Introduction to Assay Design
Genetic toxicology assays are used to identify germ cell mutagens, somatic cell mutagens, and potential carcinogens. These assays can detect diverse kinds of genetic alterations (eg, gene mutations, chromosome aberrations, and aneuploidy) that are relevant to the production of adverse human health outcomes. Over the last three decades, hundreds of chemicals and complex mixtures have been evaluated for genotoxic effects. Genetic toxicology assays serve two interrelated but distinct purposes in the toxicologic evaluation of chemicals: (1) identifying mutagens for purposes of hazard identification and (2) characterizing dose–response relationships and mutagenic mechanisms, both of which contribute to an understanding of genetic and carcinogenic risks.
A common experience when surveying the genetic toxicology literature is encountering a bewildering array of assays in viruses, bacteria, fungi, cultured mammalian cells, plants, insects, and mammals. More than 200 assays for mutagens have been proposed, and useful information has been obtained from many of them. Although most genetic toxicology testing and evaluation relies on relatively few assays, data from relatively obscure assays can sometimes contribute to a judgment about the genetic activity of a compound.
Table 9-1 lists key assays that have a prominent place in genetic toxicology. Table 9-2 is a more comprehensive list that provides literature citations to many of the assays that one might encounter in the genetic toxicology literature. Even this extensive table is not exhaustive, in that it emphasizes methods in applied genetic toxicology and not those assays whose use has been largely restricted to studies of mutational mechanisms. The commonly used assays rely on phenotypic effects as indicators of gene mutations or small deletions and on cytological methods for observing gross chromosomal damage. Detailed information on assay design, testing data, controls, sample sizes, and other factors in effective testing is found in the references cited.
Table 9-1Principal Assays in Genetic Toxicology ||Download (.pdf) Table 9-1 Principal Assays in Genetic Toxicology
A well-characterized assay for gene mutations The Salmonella/mammalian microsome assay (Ames test)
A mammalian assay for chromosome damage in vivo: metaphase analysis in rodent bone marrow or micronucleus assay in rodent bone marrow or blood
Other assays offering an extensive database or unique genetic endpoint
Assays for gene mutations
E coli WP2 tryptophan reversion assay
TK or HPRT forward mutation assays in cultured mammalian cells
Drosophila sex-linked recessive lethal assay
Gene-mutation assays in rodent somatic cells or transgenic animals
Cytogenetic analysis in cultured Chinese hamster or human cells
Other indicators of genetic damage
Mammalian germ cell assays
Mouse specific-locus tests
Cytogenetic analysis and heritable translocation assays
DNA damage and repair in rodent germ cells
Mutation analysis in tandem-repeat loci in mice
Table 9-2Overview of Genetic Toxicology Assays ||Download (.pdf) Table 9-2 Overview of Genetic Toxicology Assays
|ASSAYS ||SELECTED LITERATURE CITATIONS |
|I. Prediction of genotoxicity || |
| A. Interpretation of chemical structure || |
| Structural alerts to genotoxicity ||Ashby, (1994), Ashby and Tennant (1991), Ashby and Paton (1993) |
| B. In silico predictive models || |
| Computational and structural programs: MCASE, TOPKAT, DEREK ||Snyder et al. (2004), Snyder and Smith (2005), Snyder (2009), Mahadevan et al. (2011) |
| Quantitative structure–activity relationship (QSAR) modeling ||Votano et al. (2004), Snyder and Smith (2005), Mahadevan et al. (2011) |
|II. DNA damage and repair assays || |
| A. Direct detection of DNA damage: || |
| Alkaline elution assays for DNA strand breakage in hepatocytes ||Storer et al. (1996), Gealy et al. (2007) |
| Comet assay (single-cell gel electrophoresis) for DNA strand breakage ||Fairbairn et al. (1995), Singh (2000), Tice et al. (2000), Collins (2004), Olive (2009) |
| Comet-FISH assay for region-specific DNA damage and repair ||Glei et al. (2009), Shaposhnikov et al. (2009) |
| Nonmammalian comets in ecotoxicology ||Cotelle and Férard (1999), Lee and Steinert (2003), Jha (2004) |
| Assays for chemical adducts in DNA ||Phillips et al. (2000), Farmer and Singh (2008), Himmelstein et al. (2009) |
| B. DNA repair, recombination, and genotoxic stress responses as indicators of damage: || |
| Differential killing of repair-deficient and wild type bacteria ||Takigami et al. (2002) |
| Induction of the bacterial SOS system ||Quillardet and Hofnung (1993), Yasunaga et al. (2004), Oda et al. (2009) |
| “Green Screen” for GADD45a gene induction in TK6 human cells ||Hastwell et al. (2009), Jagger et al. (2009) |
| Unscheduled DNA synthesis (UDS) in isolated rat hepatocytes or rodents in vivo ||Madle et al. (1994), Kirkland and Speit (2008) |
| Induction of mitotic recombination ||Zimmermann (1992), Hoffmann (1994), Vogel and Nivard (2000), Turner et al. (2003), Reliene et al. (2007) |
|III. Prokaryote gene mutation assays || |
| A. Bacterial reverse mutation assays: || |
| Salmonella/mammalian microsome assay (Ames test) ||Ames et al. (1975), Kier et al. (1986), Maron and Ames (1983), Gatehouse et al. (1994), Mortelmans and Zeiger (2000), Seifried et al. (2006), Claxton et al. (2010) |
| E coli WP2 tryptophan reversion assay ||Gatehouse et al. (1994), Mortelmans and Riccio (2000) |
| Salmonella-specific base-pair substitution assay (Ames II assay) ||Gee et al. (1994), Kamber et al. (2009) |
| E coli lacZ-specific reversion assay ||Cupples and Miller (1989), Cupples et al. (1990), Josephy (2000), Hoffmann et al. (2003) |
| B. Bacterial forward mutation assays: || |
| E coli lacI assay ||Calos and Miller (1981), Halliday and Glickman (1991) |
| Resistance to toxic metabolites or analogs in Salmonella ||Jurado et al. (1994), Vlasakova et al. (2005) |
|IV. Assays in nonmammalian eukaryotes: || |
| A. Fungal assays: || |
| Forward mutations, reversion, and small deletions ||Zimmermann et al. (1984), Crouse (2000) |
| Mitotic crossing over, gene conversion, and homology-mediated deletions in yeast ||Zimmermann et al. (1984), Zimmermann (1992), Howlett and Schiestl (2000), Daigaku et al. (2004), Freeman and Hoffmann (2007) |
| Genetic detection of mitotic and meiotic aneuploidy in yeast ||Zimmermann et al. (1984), Aardema et al. (1998), Howlett and Schiestl (2000), Nunoshiba et al. (2007) |
| B. Plant assays: || |
| Gene mutations affecting chlorophyll in seedlings, the waxy locus in pollen, or Tradescantia stamen-hair color ||Ma et al. (2005), Grant and Owens (2006) |
| Chromosome aberrations and micronuclei in mitotic and meiotic cells of corn, Tradescantia, and other plants ||Ma et al. (2005), Grant and Owens (2006), MisÍk et al. (2011) |
| C. Drosophila assays: || |
| Sex-linked recessive lethal test in germ cells ||Lee et al. (1983), Mason et al. (1987), Vogel et al. (1999) |
| Heritable translocation assays ||Mason et al. (1987), Vogel et al. (1999) |
| Mitotic recombination and LOH in eyes or wings ||Vogel et al. (1999), Vogel and Nivard (2000) |
|V. Mammalian gene mutation assays || |
| A. In vitro assays for forward mutations: || |
| tk mutations in mouse lymphoma or human cells ||Clements (2000), Seifried et al. (2006), Wang et al. (2009), Moore et al. (2011) |
| hprt or xprt mutations in Chinese hamster or human cells ||Li et al. (1988), DeMarini et al. (1989), Parry et al. (2005) |
| CD59 mutations in CHO-human hybrid AL cells ||Zhou et al. (2006), Ross et al. (2007) |
| B. In vivo assays for gene mutations in somatic cells: || |
| Mouse spot test (somatic cell specific-locus test) ||Styles and Penman (1985), Lambert et al. (2005) |
| hprt mutations (6-thioguanine-resistance) in rodent lymphocytes ||Cariello and Skopek (1993), Casciano et al. (1999), Lambert et al., 2005 |
| Pig-a mutations (immunological detection of mutations blocking glycosylphosphatidylinositol synthesis) ||Dobrovolsky et al. (2010), Miura et al. (2011), Dobo et al. (2011) |
| C. Transgenic assays: || |
| Mutations in the bacterial lacI gene in “Big Blue” mice and rats ||Lambert et al. (2005), Singer et al. (2006) |
| Mutations in the bacterial lacZ gene in the “Muta Mouse” ||Lambert et al. (2005), Singer et al. (2006) |
| Mutations in the phage cII gene in lacI or lacZ transgenic mice ||Swiger (2001), Lambert et al. (2005) |
| Point mutations and deletions in the lacZ plasmid mouse ||Lambert et al. (2005), Singer et al. (2006) |
| Point mutations and deletions in delta gpt mice and rats ||Okada et al. (1999), Lambert et al. (2005) |
| Forward mutations and reversions in ΦX174 transgenic mice ||Valentine et al. (2010) |
| Inversions and deletions arising in pKZ1 mice by intrachromosomal recombination ||Sykes et al. (2006) |
|VI. Mammalian cytogenetic assays || |
| A. Chromosome aberrations: || |
| Metaphase analysis in cultured Chinese hamster or human cells ||Ishidate et al. (1988), Kirkland et al. (1990), Galloway et al. (1994), Galloway (2000), Corvi et al. (2008), Galloway et al. (2011) |
| Metaphase analysis of rodent bone marrow or lymphocytes in vivo ||Preston et al. (1981), Kirkland et al. (1990), Tice et al. (1994) |
| Chromosome painting and other FISH applications in vitro and in vivo ||Tucker et al. (1993b, 2005), Paccierotti and Sgura (2008) |
| B. Micronuclei: || |
| Cytokinesis-block micronucleus assay in human lymphocytes ||Fenech et al. (2003), Corvi et al. (2008), Fenech (2008), Fenech et al. (2011a,b) |
| Micronucleus assay in mammalian cell lines ||Kirsch-Volders et al. (2003), Corvi et al. (2008), Galloway et al. (2011) |
| In vivo micronucleus assay in rodent bone marrow or blood ||Krishna and Hayashi (2000), Hayashi et al. (2007), Tweats et al. (2007), Dertinger et al. (2011) |
