solution. Blood collected on Days -1, 15, 29, and 56 was used for the Pig-a assay. Blood collected on Days 4 and 29 was used to measure percent micronucleated reticulocytes (%MN-RETs). On Day 56, the animals were sacrificed and their spleens were collected for the lymphocyte Hprt assay. In Experiment 3, groups of 7- to 8-week-old male rats (n = 6) were treated with 0, 11, 22, and 30 mg/kg/day of the AAs mixture for 3 consecutive days. Eighty ␮L peripheral blood samples were collected as in Experiment 2, but on Days -1, 4, 15, 29, and 42. %MN-RETs were measured with the blood collected on Day 4, and the Piga assay was conducted with blood collected on Days -1, 15, 29, and 42. The animals were sacrificed on Day 42 and spleen lymphocytes isolated from the animals were used for the lymphocyte Hprt assay. In Experiment 4, the rats were treated for three consecutive days with AAs as in Experiment 3, but they were given an additional dose on Day 4 and the animals were euthanized 3 h later. As a positive control for the Comet assay, an additional group of six age-matched rats was treated with a single dose of 100 mg/kg MMS at the time of the last AAs treatment on Day 4 and sacrificed together with the AAs-treated rats. Bone marrow, kidney, and liver were collected and used for the Comet assay and pieces of liver and kidney were preserved in 10% formaldehyde for subsequent histopathological examination. In order to facilitate processing the samples, the treatments were staggered so that 2 animals from each treatment and control group were treated and assayed on three consecutive weeks.

Hprt mutant frequencies were measured in rat spleen T-cells using a modification of a method previously described for mice [30]. Spleen white blood cells were stimulated to proliferate by culturing overnight in a medium supplemented with 4 ␮g/mL concanavalin A (Worthington Biochemicals, Freehold, NJ), followed by dispensing into 96-well plates for limiting-dilution cloning in the presence and absence of 6-thioguanine (Sigma). After 10-days of incubation for clone development, the culture medium was supplemented with alamarBlue® viability indicator (1% final concentration; TREK Diagnostic Systems, Cleveland, OH); the plates then were incubated for an additional 24 h, and the number of wells containing viable clones was identified on each plate using a Tecan SpectraFluor fluorescence plate reader (Tecan U.S., Durham, NC). Frequencies of 6-thioguanine-resistant Hprt mutants were calculated by applying Poisson statistics.

2.3. Reticulocyte/total red blood cell Pig-a assay 2.3.1. Blood labeling and mutant enrichment The prototype “High Throughput” (HT) version of the Pig-a assay described by Dertinger et al. [29] was used in this study. Note that this version of the assay passes counting beads through the magnetic columns used for mutant enrichment, and is different in this regard from the protocol given in the Instruction Manual (Version 130131) that currently is included with the MutaFlow kit used for the assays. The current Instruction Manual recommends adding counting beads only after magnetic mutant enrichment. 2.3.2. Pig-a data acquisition A FACSCanto II flow cytometer (BD Biosciences), equipped with 488 nm and 633 nm lasers and running FACSDivaTM 6.1.2 software, was used for data acquisition and analysis. Default filter sets were used; nucleic acid dye (SYTO 13) fluorescence excited by the 488 nm laser was evaluated with a photomultiplier tube (PMT) equipped with a 530/30 nm emission filter, anti-CD59-PE fluorescence excited by the 488 nm laser was evaluated with a PMT equipped with a 585/42 filter, and counting bead fluorescence excited by the 633 nm laser was evaluated with a PMT equipped with a 660/20 filter. For “Pre-Column” samples, data acquisition was stopped after counting 200 beads. For “Post-Column” samples, data acquisition was limited by time; the samples were processed for 4 min at the high flow rate, which consumed almost the entire sample. This timing was sufficient to acquire ≥20,000 beads, and to interrogate the equivalent of ∼100 × 106 total RBCs and up to 3 × 106 reticulocytes (RETs) for CD59 deficiency (Pig-a mutation). 2.4. Reticulocyte micronucleus assay MN assays were performed on blood specimens collected on Days 4 and 29 in Experiment 2, and on Day 4 in Experiment 3 using In Vivo Rat MicroFlow® PLUS kits and by following the manufacturer’s instructions. The %MN-RET frequency was determined among ∼20,000 CD71-positive RETs for each animal; the %RETs out of total RBCs for each sample was recorded as an index of test agent toxicity to the hematopoietic system.