| In vivo micronucleus assay in tissues other than marrow or blood ||Hayashi et al. (2007), Coffing et al. (2011), Morita et al. (2011) |
| C. Sister chromatid exchange: || |
| SCE in human cells or Chinese hamster cells ||Tucker et al. (1993a), Wilson and Thompson (2007) |
| SCE in rodent tissues, especially bone marrow ||Tucker et al. (1993a) |
| D. Aneuploidy in mitotic cells: || |
| Hyperploidy detected by chromosome counting or FISH in cell cultures or bone marrow ||Galloway and Ivett (1986), Natarajan (1993), Aardema et al. (1998), Paccierotti and Sgura (2008) |
| Micronucleus assay with centromere/kinetochore labeling in cell cultures ||Lynch and Parry (1993), Natarajan (1993), Aardema et al. (1998), Fenech (2008), Paccierotti and Sgura (2008) |
| Altered parameters in flow-cytometric detection of micronuclei in CHO cells ||Bryce et al. (2011) |
| Mouse bone marrow micronucleus assay with centromere labeling ||Adler (1993), Aardema et al. (1998), Krishna and Hayashi (2000) |
|VII. Germ cell mutagenesis || |
| A. Measurement of DNA damage || |
| Molecular dosimetry based on mutagen adducts in reproductive cells ||Russell and Shelby (1985), Olsen et al. (2010), Verhofsrad et al. (2011) |
| UDS in rodent germ cells ||Bentley et al. (1994), Sotomayor and Sega (2000) |
| Alkaline elution assays for DNA strand breaks in rodent testes ||Bentley et al. (1994) |
| Comet assay in sperm and gonadal tissue ||Speit et al. (2009) |
| B. Gene mutations || |
| Mouse specific-locus test for gene mutations and deletions ||Russell et al. (1981), Ehling (1991), Russell and Russell (1992), Favor (1999), Russell (2004), Singer et al. (2006) |
| Mouse electrophoretic specific-locus test ||Lewis (1991) |
| Dominant mutations causing mouse skeletal defects or cataracts ||Ehling (1991), Selby et al. (2004) |
| ESTR assay in mice ||Yauk (2004), Dubrova (2005), Singer et al. (2006), Somers (2006) |
| Germ cell mutations in transgenic assays ||Lambert et al. (2005), Singer et al. (2006) |
| C. Chromosomal aberrations || |
| Cytogenetic analysis of oocytes, spermatogonia, spermatocytes, or zygotes ||Kirkland et al. (1990), Tease (1992), Russo (2000), Marchetti et al. (2001) |
| Direct detection in sperm by FISH ||Marchetti et al. (2006, 2008) |
| Micronuclei in mouse spermatids ||Russo (2000), Hayashi et al. (2007) |
| Mouse heritable translocation test ||Russell and Shelby (1985), Singer et al. (2006) |
| D. Dominant lethal mutations || |
| Mouse or rat dominant lethal assay ||Adler et al. (1994), Singer et al. (2006) |
| E. Aneuploidy || |
| Cytogenetic analysis for aneuploidy arising by nondisjunction ||Allen et al. (1986), Adler (1993), Adler et al. (1994), Aardema et al. (1998), Russo (2000), Marchetti et al. (2001) |
| Sex chromosome loss test for nondisjunction or breakage ||Russell and Shelby (1985) |
| Micronucleus assay in spermatids with centromere labeling ||Aardema et al. (1998) |
| FISH with probes for specific chromosomes in sperm ||Russo (2000), Marchetti et al. (2006, 2008) |
Some assays for gene mutations detect forward mutations whereas others detect reversion. Forward mutations, such as those detected in the thymidine kinase gene (tk) in the widely used assay in mouse lymphoma cells (Clements, 2000; Wang et al., 2009), are genetic alterations in a wild type gene that are detected by a change in phenotype caused by the alteration or loss of gene function. In contrast, back mutations are mutations that restore gene function in a mutant, such as the histidine revertants detected in the Ames assay in Salmonella (Ames et al., 1975; Mortelmans and Zeiger, 2000; Claxton et al., 2010). Thus, a back mutation or reversion that restores gene function in a mutant brings about a return to the wild type phenotype. In principle, forward-mutation assays should respond to a broad spectrum of mutagens because any mutation that interferes with gene expression should confer the detectable phenotype. A reversion assay might be expected to have a more restricted mutational response because only mutations that correct or compensate for the specific mutation in a particular mutant will be detected. In fact, some reversion assays respond to a broader spectrum of mutational changes than one might expect because mutations at a site other than that of the original mutation, either within the test gene or in a different gene (ie, a suppressor mutation), can sometimes confer the selected phenotype. Both forward mutation assays and reversion assays are used extensively in genetic toxicology.
The simplest gene mutation assays rely on selection techniques to detect mutations. A selection technique is a means of imposing experimental conditions under which only cells or organisms that have undergone mutation can grow. For example, only cells that have a mutation in the tk gene can grow in medium containing the inhibitory chemical trifluorothymidine (Seifried et al., 2006; Wang et al., 2009). Selection techniques greatly facilitate the identification of rare cells that have experienced mutation among the many cells that have not. Forward mutations (Clements, 2000; Vlasakova et al., 2005) and reversions (Josephy, 2000; Mortelmans and Riccio, 2000; Mortelmans and Zeiger, 2000; Kamber et al., 2009) can both be detected by selection techniques in microorganisms and cultured mammalian cells. Because of their speed, low cost, and ease of detecting events that occur at low frequency (ie, mutation), assays in microorganisms and cell cultures have figured prominently in genetic toxicology.
Studying mutagenesis in intact animals requires assays of more complex design than the simple selection methods used in microorganisms and cultured cells. Genetic toxicology assays therefore range from inexpensive short-term tests (Zeiger, 2010) that can be performed in a few days to complicated assays for mutations in mammalian germ cells (Favor, 1999; Russell, 2004; Singer et al., 2006). Even in multicellular organisms, there has been an emphasis on designing assays that detect mutations with great efficiency (Vogel et al., 1999; Casciano et al., 1999; Lambert et al., 2005; Dobrovolsky et al., 2010). Nevertheless, there remains a gradation in which an increase in relevance for human risk entails more elaborate and costly tests (Table 9-2). The most expensive mammalian tests are typically reserved for agents of special importance in basic research or risk assessment, whereas the simpler assays can be applied more broadly.
Cytogenetic assays differ in design from typical gene mutation assays because of their reliance on cytological rather than genetic methods. The goal in cytogenetic methods is the unequivocal visual recognition of cells that have experienced genetic damage. The alterations measured include chromosome aberrations (Preston et al., 1981; Ishidate et al., 1988; Corvi et al., 2008), micronuclei (Hayashi et al., 2007; Corvi et al., 2008; Fenech et al., 2011a,b), SCEs (Tucker et al., 1993a; Wilson and Thompson, 2007), and changes in chromosome number (Aardema et al., 1998; Paccierotti and Sgura, 2008). The latter include ploidy changes (eg, polyploid cells) and aneuploidy, in which one or a few chromosomes have been gained (ie, hyperploidy) or lost (ie, hypoploidy) relative to the normal chromosome number. Aneuploidy is of great interest because of its implications for human health, but assays for aneuploidy are not yet as refined or systematically applied as those for other classes of chromosomal alterations.
In all mutagenicity testing, one must be aware of possible sources of error. Factors to consider in the application of mutagenicity assays are the choice of suitable organisms and growth conditions, appropriate monitoring of genotypes and phenotypes, effective experimental design and treatment conditions, inclusion of proper positive and negative controls, and sound methods of data analysis. The articles cited in Table 9-2 discuss these aspects of genetic toxicology testing.
Many compounds that are not themselves mutagenic or carcinogenic can be activated into mutagens and carcinogens by metabolism (Guenngerich, 2001; Malling, 2004). Such compounds are called promutagens and procarcinogens. Because microorganisms and mammalian cell cultures lack many of the metabolic capabilities of intact mammals, provision must be made for metabolic activation in order to detect promutagens in many genetic assays. Incorporating an in vitro metabolic activation system derived from a mammalian tissue homogenate is the most common means of adding metabolic activation to microbial or cell culture assays (Malling and Frantz, 1973; Ames et al., 1975; Clements, 2000; Malling, 2004). For example, the promutagens dimethylnitrosamine and benzo[a]pyrene are not themselves mutagenic in bacteria, but they are mutagenic in bacterial assays if the bacteria are treated with the promutagen in the presence of a homogenate from mammalian liver.
The most widely used metabolic activation system in microbial and cell culture assays is a postmitochondrial supernatant from a rat liver homogenate, along with appropriate buffers and cofactors (Maron and Ames, 1983; Kirkland et al., 1990). The standard liver metabolic activation system is called an S9 mixture, designating a supernatant from centrifugation at 9000g (Malling and Frantz, 1973; Maron and Ames, 1983). Most of the short-term assays in Table 9-2 require exogenous metabolic activation to detect promutagens. Exceptions are assays in intact mammals and a few simpler assays that have a high level of endogenous cytochrome P450 metabolism, such as the detection of UDS or DNA strand breakage in cultured hepatocytes (Madle et al., 1994; Gealy et al., 2007).
Rat liver S9 provides a broad assemblage of metabolic reactions, but they are not necessarily the same as those of hepatic metabolism in an intact rat. Metabolic activation systems based on homogenates from mice, guinea pigs, hamsters, or monkeys and preparations from organs other than liver have found some use in mutagenicity testing (Mortelmans and Zeiger, 2000). In some cases, these systems detect mutagenicity more efficiently than rat liver S9. However, like a homogenate from rat liver, these systems may differ from the species or organs of their origin. Therefore, alternative metabolic activation systems tend to be more useful if chosen for mechanistic reasons rather than simply testing another species or organ. Such systems include the use of intact hepatocytes (Madle et al., 1994; Storer et al., 1996; Gealy et al., 2007) to preserve the cellular compartmentalization of reactions; an in vitro system that can carry out the reduction reactions needed to detect the activity of substances whose mutagenicity requires reductive metabolism, such as some azo compounds (Mortelmans and Zeiger, 2000; Seifried et al., 2006); and the use of mammalian cells or bacteria engineered to express foreign genes that encode enzymes of metabolic activation (Sawada and Kamataki, 1998; Crespi and Miller, 1999; Josephy, 2000, 2002; Oda et al., 2009).
Besides metabolic activation, some chemicals are subject to photochemical activation. The genotoxicity of such chemicals depends on the chemical being irradiated with ultraviolet or visible light. Many of the assays listed in Table 9-2, including gene-mutation assays in bacteria and cultured mammalian cells, cytogenetic assays, and the comet assay, have been adapted so that they can measure photogenotoxic effects (Brendler-Schwaab et al., 2004; Lynch et al., 2011).
Genes encoding enzymes of xenobiotic metabolism have been incorporated by recombinant DNA technology into microorganisms or cell cultures to expand their capacity for metabolic activation. The genes incorporated into bacteria may be derived from other bacteria or from humans (Josephy, 2002; Oda et al., 2009). For example, bacterial genes that cause overexpression of N-acetyltranserase enhance the sensitivity of the bacteria to the mutagenicity of aromatic amines or nitroarenes (Josephy, 2002). The expression of human cytochrome P450 enzymes in Salmonella tester strains from the Ames assay (Josephy 2002; Yamazaki et al., 2004), a Salmonella SOS-induction assay (Oda et al., 2009), or E coli strains of the lacZ reversion assay (Josephy, 2000) permits the activation of such promutagens as 2-aminoanthracene and 2-aminofluorene without an S9 mixture. Mammalian cell lines have also been genetically engineered to express human Phase-I and Phase-II enzymes, including those catalyzing reactions of metabolic activation (Sawada and Kamataki, 1998). Many cell lines stably expressing a single form of P450 have been established. Mutagenesis can be measured through such endpoints as HPRT mutations and cytogenetic alterations, and the cells are well suited to analyzing the contribution of different enzymes to the activation of promutagens (Crespi and Miller, 1999).
Metabolic activation is so central to genetic toxicology that all mutagenicity testing programs must provide for it in the choice of assays and procedures. Despite their usefulness, in vitro metabolic activation systems, however well refined, cannot mimic mammalian metabolism perfectly. There are differences among tissues in reactions that activate or inactivate foreign compounds, and organisms of the normal flora of the gut can contribute to metabolism in intact mammals. Agents that induce enzyme systems or otherwise alter the physiological state can also modify the metabolism of toxicants, and the balance between activation and detoxication reactions in vitro may differ from that in vivo.
Structural Alerts and In Silico Assays
The first indication that a chemical is a mutagen often lies in chemical structure. Potential electrophilic sites in a molecule serve as an alert to possible mutagenicity and carcinogenicity because such sites confer reactivity with nucleophilic sites in DNA (Tennant and Ashby, 1991). Structural alerts in combination with critical interpretation are a valuable adjunct to mutagenicity testing (Tennant and Ashby, 1991; Ashby and Paton, 1993). Attempts to formalize the structural prediction through automated computer programs have not yet led to an ability to predict mutagenicity and carcinogenicity of new chemicals with great accuracy (Snyder et al., 2004), but promising developmental work on such systems continues (Votano et al., 2004; Snyder and Smith, 2005).
Computer-based systems for predicting genotoxicity based on chemical properties are sometimes called in silico assays. These assays include computational and structural programs (Knudsen et al., 2011; Snyder and Smith, 2005; Snyder, 2009; Mahadevan et al., 2011) and the modeling of quantitative structure–activity relationships (Votano et al., 2004; Snyder and Smith, 2005; Mahadevan et al., 2011). Although there is much skepticism that such approaches can replace biological testing, they hold promise of improving the efficiency of testing strategies and reducing current levels of animal use (Guha, 2011).
DNA Damage and Repair Assays
Some assays measure DNA damage itself, rather than mutational consequences of DNA damage. They may do so directly, through such indicators as chemical adducts or strand breaks in DNA, or indirectly, through the measurement of biological repair processes. Adducts in DNA are detected by 32P-postlabeling, high-performance liquid chromatography (HPLC), fluorescence-based methods, mass spectrometry, immunological methods using antibodies against specific adducts, isotope-labeled DNA binding, and electrochemical detection (Phillips et al., 2000; Farmer and Singh, 2008; Himmelstein et al., 2009). The 32P-postlabeling method is highly versatile, in that it is sensitive and can be applied to diverse mutagens, but it may fall short of other methods for quantitative accuracy (Farmer and Singh, 2008). Through a combination of methods, many classes of adducts, including those of such environmentally widespread compounds as polynuclear aromatic hydrocarbons, can be detected. The measurement of adducts after human chemical exposures has proven useful in human monitoring, molecular dosimetry, and risk assessment Phillips et al., 2000; Farmer and Singh, 2008; Himmelstein et al., 2009). Adducts in somatic cells are relevant to carcinogenesis (Himmelstein et al., 2009), whereas those in reproductive cells permit molecular dosimetry for germ cell mutagenesis (Olsen et al., 2010; 2011; Verhofsrad et al., 2011).
DNA strand breakage can be measured by alkaline elution (Storer et al., 1996; Gealy et al., 2007) and electrophoretic methods. Single-cell gel electrophoresis, also called the comet assay, is a widely used, rapid method of measuring DNA damage (Fairbairn et al., 1995; Singh, 2000; Tice et al., 2000; Collins, 2004; Olive, 2009). In this assay cells are incorporated into agarose on slides, lysed so as to liberate their DNA, and subjected to electrophoresis. The DNA is stained with a fluorescent dye for observation and image analysis. Because broken DNA fragments migrate more quickly than larger pieces of DNA, a blur of fragments (a “comet”) is observed when the DNA is extensively damaged. The extent of DNA damage can be estimated from the length and other attributes of the comet tail (Collins, 2004). Variations in the procedure permit the general detection of DNA strand breakage under alkaline conditions (Fairbairn et al., 1995; Singh, 2000; Tice et al., 2000) or the preferential detection of double-strand breaks under neutral conditions (Fairbairn et al., 1995; Olive, 2009). A recent development is the combination of the comet assay with FISH to detect damage in specific regions of the genome (Glei et al., 2009; Shaposhnikov et al., 2009). Although the comet assay is relatively new, it has proven to be a sensitive indicator of DNA damage with broad applicability. It has been used most commonly with human lymphocytes (Fairbairn et al., 1995; Singh, 2000; Collins, 2004) and other mammalian cells (Tice et al., 2000), but it can be adapted to diverse species, including plants, worms, mollusks, fish, and amphibians (Cotelle and Férard, 1999; Lee and Steinert, 2003; Jha, 2004). This adaptability makes it well suited to use in environmental genetic toxicology. The applicability of the comet assay and other DNA damage assays to rodent testes (Bentley et al., 1994; Speit et al., 2009) makes these methods helpful in interpreting risks to germ cells.
The occurrence of DNA repair can serve as an easily measured indicator of DNA damage. Repair assays have been developed in microorganisms, cultured mammalian cells, and intact mammals (Table 9-2). Greater toxicity of a chemical in DNA-repair-deficient strains than in their repair-proficient counterparts (eg, rec+ and rec− in Bacillus subtilis) can serve as an indicator of DNA damage in bacteria (Takigami et al., 2002). Bacterial repair assays find occasional application but are used less commonly today than historically. The measurement of UDS, which is a measure of excision repair, is a mammalian DNA repair assay. The occurrence of UDS indicates that DNA has been damaged (Madle et al., 1994). The absence of UDS, however, does not provide evidence that DNA has not been damaged because some classes of damage are not readily excised, and some excisable damage may not be detected as a consequence of assay insensitivity (Kirkland and Speit, 2008). Despite these limitations, UDS assays continue to find some use because of their applicability to cultured hepatocytes with endogenous cytochrome P450 enzyme activities and to tissues of intact animals, including hepatocytes (Madle et al., 1994) and germinal tissue (Bentley et al., 1994; Sotomayor and Sega, 2000).
Besides specific DNA repair processes, the induction of general responses to genotoxic stress has been used as an indicator of genetic damage. The induction of SOS functions, indicated by phage induction or by colorimetry, can serve as an indicator of genetic damage in bacteria (Quillardet and Hofnung, 1993; Yasunaga et al., 2004; Oda et al., 2009). The GADD45a-GFP assay, also called “Green Screen,” detects genotoxic stress in human lymphoblastoid TK6 cells (Hastwell et al., 2009). The stress response is detected by a green-fluorescent protein reporter in the genetic construct, and the induction of fluorescence has been observed with mutagens, clastogens, and aneugens (Hastwell et al., 2009). The assay can be conducted with S9 metabolic activation and lends itself to automated detection by flow cytometry (Jagger et al., 2009).
Gene Mutations in Prokaryotes
The most common means of detecting mutations in microorganisms is selecting for reversion in strains that have a specific nutritional requirement differing from wild type members of the species; such strains are called auxotrophs. For example, the widely used assay developed by Bruce Ames and his colleagues is based on measuring reversion in several histidine auxotrophs in Salmonella enterica serovar Typhimurium, commonly called S typhimurium (Ames et al., 1975).