2.6. Alkaline Comet assay 2.6.1. Tissue dissociation For liver and kidney, cells were dissociated by mincing the tissues in mincing solution (Hank’s Balanced Salt Solution with 20 mM EDTA and 10% dimethylsulfoxide) using sharp scissors and then filtering the minced tissues through a 40 ␮m cell strainer (Fisher Scientific, Pittsburgh, PA). Bone marrow cells were isolated from femurs by flushing directly into mincing solution. 2.6.2. In vivo alkaline Comet assay The alkaline Comet assay was performed as described by Ding et al. [31]. Two slides were scored for each tissue/treatment; at least 100 cells were selected randomly from each slide and scored using a system consisting of a Nikon 50i fluorescence microscope and Comet IV imaging software (Perceptive Instruments, Wiltshire, UK). Percent of DNA in tail (% Tail DNA) was used as the parameter for assessing overall DNA damage. 2.6.3. Histopathology studies As an aid for interpreting the Comet assay results, liver and kidney tissues were evaluated for cytotoxicity by histopathology examination. Tissues were embedded in paraffin blocks using a paraffin-based infiltrating medium (Formula R® ; Leica Microsystems, Buffalo Grove, IL). Embedded tissues were sectioned at approximately 5 microns, stained with hematoxylin and eosin, and examined by microscopy. Non-neoplastic lesions were graded for severity as 1 (minimal), 2 (mild), 3 (moderate), or 4 (marked). 2.7. Statistical analysis RBCCD59− , RETCD59− , %MN-RET, Hprt lymphocyte, and Comet data were squareroot transformed (when necessary) to approximate normal distributions. The data were analyzed using one-way analysis of variance (ANOVA), followed by pair-wise comparisons of responses for individual treatment groups to the vehicle control using Dunnett’s test. All statistical tests were two-tailed, used P <0.05 to indicate significance, and were conducted using SigmaPlot version 11.0 software (Systat Software, San Jose, CA).

3. Results 3.1. Toxicity of AAs in rats 3.1.1. Weight gain In Experiment 1, the 30 mg/kg dose produced considerable weight loss over the 28 days of treatment, but was tolerated as an acute dose (3 or 4 daily treatments; Fig. 2a); 20 and 25 mg/kg/day resulted in little or no weight gain over the 28 days of treatment, and 22 mg/kg/day was initially chosen as the top dose for Experiment 2. Weight gain data obtained in Experiment 2 (28-day treatment) are shown in Fig. 2b. Initially, rats in this experiment were dosed with 5.5, 11, and 22 mg/kg AAs. Because of a precipitous loss of weight in animals given 22 mg/kg/day AAs, dosing was terminated on Day 7 and a second trial was initiated that included a vehicle control and a group of rats dosed at 2.75 mg/kg/day. Data from the two trials were analyzed separately although, for simplicity, they are pooled for the graphical representations shown in the figures. Overall, 28 days of dosing with 2.75, 5.5, and 11 mg/kg/day AAs resulted in weight gains that were approximately 88, 54, and 10% of the vehicle control group. In Experiment 3, rat weights were monitored over the course of the mutant manifestation period, until Day 42 (Fig. 2c). After the 3-day dosing regimen, rats in the

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Fig. 2. F344 rat weight gains during the dosing and mutant manifestation periods in Experiments 1–3. The administered doses are in mg/kg/day. (A) Weight gain during 28 days of daily dosing with AAs to identify a maximum tolerated dose (MTD). Data are the average of 2 rats/dose. (B) Weight gain during the 28 days of daily dosing with AAs in Experiment 2. Data are means calculated from 6 rats/dose. Dosing was stopped at Day 7 for the group given 22 mg/kg. (C) Weight gains during the 3 days of treatment with AAs and during the six weeks of mutant manifestation after the last treatment in Experiment 3. Data are means calculated from measurements made on 6 rats/dose.

30 mg/kg/day and 22 mg/kg/day groups initially experienced loss of weight; however, animals in both groups began gaining back weight by Day 7. At the termination of the study on Day 42, rats in the 11, 22, and 30 mg/kg/day groups had gained 72, 31, and 18% of the weight gained by the vehicle controls.