In the Ames assay one measures the frequency of histidine-independent bacteria that arise in a histidine-requiring strain in the presence or absence of the chemical being tested. Auxotrophic bacteria are treated with the chemical of interest by one of several procedures (eg, the standard plate-incorporation assay) and plated on medium that is deficient in histidine (Ames et al., 1975; Maron and Ames, 1983; Mortelmans and Zeiger, 2000). The assay is conducted using genetically different strains so that reversion by base pair substitutions and frameshift mutations in several DNA sequence contexts can be detected and distinguished. Because Salmonella does not metabolize promutagens in the same way as mammalian tissues, the assay is generally performed in the presence and absence of a rat liver S9 metabolic activation system. Hence, the Ames assay is also called the Salmonella/microsome assay.
The principal strains of the Ames test and their characteristics are summarized in Table 9-3. In addition to the histidine alleles that provide the target for measuring mutagenesis, the Ames tester strains contain other genes and plasmids that enhance the assay. Part I of the table gives genotypes, and Part II explains the rationale for including specific genetic characteristics in the strains. Part III summarizes the principal DNA target in each strain and the mechanisms of reversion. Taken together, the Ames strains detect a broad array of mutations, and they complement one another. For example, strains TA102 and TA104, which are sensitive to agents that cause oxidative damage in DNA, detect the A:T → G:C base pair substitutions that are not detected by hisG46 strains (Mortelmans and Zeiger, 2000). TA102 also detects agents that cause DNA cross-links because it has an intact excision repair system whereas the other common tester strains do not.
Table 9-3The Ames Assay: Tester Strains and Their Characteristics ||Download (.pdf) Table 9-3 The Ames Assay: Tester Strains and Their Characteristics
|I. STANDARD TESTER STRAINS OF SALMONELLA TYPHIMURIUM |
|STRAIN ||TARGET ALLELE ||CHROMOSOMAL GENOTYPE ||PLASMIDS |
|TA1535 ||hisG46 ||hisG46 rfa ΔuvrB ||None |
|TA100 ||hisG46 ||hisG46 rfa ΔuvrB ||pKM101 (mucAB Apr) |
|TA1538 ||hisD3052 ||hisD3052 rfa ΔuvrB ||None |
|TA98 ||hisD3052 ||hisD3052 rfa ΔuvrB ||pKM101 (mucAB Apr) |
|TA1537 ||hisC3076 ||hisC3076 rfa ΔuvrB ||None |
|TA97 ||hisD6610 ||hisD6610 hisO1242 rf ΔuvrB ||pKM101 (mucAB Apr) |
|TA102 ||hisG428 ||hisΔ (G)8476 rfa ||pKM101 (mucAB Apr); pAQ1 (hisG428 Tcr) |
|TA104 ||hisG428 ||hisG428 rfa ΔuvrB ||pKM101 (mucAB Apr) |
|II. GENETIC CHARACTERISTICS OF THE AMES TESTER STRAINS |
|CHARACTERISTIC ||RATIONALE FOR INCLUSION IN THE TESTER STRAIN |
|rfa ||Alters the lipopolysaccharide wall, enhancing permeability to mutagens. |
|ΔuvrB ||Deletes the excision repair system, increasing sensitivity to many mutagens; retention of excision in TA102 permits the detection of DNA cross-linking agents. |
|mucAB ||Enhances sensitivity to some mutagens whose activity depends on the SOS system. |
|Apr ||Permits selection for the presence of pKM101 by ampicillin resistance. |
|hisO1242 ||Affects regulation of histidine genes, enhancing revertibility of hisD6610 in TA97. |
|His Δ (G)8476 ||Eliminates the chromosomal hisG gene, allowing detection of the reversion of hisG428 on pAQ1 in TA102. |
|Tcr ||Permits selection for the presence of pAQ1 in TA102 by tetracycline resistance. |
| ||III. MECHANISMS OF REVERSION DETECTED BY THE AMES TESTER STRAINS |
|STRAIN ||PRIMARY TARGET* ||MUTATIONS DETECTED |
|TA1535, TA100 ||GGG/CCC ||Base-pair substitutions, principally those beginning at G:C base pairs (G:C→A:T; G:C→T:A; G:C→C:G). These strains also detect A:T→C:G but not A:T→G:C. |
|TA1538, TA98 ||CGCGCGCG/GCGCGCGC ||Frameshift mutations, especially −2 frameshifts (–GC or –CG), +1 frameshifts, other small deletions, and some complex mutations. |
|TA1537 ||GGGGG/CCCCC ||Frameshift mutations, mainly –1 (–G or –C; less frequently –T), but also some +CG frameshifts. |
|TA97 ||GGGGGG/CCCCCC ||Frameshift mutations, combining the specificity of TA1537 at the primary target and with some characteristics of TA98. |
|TA102, TA104 ||TAA/ATT ||Base-pair substitutions, principally those beginning at A:T base pairs (A:T→G:C; A:T→T:A; A:T→C:G), but also G:C→T:A and G:C→A:T. |
The most common version of the Ames assay is the plate-incorporation test (Ames et al., 1975; Maron and Ames, 1983; Mortelmans and Zeiger, 2000). In this procedure, the bacterial tester strain, the test compound (or solvent control), and the S9 metabolic activation system (or buffer for samples without S9) are added to 2 mL of molten agar containing biotin and a trace of histidine to allow a few cell divisions, mixed, and immediately poured onto the surface of a petri dish selective for histidine-independent revertants. For general testing it is recommended that at least three plates per dose and five doses be used with and without S9, along with appropriate concurrent positive and negative controls (Mortelmans and Zeiger, 2000). Variations on the standard plate-incorporation assay confer advantages for some applications. These include a preincubation assay that facilitates the detection of unstable compounds and short-lived metabolites, a desiccator assay for testing volatile chemicals and gases, a microsuspension assay for working with small quantities of test agent, assays incorporating reductive metabolism rather than the conventional S9 system, and assays under hypoxic conditions (Mortelmans and Zeiger, 2000).
Although simplicity is a great merit of microbial assays, it can also be deceptive. Even assays that are simple in design and application can be performed incorrectly. For example, in the Ames assay one may see very small colonies in the petri dishes at highly toxic doses (Maron and Ames, 1983; Kirkland et al., 1990; Mortelmans and Zeiger, 2000). Counting such colonies as revertants would be an error because they may actually be nonrevertant survivors that grew on the low concentration of histidine in the plates. Were there millions of survivors, the amount of histidine would have been insufficient to allow any of them (except real revertants) to form colonies. This artifact is easily avoided by checking that there is a faint lawn of bacterial growth in the plates; one can also confirm that colonies are revertants by streaking them on medium without histidine to be sure that they grow in its absence. Such pitfalls exist in all mutagenicity tests. Therefore, anyone performing mutagenicity tests must have detailed familiarity with the laboratory application and literature of the assay and be observant about the responsiveness of the assay.
Although information from the Ames assay has become a standard in genetic toxicology testing, equivalent information can be obtained from other bacterial assays. Like the Ames assay, the WP2 tryptophan reversion assay in E coli (Kirkland et al., 1990; Mortelmans and Riccio, 2000) incorporates genetic features that enhance assay sensitivity, can accommodate S9 metabolic activation, and performs well in many laboratories. Mutations are detected by selecting for reversion of a trpE allele from Trp− to Trp+. Its responsiveness to mutagens most closely resembles TA102 among the Ames strains (Mortelmans and Riccio, 2000).
Bacterial reversion assays are commonly used for testing purposes, but they also provide information on molecular mechanisms of mutagenesis. The broader understanding of mutational mechanisms that comes from refined genetic assays and molecular analysis of mutations can contribute to the interpretation of mutational hazards. The primary reversion mechanisms of the Ames strains, summarized in Table 9-3, were initially determined by genetic and biochemical means (Maron and Ames, 1983). An ingenious method called allele-specific colony hybridization greatly facilitated the molecular analysis of revertants in the Ames assay (Koch et al., 1994), and many spontaneous and induced revertants have been cloned or amplified by the polymerase chain reaction (PCR) and sequenced (Levine et al., 1994; DeMarini, 2000).
Part III of Table 9-3 is by necessity a simplification with respect both to targets and mechanisms of reversion of the Ames strains. Some mutations that bring about reversion to histidine independence fall outside the primary target, and the full target has been found to be as much as 76 base pairs in hisD3052 (DeMarini et al., 1998). Other revertants can arise by suppressor mutations in other genes. It has been shown that hisG46, hisG428, hisC3076, hisD6610, and hisD3052 all revert by multiple mechanisms and that the spectrum of classes of revertants may vary depending on the mutagen, experimental conditions, and other elements of the genotype (Cebula and Koch, 1990; Prival and Cebula, 1992; DeMarini et al., 1998; Mortelmans and Zeiger, 2000).
The development of Salmonella strains that are highly specific with respect to mechanisms of reversion has made the identification of particular base pair substitutions more straightforward. These strains (TA7001–TA7006) each revert from his− to his+ by a single kind of mutation (eg, G:C to T:A), and collectively they permit the specific detection of all six possible base pair substitutions (Gee et al., 1994; Mortelmans and Zeiger, 2000; Kamber et al., 2009). The assay is usually conducted using a fluctuation test in 24-well plates, rather than the plate-incorporation or preincubation method. The ability to discern mutagens and nonmutagens is comparable to the standard Ames assay (Kamber et al., 2009).
Specific reversion assays are also available in E coli. A versatile system based on reversion of lacZ mutations in E coli permits the specific detection of all six possible base pair substitutions (Cupples and Miller, 1989; Josephy, 2000) and frameshift mutations for which one or two bases have been added or deleted in various sequence contexts (Cupples et al., 1990; Josephy, 2000; Hoffmann et al., 2003). The versatility of the lacZ assay has been expanded through the introduction of useful characteristics into the strains parallel to those incorporated into the Ames strains. Among the features added to the lacZ assay are DNA repair deficiencies, permeability alterations, plasmid-enhanced mutagenesis, and enzymes of mutagen metabolism (Josephy, 2000).
Bacterial forward mutation assays, such as selections for resistance to arabinose or to purine or pyrimidine analogs in Salmonella (Jurado et al., 1994; Vlasakova et al., 2005), are also used in research and testing, although less extensively than reversion assays. A versatile forward mutation assay that has contributed greatly to an understanding of mechanisms of mutagenesis is the lacI system in E coli (Calos and Miller, 1981; Halliday and Glickman, 1991). Mutations in the lacI gene, which encodes the repressor of the lactose operon, are easily identified by phenotype, cloned or amplified by PCR, and sequenced. The lacI gene is widely used as a target for mutagenesis both in E coli and in transgenic mice, and thousands of lacI mutants have been sequenced.
Genetic Alterations in Nonmammalian Eukaryotes
Gene Mutations and Chromosome Aberrations
Many early studies of mutagenesis used yeasts, mycelial fungi, plants, and insects as experimental organisms. Even though well-characterized genetic systems permit the analysis of a diverse array of genetic alterations in these organisms (Table 9-2), they have been largely supplanted in genetic toxicology by bacterial and mammalian systems. Exceptions are to be found where the assays in nonmammalian eukaryotes permit the study of genetic endpoints that are not readily analyzed in mammals or where the organism has special attributes that fit a particular application.
The fruit fly, Drosophila, has long occupied a prominent place in genetic research. In fact, the first unequivocal evidence of chemical mutagenesis was obtained in Scotland in 1941 when Charlotte Auerbach and J.M. Robson demonstrated that mustard gas is mutagenic in Drosophila. Drosophila continues to be used in modern mutation research (Potter et al., 2000) but its role in genetic toxicology is now more limited. The Drosophila assay of greatest historical importance is the sex-linked recessive lethal (SLRL) test. A strength of the SLRL test is that it permits the detection of recessive lethal mutations at 600 to 800 different loci on the X chromosome by screening for the presence or absence of wild type males in the offspring of specifically designed crosses (Mason et al., 1987; Vogel et al., 1999). The genetic alterations include gene mutations and small deletions. The spontaneous frequency of SLRL is about 0.2%, and a significant increase over this frequency in the lineages derived from treated males indicates mutagenesis. Although it requires screening large numbers of fruit fly vials, the SLRL test yields information about mutagenesis in germ cells, which is lacking in all microbial and cell culture systems. However, means of exposure, measurement of doses, metabolism, and gametogenesis in insects differ from those in mammalian toxicology, thereby introducing doubt about the relevance of Drosophila assays to human genetic risk. Drosophila assays are also available for studying the induction of chromosome abnormalities in germ cells, specifically heritable translocations (Mason et al., 1987; Vogel et al., 1999).
Genetic and cytogenetic assays in plants (Ma et al., 2005; Grant and Owens, 2006; Misík et al., 2011) also occupy a more restricted niche in modern genetic toxicology than they did years ago. However, plant assays continue to find use in special applications, such as in situ monitoring for mutagens and exploration of the metabolism of promutagens by agricultural plants. In situ monitoring entails looking for evidence of mutagenesis in organisms that are grown in the environment of interest. Natural plant populations can also be examined for evidence of genetic damage, but doing so requires utmost precaution when characterizing the environments and defining appropriate control populations.
Assays in nonmammalian eukaryotes continue to be important in the study of induced recombination. Recombinogenic effects in yeast have long been used as a general indicator of genetic damage (Zimmermann et al., 1984), and interest in the induction of recombination has increased as recombinational events have been implicated in the etiology of cancer (Sengstag, 1994; Reliene et al., 2007). LOH is central to the expression of the mutant alleles of tumor-suppressor genes, and mitotic recombination is a major mechanism of LOH. Widely used assays for recombinogens detect mitotic crossing over and mitotic gene conversion in the yeast Saccharomyces cerevisiae (Zimmermann, 1992), and hundreds of chemicals have been tested for recombinogenic effects in straightforward yeast assays (Zimmermann et al., 1984). In yeast strain D7, for example, mitotic crossing over involving the ade2 locus is detected on the basis of pink and red colony color, mitotic gene conversion at the trp5 locus by selection for growth without tryptophan, and gene mutations by reversion of the ilv1-92 allele (Zimmermann, 1992; Freeman and Hoffmann, 2007). Newer yeast assays have been constructed to discern whether LOH has occurred by mitotic recombination or chromosome loss (Daigaku et al., 2004; Nunoshiba et al., 2007). Strategies have also been devised to detect recombinogenic effects in human lymphocytes (Turner et al., 2003), other mammalian cells, mice, plants, and mycelial fungi (Hoffmann, 1994; Reliene et al., 2007). At least 350 chemicals have been evaluated in Drosophila somatic cell assays in which recombinogenic effects are detected by examining wings or eyes for regions in which recessive alleles are expressed in heterozygotes (Vogel et al., 1999; Vogel and Nivard, 2000).
Gene Mutations in Mammals
Mutagenicity assays in cultured mammalian cells have some of the same advantages as microbial assays with respect to speed and cost, and they use similar approaches. The most widely used assays for gene mutations in mammalian cells detect forward mutations that confer resistance to a toxic chemical. For example, mutations in the gene encoding hypoxanthine-guanine phosphoribosyltransferase (HPRT enzyme; HPRT gene) confer resistance to the purine analogue 6-thioguanine (Li et al., 1988; Parry et al., 2005), and thymidine kinase mutations (TK enzyme; TK gene) confer resistance to the pyrimidine analogue trifluorothymidine (Clements, 2000; Wang et al., 2009). HPRT and TK mutations may therefore be detected by attempting to grow cells in the presence of purine analogues and pyrimidine analogues, respectively. For historical reasons, HPRT assays have most commonly been conducted in Chinese hamster cells or human cells, whereas TK assays have used mouse lymphoma cells or human cells. The mouse lymphoma assay, long used for detecting gene mutations, is now also used to detect other endpoints, including recombination, deletion, and aneuploidy (Wang et al., 2009). Forward-mutation assays typically respond to diverse mechanisms of mutagenesis, but there are exceptions such as resistance to ouabain, which only occurs by a specific alteration (DeMarini et al., 1989). Assays that do not detect various kinds of mutations are not useful for general mutagenicity testing. Genetic or molecular evidence that an assay is responsive to diverse mechanisms of mutagenesis is essential.
Instead of using selective media, flow cytometry can be used to detect gene mutations by immunological methods in mammalian cell cultures and intact animals. An in vitro assay for CD59 mutations is performed in CHO-human hybrid AL cells (Zhou et al., 2006). AL cells contain a single human chromosome 11 along with the Chinese hamster chromosome complement of the Chinese hamster ovary (CHO) cells. CD59 is a cell-surface protein encoded by a gene on the human chromosome. Fluorescent anti-CD59 antibody is used to quantify CD59 cells by flow cytometry. Besides CD59 itself, mutations in other CD genes on chromosome 11 and in their glycosylphosphatidylinositol (GPI) anchor can be detected in the assay (Ross et al., 2007). An assay for mutations in Pig-a, the GPI anchor gene, is discussed with in vivo assays.
In vivo assays involve treating intact animals and analyzing appropriate tissues for genetic effects. The choice of suitable doses, treatment procedures, controls, and sample sizes is critical in the conduct of in vivo tests. Mutations may be detected either in somatic cells or in germ cells. The latter are of special interest with respect to risk for future generations.
The mouse spot test is a traditional genetic assay for gene mutations in somatic cells (Styles and Penman, 1985; Lambert et al., 2005). Visible spots of altered phenotype in mice heterozygous for coat-colored genes indicate mutations in the progenitor cells of the altered regions. Although straightforward in design, the spot test is less used today than other somatic cell assays or than its germ cell counterpart, the mouse specific-locus test.
Cells that are amenable to positive selection for mutants when collected from intact animals form the basis for efficient in vivo mutation-detection assays analogous to those in mammalian cell cultures. Lymphocytes with mutations in the HPRT gene are readily detected by selection for resistance to 6-thioguanine. The hprt assay in mice, rats, and monkeys (Casciano et al., 1999) is of special interest because it permits comparisons to the measurement of HPRT mutations in humans in mutational monitoring (Cole and Skopek, 1994; Albertini and Hayes, 1997; Albertini, 2001).
The Pig-a assay is a newer assay that has versions suitable for human monitoring and laboratory studies. The assay detects mutations that block GPI synthesis (Dobrovolsky et al., 2010). Pig-a is a sex-linked gene whose gene product functions as an anchor for cell-surface proteins. Mutations in Pig-a can be detected in red blood cells from rats, mice, monkeys, and humans by means of fluorescent antibodies against GPI-anchored cell-surface proteins, such as CD59. Using antibodies to more than one GPI-linked marker has been suggested as a means of making the assay specific to Pig-a rather than also detecting mutants for a particular cell-surface protein. Frequencies can be measured by clonal growth of Pig-a cells using proaerolysin (ProAER) selection or by flow cytometry (Dobrovolsky et al., 2010; Miura et al., 2011; Dobo et al., 2011). The fact that the same assay can be performed in several species makes this a promising assay for comparisons of human monitoring and controlled exposures in laboratory animals.