3.1.2. Erythropoietic system toxicity %RETs were monitored during the Pig-a and MN assays conducted in Experiments 2 and 3 as a measure of toxicity to the erythropoietic system. Dose-related decreases in %RETs were observed on Day 4 in both Experiments. The %RET frequencies for all

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Fig. 3. Percent micronucleated reticulocytes (%MN-RETs; shaded bars) and percent reticuloctyes (%RETs; line) in F344 rats treated with (A) 28 and (B) 3 daily doses of AAs. %RETs (plotted with solid lines) were measured by anti-CD71 staining. Data given as mean ± SEM (n = 6). Responses significantly different from the vehicle control are marked with an asterisk (*).

dose groups recovered to near vehicle control levels at later sampling times for both the 28- and 3-day treatment groups (Fig. 3a and b; MN-related data also shown in Fig. 4).

frequencies in rats treated with 11, 22, and 30 mg/kg/day AAs were all similar to the vehicle control (Fig. 3b). 3.3. Reticulocyte/total red blood cell Pig-a assays

3.1.3. Histopathology observations Sections of liver and kidney from rats sacrificed for the Comet assay (Experiment 4) were stained and observed microscopically. The only histopathology findings were dose-related minimal to mild degeneration of the renal tubules in the kidney (Table 1; also see Supplementary Figure). The degeneration involved the renal tubular epithelial cells lining the proximal tubules. No AAs-related microscopic findings were recorded for the liver. 3.2. Micronucleus assays A small increase in %MN-RETs was observed for the 11 mg/kg/day dose group in the assays conducted on Day 4 in Experiment 2; on Day 29, all dose groups (including the 11 mg/kg/day group) had %MN-RET frequencies that were similar to the vehicle control (Fig. 3a). In Experiment 3, the Day 4 %MN-RET

Pig-a mutant frequencies (MFs) were determined on Days -1, 15, 29, and 56 in rats treated for 28 days as part of Experiment 2, and on Days -1, 15, 29, and 42 in rats treated for 3 days as part of Experiment 3. Pig-a MFs were calculated using the numbers of CD59-deficient mutant total RBCs (RBCCD59− ) and mutant RETs (RETCD59− ) detected in magnetically enriched samples and the corresponding numbers of total RBC and RET equivalents processed. In both experiments, RBCCD59− and RETCD59− frequencies were low in all dose groups at Day -1 (before the treatment with AAs) and in the vehicle control rats at all sampling time-points, with most MFs being less than 5 × 10−6 . RBCCD59− and RETCD59− frequencies for the AAs-treated animals displayed significant, time- and dosedependent increases in both experiments (Figs. 5 and 6). In Experiment 2 (28-day dosing), rats treated with 5.5 and 11 mg/kg/day AAs had significantly elevated RBCCD59− and

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Fig. 4. Relative reticulocyte (RET) fraction in peripheral blood from AAs-treated F344 rats during (A) Experiment 2 and (B) Experiment 3. The fraction of RETs in the treated animals is expressed as a percent of the corresponding vehicle control frequency, with the control frequency set at 100%. %RETs for the Day 4 data were derived from anti-CD71 staining, while data for the other sampling times were from SYTO 13 staining.

RETCD59− frequencies on Day 15, and all the groups treated with AAs had elevated RBCCD59− and RETCD59− frequencies on Days 29 and 56 (Fig. 5a, b). While both the RBCCD59− and RETCD59− frequencies increased with sampling time, the RETCD59− frequencies generally were greater than the RBCCD59− frequencies at the Day 15 and Day 29 sampling time-points; by Day 56, the RBCCD59− and RETCD59− frequencies were similar. Table 1 Histopathology data obtained from kidney and liver of vehicle control and AAs treated F344 rats. Organ

Kidney

Liver

a

Treatment group

Vehicle 11 mg/kg AAs 22 mg/kg AAs 30 mg/kg AAs Vehicle 11 mg/kg AAs 22 mg/kg AAs 30 mg/kg AAs

Renal tubule degeneration.