Besides determining whether agents are mutagenic, mutation assays provide information on mechanisms of mutagenesis that contributes to an understanding of mutational hazards. Base pair substitutions and large deletions, which may be indistinguishable on the basis of phenotype, can be differentiated through the use of probes for the target gene and Southern blotting, in that base substitutions are too subtle to be detectable on the blots, whereas gross structural alterations are visible (Cole and Skopek, 1994; Albertini and Hayes, 1997). Molecular analysis has been used to determine proportions of mutations ascribable to deletions and other structural alterations in several assays, including the specific-locus test for germ cell mutations in mice (Favor, 1999) and the human HPRT assay (Cole and Skopek, 1994). Gene mutations have been characterized at the molecular level by DNA sequence analysis both in transgenic rodents (Lambert et al., 2005; Singer et al., 2006) and in endogenous mammalian genes (Cariello and Skopek, 1993). Many HPRT mutations from human cells in vitro and in vivo have been analyzed at the molecular level and classified with respect to base pair substitutions, frameshifts, small deletions, large deletions, and other alterations (Cole and Skopek, 1994).
Transgenic animals are products of DNA technology in which the animal contains foreign DNA sequences that have been added to the genome. The foreign DNA is represented in all the somatic cells of the animal and is transmitted in the germ line to progeny. Mutagenicity assays in transgenic animals combine in vivo metabolic activation and pharmacodynamics with simple microbial detection systems, and they permit the analysis of mutations induced in diverse mammalian tissues (Lambert et al., 2005; Sykes et al., 2006; Valentine et al., 2010).
The transgenic animals that have figured most heavily in genetic toxicology are rodents that carry lac genes from E coli. The bacterial genes were introduced into mice or rats by injecting a vector carrying the genes into fertilized oocytes (Lambert et al., 2005). The strains are commonly referred to by their commercial names—the “Big Blue mouse” and “Big Blue rat,” which use lacI as a target for mutagenesis, and the “Muta Mouse,” which uses lacZ (Lambert et al., 2005). After mutagenic treatment of the transgenic animals, the lac genes are recovered from the animal, packaged in phageλ, and transferred to E coli for mutational analysis. Mutant plaques are identified on the basis of phenotype, and mutant frequencies can be calculated for different tissues of the treated animals. The cII locus may be used as a second target gene in both the lacZ and lacI transgenic assays (Swiger, 2001; Lambert et al., 2005). Its use offers technical advantages as a small, easily sequenced target in which mutations are detected by positive selection, and it permits interesting comparisons both within and between assays.
A lacZ transgenic mouse that uses a plasmid-based system rather than a phage vector has the advantage that deletion mutants are more readily recovered than in the phage-based lac systems (Lambert et al., 2005). Deletions may also be detected in the gpt delta mouse and rat using a phage vector system. These transgenic animals detect two kinds of genetic events in two targets—point mutations in gpt detected by resistance to 6-thioguanine and spi deletions that permit growth on P2 lysogens (Okada et al., 1999; Lambert et al., 2005). Other transgenic assays are under development and offer the prospect of expanding the versatility of such assays (Lambert et al., 2005). These include pKZ1 mice in which inversions and deletions arising in various tissues by intrachromosomal recombination have been used to study effects of low doses of ionizing radiation (Sykes et al., 2006) and an assay that detects both forward mutations and reversion in mice that carry the genome of phage ΦX174 (Valentine et al., 2010).
Various mutagens, including alkylating agents, nitrosamines, procarbazine, cyclophosphamide, and polycyclic aromatic hydrocarbons have been studied in transgenic mouse assays, and mutant frequencies have been analyzed in such diverse tissues as liver, skin, spleen, kidney, bladder, small intestine, bone marrow, and testis (Lambert et al., 2005). Mutation frequencies in transgenes in testes have been compared to results in standard germ cell mutagenesis assays (Singer et al., 2006). Female germ cells are less amenable to study in transgenic assays because of the difficulty of collecting sufficient numbers of oocytes, but it has been suggested that granulosa cells from ovarian follicles may serve as a surrogate for the exposure of female germ cells to mutagens (Singer et al., 2006). Mutant frequencies have been compared to the formation of adducts in various tissues and to the site specificity of carcinogenesis and DNA repair capacity (Lambert et al., 2005). An important issue that remains to be resolved is the extent to which transgenes resemble endogenous genes. Although their mutational responses tend to be comparable (Lambert et al., 2005), some differences have been noted (Burkhart and Malling, 1993; Lambert et al., 2005), and questions have been raised about the relevance of mutations that might be recovered from dying or dead animal tissues (Burkhart and Malling, 1994). Therefore, transgenic animals offer promising models for the study of chemical mutagenesis, but they must be further characterized before their ultimate place in hazard assessment is clear.
Mammalian Cytogenetic Assays
Cytogenetic assays rely on the use of microscopy for the direct observation of the effects of interest. This approach differs sharply from the indirectness of traditional genetic assays in which one observes a phenotype and reaches conclusions about genes. It is only through the addition of DNA sequencing that genetic assays can approach the directness of cytogenetic assays.
In conventional cytogenetics, metaphase analysis is used to detect chromosomal anomalies, especially unstable chromosome and chromatid aberrations. A key factor in the design of cytogenetic assays is obtaining appropriate cell populations for treatment and analysis (Preston et al., 1981; Ishidate et al., 1988; Kirkland et al., 1990; Galloway et al., 1994). Cells with a stable, well-defined karyotype, short generation time, low chromosome number, and large chromosomes are ideal for cytogenetic analysis. For this reason, Chinese hamster cells have been used widely in cytogenetic testing. Other cells are also suitable, and human cells, especially peripheral lymphocytes, have been used extensively. Cells should be treated during a sensitive period of the cell cycle (typically S), and aberrations should be analyzed at the first mitotic division after treatment so that the sensitivity of the assay is not reduced by unstable aberrations being lost during cell division. Examples of chromosome aberrations are shown in Fig. 9-4.
Chromosome aberrations induced by x-rays in Chinese hamster ovary (CHO) cells. (A) A chromatid deletion (►). (B) A chromatid exchange called a triradial (►). (C) A small interstitial deletion (►) that resulted from chromosome breakage. (D) A metaphase with more than one aberration: a centric ring plus an acentric fragment (►) and a dicentric chromosome plus an acentric fragment (→).
Cytogenetic assays require careful attention to growth conditions, controls, doses, treatment conditions, and time intervals between treatment and the sampling of cells for analysis (Preston et al., 1981; Ishidate et al., 1988; Kirkland et al., 1990; Galloway et al., 2011). Data collection is a critical part of cytogenetic analysis. It is essential that sufficient cells be analyzed because a negative result in a small sample is inconclusive. Results should be recorded for specific classes of aberrations, not just an overall index of aberrations per cell. The need for detailed data is all the more important because of nonuniformity in the classification of aberrations and disagreement on whether small achromatic (ie, unstained) gaps in chromosomes are true chromosomal aberrations. Gaps should be quantified but not pooled with other aberrations.
In interpreting results on the induction of chromosome aberrations in cell cultures, one must be alert to the possibility of artifacts associated with extreme assay conditions because aberrations induced under such circumstances may not be a reflection of a chemical-specific genotoxicity (Scott et al., 1991; Galloway, 2000; Galloway et al., 2011). Questionable positive results have been found at highly cytotoxic doses (Galloway et al., 2011), high osmolality, and pH extremes (Scott et al., 1991). The possibility that metabolic activation systems may be genotoxic also warrants scrutiny (Scott et al., 1991). Although excessively high doses may lead to artifactual positive responses, the failure to test to a sufficiently high dose also undermines the utility of a test. Therefore, testing should be extended to a dose at which there is some cytotoxicity, such as a reduction in a replicative index or the mitotic index (the proportion of cells in division). If the chemical is nontoxic, testing dosages should extend up to an arbitrary limit of dosage (Galloway et al., 2011). By a consensus of cytogeneticists and genetic toxicologists, a limit of 10 mM or 5 mg/mL, whichever is lower, has been recommended (Galloway et al., 2011). Some have argued that the limit should be lowered, perhaps to 1 mM, but no consensus could be reached on this point (Galloway et al., 2011).
In vivo assays for chromosome aberrations involve treating intact animals and later collecting cells for cytogenetic analysis (Preston et al., 1981; Kirkland et al., 1990; Tice et al., 1994). The main advantage of in vivo assays is that they include mammalian metabolism, DNA repair, and pharmacodynamics. The target is typically a tissue from which large numbers of dividing cells are easily prepared for analysis. Bone marrow from rats, mice, or Chinese hamsters is most commonly used. Peripheral lymphocytes are another suitable target when stimulated to divide with a mitogen such as phytohemagglutinin. Effective testing requires dosages and routes of administration that ensure adequate exposure of the target cells, proper intervals between treatment and collecting cells, and sufficient numbers of animals and cells analyzed (Preston et al., 1981; Kirkland et al., 1990; Tice et al., 1994).
An important development in cytogenetic analysis is FISH, in which a nucleic acid probe is hybridized to complementary sequences in chromosomal DNA (Tucker et al., 1993b, 2005; Paccierotti and Sgura, 2008). The probe is labeled with a fluorescent dye so that the chromosomal location to which it binds is visible by fluorescence microscopy. Composite probes have been developed from sequences unique to specific human chromosomes, giving a uniform fluorescent label over the entire chromosome. Slides prepared for standard metaphase analysis are suitable for FISH after they have undergone a simple denaturation procedure. The use of whole-chromosome probes is commonly called “chromosome painting” (Tucker et al., 1993b; Speicher and Carter, 2005; Paccierotti and Sgura, 2008). Another significant advantage of FISH methods is that the probes can be used with interphase cells/chromosomes making any tissue potentially available for analysis (Vorsanova et al., 2010). Examples of cells showing chromosome painting and reciprocal translocations are shown in Figs. 9-5 and 9-6.
Karyotype of human cell using FISH. To produce a FISH-based karyotype of the type depicted here, appropriate chromosome probe sets are developed for each chromosome pair such that by using computer colorization each has a specific color for analysis. (Reproduced with permission from Schrock E, et al. Multicolor spectral karyotyping of human chromosomes. Science. 1996 Jul 26;273(5274):494–497.)
Chromosome aberrations identified by FISH. Human breast cancer cell with aneuploidy for some chromosomes and with reciprocal translocations (identified by color switches along a chromosome).
Chromosome painting facilitates cytogenetic analysis because aberrations are easily detected by the number of fluorescent regions in a painted metaphase. For example, if chromosome 4 were painted with a probe while the other chromosomes were counter-stained in a different color, one would see only the two homologues of chromosome 4 in the color of the probe in a normal cell. However, if there were a translocation or a dicentric chromosome and fragment involving chromosome 4, one would see three areas of fluorescence—one normal chromosome 4 and the two pieces involved in the chromosome rearrangement. Aberrations are detected only in the painted portion of the genome, but this disadvantage can be offset by painting a few chromosomes simultaneously with probes of different colors (Tucker et al., 1993b). FISH reduces the time and technical skill required to detect chromosome aberrations, and it permits the scoring of stable aberrations, such as translocations and insertions, that are not readily detected in traditional metaphase analysis without special labeling techniques. Using FISH, some chromosomal analysis can even be conducted in interphase cells (Paccierotti and Sgura, 2008; Vorsanova et al., 2010). Although FISH is not routinely used in genotoxicity testing, it is a valuable research tool for studying clastogens and is having a substantial impact in monitoring human populations for chromosomal damage.
Metaphase analysis is time consuming and requires considerable skill, so simpler cytogenetic assays have been developed, of which micronucleus assays have become especially important. Micronuclei are membrane-bound structures that contain chromosomal fragments, or sometimes whole chromosomes, that were not incorporated into a daughter nucleus at mitosis. Because micronuclei usually represent acentric chromosomal fragments, they are most commonly used as simple indicators of chromosomal damage. However, the ability to detect micronuclei containing whole chromosomes has led to their use for detecting aneuploidy as well. Micronucleus assays may be conducted in primary cultures of human lymphocytes (Fenech et al., 2003; Corvi et al., 2008; Fenech, 2008; Fenech et al., 2011b), mammalian cell lines (Kirsch-Volders et al., 2003; Corvi et al., 2008), or mammals in vivo (Krishna and Hayashi, 2000; Hayashi et al., 2007; Dertinger et al., 2011).
Micronucleus assays in lymphocytes have been greatly improved by the cytokinesis-block technique in which cell division is inhibited with cytochalasin B, resulting in binucleate and multinucleate cells (Fenech et al., 2003, 2011b; Kirsch-Volders et al., 2003; Fenech, 2008). In the cytokinesis-block assay in human lymphocytes, nondividing (G0) cells are treated with ionizing radiation or a radiomimetic chemical and then stimulated to divide with the mitogen phytohemagglutinin. Alternatively, the lymphocytes may be exposed to the mitogen first, so that the subsequent mutagenic treatment with radiation or chemicals includes the S period of the cell cycle. In either case, cytochalasin B is added for the last part of the culture period, and micronuclei are counted only in binucleate cells so as to ensure that the cells have undergone a single nuclear division that is essential for micronucleus development. The assay thereby avoids confusion owing to differences in cellular proliferation kinetics.
Although first devised using primary lymphocytes (Fenech et al., 2003; Fenech, 2008; Fenech, 2011b), the cytokinesis-block micronucleus assay has since been adapted for use in continuous cell cultures, including the Chinese hamster and mouse lymphoma cells that are widely used in other genotoxicity assays (Kirsch-Volders et al., 2003; Corvi et al., 2008). Micronucleus assays should be conducted in such a way that cellular proliferation is monitored along with the micronucleus frequency, and this is facilitated by the cytokinesis block. Reliable data have been obtained in cultured cells both with and without cytokinesis block, but scoring results only in binucleate cells after blockage of cytokinesis with cytochalasin B confers advantages with respect to the measurement of proliferation, recognizing whether an agent is cytostatic, and obtaining clear dose–response relationships (Kirsch-Volders et al., 2003).
Although micronuclei resulting from chromosome breakage comprise the principal endpoint in the cytokinesis-block micronucleus assay, the method can also provide evidence of aneuploidy, chromosome rearrangements that form nucleoplasmic bridges, inhibition of cell division, necrosis, apoptosis, and excision-repairable lesions (Fenech et al., 2003; Fenech, 2008; Fenech et al., 2011b). Micronuclei in a binucleate human lymphocyte are shown in Fig. 9-7. A recent review of the International Human Micronucleus (HUMN) Project provides a comprehensive description of standardized protocols for micronucleus assays in human lymphocytes and buccal cells together with a review of associations between micronucleus data and disease outcomes (Fenech et al., 2011a).
Micronucleus in a human lymphocyte. The cytochalasin B method was used to inhibit cytokinesis that resulted in a binucleate nucleus. The micronucleus (arrow) resulted from failure of an acentric chromosome fragment or a whole chromosome being included in a daughter nucleus following cell division. (Figure courtesy of James Allen, Jill Barnes, and Barbara Collins.)
The in vivo micronucleus assay is often performed by counting micronuclei in immature (polychromatic) erythrocytes in the bone marrow of treated mice, but it may also be based on peripheral blood (Krishna and Hayashi, 2000; Hayashi et al., 2007). Micronuclei remain in the cell when the nucleus is extruded in the maturation of erythroblasts. In vivo micronucleus assays are increasingly used in genotoxicity testing as a substitute for bone marrow metaphase chromosome analysis. Micronucleus assays have been developed for mammalian tissues other than bone marrow and blood, including skin, duodenum, colon, liver, lung, spleen, testes, bladder, buccal mucosal cells, stomach, vagina, and fetal tissues (Coffing et al., 2011; Morita et al., 2011). Although assays in bone marrow and blood are the mainstay of genotoxicity testing, the new assays are important for mechanistic studies and research on the site specificity of genetic damage and carcinogenesis.
Micronuclei are most commonly visualized through microscopy, but automated means of detecting micronuclei are being developed through the application of flow cytometry. Flow cytometric detection is effective in micronucleus assays in rodent bone marrow or blood (Dertinger et al., 2011). It can also be used to detect micronuclei in Chinese hamster CHO cells, where altered flow cytometric parameters can reveal whether the micronuclei arose primarily by chromosome breakage or by aneuploidy (Bryce et al., 2011).
Sister Chromatid Exchanges
SCE, in which there has been an apparently reciprocal exchange of segments between the two chromatids of a chromosome, are visible cytologically through differential staining of chromatids. Fig. 9-8 shows SCE in human cells. Many mutagens induce SCE in cultured cells and in mammals in vivo (Tucker et al., 1993a; Wilson and Thompson, 2007). Despite the convenience and responsiveness of SCE assays, data on SCE are less informative than data on chromosome aberrations. There is uncertainty about the underlying mechanisms by which SCEs are formed and how DNA damage or perturbations of DNA synthesis stimulate their formation (Preston, 1991). SCE assays are therefore best regarded as general indicators of mutagen exposure, analogous to DNA damage and repair assays, rather than measures of a mutagenic effect.
Sister chromatid exchanges (SCEs) in human lymphocytes. (A) SCE in untreated cell. (B) SCE in cell exposed to ethyl carbamate. The treatment results in a very large increase in the number of SCE. (Figure courtesy of James Allen and Barbara Collins.)
Although assays for the induction of aneuploidy are not yet as refined as those for gene mutations and chromosome aberrations, they are being developed (Aardema et al., 1998). The targets of aneugens are often components of the mitotic or meiotic apparatus, rather than DNA. Therefore, aneugens should not be expected to overlap strongly with mutagens and clastogens. For example, a chemical that interferes with the polymerization of tubulin and thereby disrupts the formation of a mitotic spindle is likely to show specificity as an aneugen. Assays for chemicals that induce aneuploidy should therefore encompass all the relevant cellular targets that are required for the proper functioning of the mitotic and meiotic process. Means of detecting aneuploidy include chromosome counting (Galloway and Ivett, 1986; Natarajan, 1993; Aardema et al., 1998; Paccierotti and Sgura, 2008), the detection of micronuclei that contain kinetochores (Lynch and Parry, 1993; Natarajan, 1993; Aardema et al., 1998; Fenech, 2008; Paccierotti and Sgura, 2008), and the observation of abnormal spindles or spindle–chromosome associations in cells in which spindles and chromosomes have been differentially stained (Parry, 1998). FISH-based assays have also been developed for the assessment of aneuploidy in interphase somatic cells (Rupa et al., 1997; Paccierotti and Sgura, 2008) and in sperm (Russo, 2000; Marchetti et al., 2006, 2008).
A complication in chromosome counting is that a metaphase may lack chromosomes because they were lost during cell preparation for analysis, rather than having been absent from the living cell. To avoid this artifact, cytogeneticists generally use extra chromosomes (ie, hyperploidy) rather than missing chromosomes (ie, hypoploidy) as an indicator of aneuploidy in chromosome preparations from mammalian cell cultures (Galloway and Ivett, 1986; Aardema et al., 1998) or mouse bone marrow (Adler, 1993). Techniques for counting chromosomes in intact cells may allow reliable measures of hypoploidy (Natarajan, 1993), but the detection of hyperploidy remains the norm in lieu of clear evidence that artifactual chromosome loss has been avoided. It has been suggested that counting polyploid cells, which is technically straightforward, may be an efficient way to detect aneugens (Aardema et al., 1998), but there is disagreement on the point (Parry, 1998).
Micronucleus assays can detect aneugens as well as clastogens. Micronuclei that contain whole chromosomes tend to be somewhat larger than those containing chromosome fragments, but the two categories are not readily distinguished in typically stained preparations (Natarajan, 1993). However, one can infer that a micronucleus contains a whole chromosome if it is shown to contain a kinetochore or centromeric DNA. Aneuploidy may therefore be detected by means of antikinetochore antibodies with a fluorescent label or FISH with a probe for centromere-specific DNA (Lynch and Parry, 1993; Natarajan, 1993; Krishna and Hayashi, 2000; Fenech, 2008; Paccierotti and Sgura, 2008). Micronuclei containing kinetochores or centromeric DNA may be detected in cultured cells (Lynch and Parry, 1993; Aardema et al., 1998; Fenech, 2008; Paccierotti and Sgura, 2008) and in mouse bone marrow in vivo (Adler, 1993; Krishna and Hayashi, 2000). Frequencies of micronuclei ascribable to aneuploidy and to clastogenic effects may therefore be determined concurrently by tabulating micronuclei with and without kinetochores.