Data # Examined

# Affected

Avg. severity

7 6 6 6 7 6 6 6

0 0 5a 6a 0 0 0 0

– – 1.4 1.7 – – – –

In Experiment 3 (3-day dosing), all RBCCD59− and RETCD59− frequencies measured in AAs-treated rats on Days 15, 29, and 42 were elevated significantly (Fig. 6a, b). While RBCCD59− frequencies in treated rats displayed a steady increase with sampling time, the RETCD59− frequencies reached dose-related maxima on Day 15. RETCD59− frequencies then decreased on Day 29 and remained at the same level on Day 42. As was the case with the last sampling time-point in Experiment 2, similar RBCCD59− and RETCD59− frequencies were observed for the different dosing groups on Day 42. Even though the daily doses given to the animals were higher in Experiment 3 than Experiment 2, by the termination of both Experiments (Day 42 and Day 56), the larger cumulative low, medium, and high doses used in Experiment 2 produced approximately 2-3fold higher RBCCD59− and RETCD59− frequencies than the cumulative low, medium, and high doses in Experiment 3. 3.4. Lymphocyte Hprt assays Lymphocyte Hprt assays were performed on Day 56 in Experiment 2 and on Day 42 in Experiment 3 (Fig. 7a, b). All treated animals from both experiments had significantly higher Hprt MFs than their vehicle control. The responses measured in both

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Fig. 5. Pig-a mutant frequencies in F344 rats given 28 daily doses of AAs in Experiment 2. (A): RETCD59− frequencies and (B): RBCCD59− frequencies. Data given as the mean ± SEM (n = 6); mutant frequencies significantly different from the vehicle control are marked with an asterisk (*).

experiments were dose-related, and, even though the daily doses were nominally lower, the low, medium, and high doses used for the 28-day treatment (Experiment 2) produced higher MFs than the low, medium, and high doses used in Experiment 3. In addition, similar to the erythrocyte Pig-a MFs, the 28-day treatment with 11 mg/kg/day AAs resulted in an approximately 5-fold greater Hprt MF than did the same dose administered for 3 days only. 3.5. Comet assay The alkaline Comet assay conducted in Experiment 4 detected significant increases in DNA damage in bone marrow, liver, and kidney (Fig. 8). There were no obvious dose-related trends in % Tail DNA in kidney and bone marrow samples, and a somewhat weak dose-response in liver samples. The magnitude of the increases in DNA damage was similar in the three tissues. A robust response

was observed in all tissues from the MMS-treated animals (positive control). 4. Discussion Previous studies indicate that AAs are strong in vitro genotoxins and strong in vivo gene mutagens; yet, when measured in mice, AAs are poor inducers of micronuclei [14,18]. Here we confirmed the negative/weak findings for in vivo MN induction in rats. With one exception, AAs tested negative in the three MN assays that we conducted in this study. The 11 mg/kg/day AAs treatment produced an ∼2-fold increase in %MN-RETs in the MN assays conducted on Day 4 of Experiment 2. No increase in MN frequency was observed in animals treated with this dose on Day 29 in Experiment 2 and at this or any higher dose on Day 4 in Experiment 3. These studies were conducted at doses approaching the limits of toxicity allowed by our IACUC guidelines (Fig. 2). Although AAs produced micronuclei,

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Fig. 6. Pig-a mutant frequencies in F344 rats given 3 daily doses of AAs in Experiment 3. (A): RETCD59− mutant frequencies and (B): RBCCD59− mutant frequencies. Data given as the mean ± SEM (n = 6); mutant frequencies significantly different from the vehicle control are marked with an asterisk (*).

chromosomal aberrations, and sister chromatid exchange in vitro [10,11], the available data indicate that the MN assay is at best only weakly sensitive to the genotoxicity of AAs in vivo. The weak MN responses contrast with the relatively strong responses for gene mutation in the erythrocyte Pig-a assay and lymphocyte Hprt assay. Pig-a mutation was measured with the recently developed HT version of the assay. This assay is capable of detecting 2- to 3-fold increases in Pig-a MF above background due to its ability to interrogate high numbers of RETs and total RBCs [29]. The power of the HT assay, however, probably was unnecessary to detect the in vivo mutagenicity of AAs. The maximum fold-increases for RBCCD59− and RETCD59− frequencies in Experiment 2 were 137 and 128, and in Experiment 3 were 25 and 19. Also the responses in the Hprt lymphocyte assay were unambiguously positive and doserelated. The maximum fold-increases for the lymphocyte Hprt assay (55 in Experiment 2 and 8 in Experiment 3) were somewhat less than for the Pig-a assay, probably due to higher background MFs in the Hprt assay.