Germ cell mutagenesis assays are of special interest as indicators of genetic damage that can enter the gene pool and be transmitted through generations. Mammalian germ cell assays provide the best basis for assessing risks to human germ cells and therefore hold a central place in genetic toxicology despite their relative complexity and expense. The design of the test must compensate for the fact that mutations occur at low frequency, and even the simplest animal systems face the difficulty of their having a sufficiently large sample size. One can easily screen millions of bacteria or cultured cells by selection techniques, but screening large numbers of mice poses practical limitations. Therefore, a germ cell assay must offer a straightforward, unequivocal identification of mutants with minimal labor (Singer and Yauk, 2010).
The mouse specific-locus test detects recessive mutations that produce easily analyzed, visible phenotypes (coat pigmentation and ear size) conferred by seven genes (Russell and Shelby, 1985; Ehling, 1991; Russell and Russell, 1992; Favor, 1999; Russell, 2004; Singer et al., 2006). Mutants may be classified as having point mutations or chromosomal alterations on the basis of genetic and molecular analysis (Favor, 1999). The assay has been important in assessing genetic risks of ionizing radiation and has been used to study various chemical mutagens. Although they have been used less extensively, there are other gene mutation assays in mouse germ cells based on dominant mutations that cause skeletal abnormalities (Selby et al., 2004) or cataracts (Ehling, 1991) and recessive mutations that cause electrophoretic changes in proteins (Lewis, 1991).
Mammalian assays permit the measurement of mutagenesis at different germ cell stages (Favor, 1999; Russell, 2004). Late stages of spermatogenesis are often found to be sensitive to mutagenesis, but effects in spermatocytes, spermatids, and spermatozoa are transitory. Mutagenesis in stem-cell spermatogonia and resting oocytes is of special interest in genetic risk assessment because of the persistence of these stages throughout reproductive life. Chemical mutagens show specificity with respect to germ cell stages. For example, ethylnitrosourea and chlorambucil are both potent mutagens in the mouse specific-locus test, but the former induces primarily point mutations in spermatogonia, whereas the latter mostly induces deletions in spermatids (Russell and Russell, 1992). The ratio of deletions to point mutations is not only a function of the nature of the mutagen but depends on germ cell stage, as some mutagens induce higher proportions of gross alterations in late stages of spermatogenesis than in spermatogonia (Lewis, 1991; Favor, 1999; Russell, 2004).
There is currently no unequivocal evidence of induced gene mutations in human germ cells, but studies in mice leave little doubt about the susceptibility of mammalian germ cells to mutagenesis by radiation and chemicals. New molecular methods, particularly those involving the assessment of changes in tandem repeat loci (Yauk, 2004; Dubrova, 2005; Singer et al., 2006; Somers, 2006), hold great promise for the development of systems that will permit the efficient detection of genetic alterations in human germ cells. The development of methods based on expanded simple tandem repeat (ESTR) loci in mice and other species is important, in that it permits in situ monitoring for environmental mutagens and the quantification of mutagenesis after controlled exposures of laboratory animals using systems parallel to those being developed for human monitoring (Yauk, 2004; Dubrova, 2005; Wu et al., 2006; Somers, 2006). Basic research on mechanisms underlying ESTR changes is essential, as it is still unclear how ESTR changes relate to the gene mutations that have been long studied by population geneticists and are detected in classical gene-mutation assays.
Cytogenetic assays in germ cells are not routinely included in mutagenicity testing, but they are an important source of information for assessing risks to future generations posed by the induction of chromosome aberrations. Metaphase analysis of germ cells is feasible in rodent spermatogonia, spermatocytes, or oocytes (Kirkland et al., 1990; Tease, 1992; Russo, 2000; Marchetti et al., 2001). A micronucleus assay has also been developed in which chromosomal damage induced in meiosis is measured by the observation of rodent germ cells, principally spermatids (Russo, 2000; Hayashi et al., 2007).
Aneuploidy originating in mammalian germ cells may be detected cytologically through chromosome counting for hyperploidy (Allen et al., 1986; Adler, 1993; Aardema et al., 1998; Russo, 2000; Marchetti et al., 2001) or genetically in the mouse sex-chromosome loss test (Russell and Shelby, 1985), but these methods are not widely used in toxicological testing. A promising development is the detection of aneuploidy in the sperm of mice or rats by FISH with chromosome-specific probes (Baumgarthner et al., 1999; Russo, 2000; Marchetti et al., 2006, 2008). The presence of two fluorescent spots indicates the presence of an extra copy of the chromosome identified by the probe; probes for several chromosomes are used simultaneously so that aneuploid sperm are distinguishable from diploid sperm.
Besides cytological observation, indirect evidence for chromosome aberrations is obtained in the mouse heritable translocation assay, which measures reduced fertility in the offspring of treated males (Russell and Shelby, 1985; Singer et al., 2006). This presumptive evidence of chromosomal rearrangements can be confirmed through cytogenetic analysis. Data from the mouse heritable translocation test in postmeiotic male germ cells have been used in an attempt to quantify human germ cell risk for ethylene oxide, a mutagen used as a fumigant, sterilizing agent, and reactant in chemical syntheses (Rhomberg et al., 1990; Preston et al., 1995).
Dominant Lethal Mutations
The mouse or rat dominant lethal assay (Adler et al., 1994; Singer et al., 2006) offers an extensive database on the induction of genetic damage in mammalian germ cells. In the most commonly used version of the assay, males are treated on an acute or subchronic basis with the chemical of interest and then mated with virgin females at appropriate intervals. The females are killed and necropsied during pregnancy so that embryonic mortality may be characterized and quantified. Most dominant lethal mutations, manifested as intrauterine deaths, are thought to arise from chromosomal anomalies.
Development of Testing Strategies
Concern about adverse effects of mutation on human health, principally carcinogenesis and the induction of transmissible damage in germ cells, has provided the impetus to identify environmental mutagens. Priorities must be set for testing because it is not feasible to conduct multiple tests of all chemicals to which people are exposed. Such factors as production volumes, intended uses, the extent of human exposure, environmental distribution, and effects that may be anticipated on the basis of chemical structure or previous testing must be considered in order to ensure that compounds with the greatest potential for adverse effects receive the most comprehensive study. The most obvious use of genetic toxicology assays is screening chemicals to detect mutagens, but they are also used to obtain information on mutagenic mechanisms and dose–responses that contribute to an evaluation of hazards. Besides testing pure chemicals, environmental samples are tested because many mutagens exist in complex mixtures (DeMarini 1998; Ohe et al., 2003; White, 2004). The analysis of complex mixtures often requires a combination of mutagenicity assays and refined analytical methods (White, 2004; Hewitt and Marvin, 2005). Assessment of a chemical's genotoxicity requires data from well-characterized assays. Assays are said to be validated when they have been shown to perform reproducibly and reliably with many compounds from diverse chemical classes in several laboratories. An evaluation of test performance, however, sometimes extends beyond determining whether the assay effectively detects the specific endpoint that it actually measures to whether it is predictive of other endpoints of interest. For example, there is great interest in the ability of mutagenicity tests, which do not measure carcinogenicity per se, to predict whether chemicals are carcinogens.
Mutagenicity testing, combined with an evaluation of chemical structure, has been found to identify a large proportion of trans-species, multiple-site carcinogens (Tennant and Ashby, 1991; Gold et al., 1993). In contrast, some carcinogens are not detected as mutagens. Putatively nongenotoxic carcinogens often give responses that are more specific with respect to species, sites, and conditions (Ashby and Paton, 1993; Gold et al., 1993). In predicting carcinogenicity, one should consider both the sensitivity and the specificity of an assay. Sensitivity refers to the proportion of carcinogens that are positive in the assay, whereas specificity is the proportion of noncarcinogens that are negative (Tennant et al., 1987; McGregor et al., 1999). Sensitivity and specificity both contribute to the predictive reliability of an assay. The commonly held view that deficiencies in the sensitivity or specificity of individual assays may be circumvented by using assays in complementary combinations called tiers or batteries has fallen into disfavor because, rather than offsetting each other's strengths and weaknesses, genetic toxicology assays are often consistent with one another (Tennant et al., 1987; Ashby and Tennant, 1991; Kim and Margolin, 1999).
Strategies for testing have evolved over the last few decades, such that data from a few well-chosen assays are now considered sufficient (MacGregor et al., 2000). Rather than trying to assemble extensive batteries of complementary assays, it is prudent to emphasize mechanistic considerations in choosing assays. Such an approach makes a sensitive assay for gene mutations (eg, the Ames assay) and an assay for clastogenic effects in mammals pivotal in the evaluation of genotoxicity, and this is the basis for our highlighting these assays in Table 9-1. The Ames assay has performed reliably with hundreds of compounds in laboratories throughout the world. Other bacterial assays and mammalian cell assays also provide useful information on gene mutations. Beyond gene mutations, one should evaluate damage at the chromosomal level with a mammalian in vitro or in vivo cytogenetic assay. Cytogenetic assays in rodents are especially useful for this purpose because they combine a well-validated genetic assay with mammalian pharmacodynamics and metabolism. The other assays in Table 9-1 offer an extensive database on chemical mutagenesis (ie, Drosophila SLRL), a unique genetic endpoint (ie, aneuploidy; mitotic recombination), applicability to diverse organisms and tissues (ie, DNA damage assays, such as the comet assay), or special importance in the assessment of genetic risk (ie, germ cell assays). The more extensive listing of assays in Table 9-2 provides references that can be helpful in interpreting genetic toxicology data that can be found in the scientific literature.