Given that Pig-a and Hprt mutations are largely induced by DNA damage to cells in the bone marrow, we conclude that genotoxic species are reaching this tissue. This conclusion is supported by the DNA damage detected by the Comet assay in bone marrow (Fig. 8) as well as the suppression of the erythropoietic system evidenced as reductions in %RETs in AAs-treated rats (Fig. 3). Thus, the weak responses in the MN assay indicate that the DNA damage produced by AAs (Fig. 1) is much more likely to cause gene mutation than chromosome breakage and micronucleus induction, at least in erythropoietic precursor cells at the localized exposure levels attained in the present study. We have previously argued that Hprt and Pig-a MFs are best compared using RBCCD59− frequencies and lymphocyte Hprt frequencies assayed 5 to 6 weeks after the treatment [32]. With this in mind, we considered Day 56 of a 28-day treatment protocol, as was conducted for Experiment 2, or Day 42 of an acute treatment protocol, as was conducted in Experiment 3, to provide a reasonable period for Hprt mutant manifestation and data for comparing

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Fig. 7. Hprt lymphocyte mutant frequencies in F344 rats administered (A) 28 or (B) 3 daily doses of AAs. Hprt mutant frequencies were evaluated as 6-thioguanine-resistant lymphocytes in cells isolated from spleens collected from the animals on Day 56 in Experiment 2 and on Day 42 in Experiment 3. Data are means of six animals per dose group ± SEM; responses significantly different from the vehicle control are marked with an asterisk (*).

Hprt MFs with Pig-a MFs. As indicated above, the fold-increases for AAs-induced mutation in the two assays are different, due in part to the higher vehicle control frequencies for the lymphocyte Hprt assay. However, the net increases (induced MFs) are quite similar; for instance, the 11 mg/kg/day treatment in Experiment 2 produced a net increase of 144 × 10−6 in the Pig-a total RBC assay and 166 × 10−6 in the lymphocyte Hprt assay. Previous data from rodent assays indicate that, for potent gene mutagens, the Hprt and Pig-a assays produce similar levels of induced mutants (e.g., N-ethyl-N-nitosourea and benzo[a]pyrene) [32,33], but for potent clastogens (e.g., X-rays), the Hprt assay often is more sensitive than the Pig-a assay [34]. The nearly equal net Pig-a and Hprt MFs in the present study are consistent with AAs being strong gene mutagens. Most genotoxicity safety assessments have relied primarily on in vivo assays that detect aneugenicity and clastogenicity in rodent hematopoietic cells, like the erythrocyte MN assay [21]. Many in vivo gene mutagens also induce micronuclei; however, there are many examples of rodent carcinogens that are negative in in vivo rodent MN assays [19,20,35,36]. While some of these carcinogens may be operating through a nongenotoxic mode of action, the in vivo genotoxicity of others may be missed because the agent

does not reach the bone marrow or, as appears to be the case with AAs, the agent is a much better gene mutagen than clastogen or aneugen. Option 2 of the recent International Conference on Harmonization S2(R1) Guideline [22] at least partially compensates for the relatively low sensitivity of the in vivo MN assay by requiring a second in vivo genotoxicity test in a tissue other than the bone marrow. The liver Comet assay is mentioned specifically as a second test, and our data indicate that the liver (or kidney) Comet assay is successful in complementing the erythrocyte MN assay in detecting the in vivo genotoxicity of AAs. It is unclear if toxicity is a confounding factor for the in vivo Comet assay [37], and even though toxicity was detected in this study, it was moderate, and detected only in kidney and thus unlikely to account for the positive responses in the assay in the liver. The absence of a clear dose response for DNA damage may be due to saturating a pathway involved in producing the DNA damage (as might be expected if AAs-induced damage were produced indirectly). It also could be due to a narrow window for detecting AAs-induced DNA damage with the in vivo Comet assay. Nesslany et al. [23] found that AAs-induced DNA damage in rat kidney was detected only 22–26-h following the treatment. In

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Fig. 8. Induction of DNA damage in F344 rats treated with AAs for four consecutive days and sacrificed 3 hr after the last treatment. A single dose of MMS (100 mg/kg) was used as the positive control. Data obtained for kidney, bone marrow, and liver are expressed as % Tail DNA (n = 6 or 7 ± SEM). Dose groups significantly different from the vehicle control are marked with an asterisk (*).

our studies doses were administered both 3 and 24 h prior to tissue sampling, which included the window for detecting AAs-induced DNA damage observed by Nesslany. However, the multiple treatments that we used, coupled with any toxicity to the tissues, may have blurred any dose response for AAs-induced DNA damage. In this respect, because they respond to the cumulative effects of DNA damage over time, the Hprt and Pig-a gene mutation assays may have been at an advantage in detecting the dose-response genotoxicity of AAs. In summary, there are several examples of genotoxic carcinogens that are negative in the erythrocyte MN assay because they do not damage bone marrow (e.g., diethylnitrosoamine) [38]; the Comet assay, conducted in a target tissue for toxicity, is probably the most practical way of detecting the in vivo genotoxicity of such agents. However, the results from this study indicate that AAs are negative in the erythrocyte MN assay, not because they do not damage the bone marrow, but because they are much better inducers of gene mutation than micronuclei. Stankowski et al. [39] recently reported that 28 daily doses of 4-nitroquinoline-1-oxide resulted in positive responses in the Pig-a assay but negative responses for erythrocyte MN induction (also negative for peripheral blood chromosome aberration and for liver, stomach and peripheral blood Comet). Given these observations, and the relative ease with which the erythrocyte Pig-a assay can be integrated with the MN assay in repeat dose general toxicology studies, it might be prudent to complete the analysis of systemic genotoxicity in bone marrow by combining assays for clastogenicity/aneugenicity (MN) with gene mutation (Pig-a). At minimum these data demonstrate the value of assaying mechanistically different endpoints in evaluating the in vivo genotoxicity of test agents.

Funding This work was supported by institutional funds from the U.S. Food and Drug Administration/National Center for Toxicological Research. Prototype kits supplied free of charge by Litron Laboratories were used to conduct the Pig-a assays. The contribution by Litron Laboratories was supported by a grant from the U.S. National Institute of Environmental Health Sciences, no. R44ES018017. JAB

was supported by a postdoctoral appointment administered by the Oak Ridge Institute for Science and Education. Conflict of Interest The authors declare that there are no conflicts of interest. Acknowledgements We are grateful to Litron Laboratories for their gift of Prototype In Vivo Mutaflow kits and to Dr. Kelly Davis of Toxicologic Pathology Associates for conducting the histopathological examination of kidney and liver. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.mrgentox. 2013.03.002. References [1] NTP (National Toxicology Program), Report on Carcinogens. U.S. Department of Health and Human Services, Public Health Service, National Toxicology Program, Twelfth edition, pp 45-49, 2011. http://ntp.niehs.nih.gov/ ntp/roc/twelfth/roc12.pdf [2] V.M. Arlt, M. Stiborova, H.H. Schmeiser, Aristolochic acid as a probable human cancer hazard in herbal remedies: a review, Mutagen. 17 (2002) 265–277. [3] G. Xing, X. Qi, M. Chen, Y. Wu, J. Yao, L. Gong, T. Nohmi, Y. Luan, J. Ren, Comparison of the mutagenicity of aristolochic acid I and aristolochic acid II in the gpt delta transgenic mouse kidney, Mutat. Res. 743 (2012) 52–58. [4] M. Stiborova, E. Frei, V.M. Arlt, H.H. Schmeiser, Metabolic activation of carcinogenic aristolochic acid, a risk factor for Balkan endemic nephropathy, Mutat. Res. 658 (2008) 55–67. [5] D.A. Kessler, Cancer and herbs, N. Engl. J. Med. 342 (2000) 1742–1743. [6] FDA (U.S. Food and Drug Administration), Aristolochic Acid: FDA warns consumers to discontinue use of botanical products that contain Aristolochic acid, 2001. http://www.fda.gov/food/dietarysupplements/alerts/ucm096388.htm [7] Y. Grosse, R. Baan, K. Straif, B. Secretan, F. El Ghissassi, V. Bouvard, L. BenbrahimTallaa, N. Guha, L. Galichet, V. Cogliano, A review of human carcinogens – Part A: pharmaceuticals, Lancet Oncol. 10 (2009) 13–14. [8] G. Robisch, O. Schimmer, W. Goggelmann, Aristolochic acid is a direct mutagen in Salmonella typhimurium, Mutat. Res. 105 (1982) 201–204.

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