Although behavioral assays offer alternative endpoints for assessing the impact of contami‐ nants on soil invertebrates, it is recommended to use such evaluations along with acute and/or chronic toxicity tests [35] because in certain cases, the substances can cause 100% mortality without the observance of an avoidance effect [32]. Such cases may be related to the test substance's narcotic properties, an absence of irritating effects, or physiological adapta‐ tions associated with the species' mode of life [36]. Integrated laboratory analyses should therefore be performed when a higher precision is required when assessing the interactions between contaminants, animal species, and soil properties because such analyses decrease the amount of uncertainty when determining ecological risks.

Although the standardized guidelines for ecotoxicological tests for terrestrial environments are relatively new, compared with the guidelines for aquatic environments, the number of tests based on these guidelines has increased considerably, and such tests have been used to investigate the ecological risk assessment of fungicides, herbicides, insecticides, heavy metals, nanomaterials, agro-industrial residues, and other substances in soil [13, 20].

#### **2. Standard species for soil ecotoxicology**

These two approaches include various tests that can be classified according to exposure time (acute or chronic toxicity), observed effect (mortality, reduced growth or reproduction, bioaccumulation, or behavioral changes) or effective response (lethal or sublethal) [21]. In these tests, representative species of the soil fauna are exposed to increasing contaminant concen‐ trations and the contaminant effects are measured in one (single species) or several species

There are various methods for evaluating toxicity in invertebrates, including topical applica‐ tion, force-feeding, and immersion tests [23]. However, the main laboratory assays standar‐ dized by the norms of the Organization for Economic Cooperation and Development (OECD) and International Organization for Standardization (ISO) consist of exposing standard species to samples of contaminated soil. These protocols describe methods used to determine acute and chronic toxicity and the effects on the behavior of earthworms, collembolans, enchytraeids,

The objective of acute toxicity tests is mainly to assess whether a substance causes organism death. These tests are useful for short-term identifications of highly toxic contaminants; however, they do not consider different stages of the test organism life cycle (growth, repro‐ duction, and birth of juveniles) or determine whether particular life cycle stages present increased sensitivity to toxic substances [24]. These tests are also used as preliminary evalua‐ tions ("range-finding tests") to determine the concentration ranges to be used in definitive acute toxicity tests and/or the sublethal concentrations for chronic toxicity assays [13].

Chronic toxicity tests are medium-term tests that measure the sublethal effects of potentially toxic substances on organisms, such as changes in reproduction and growth, and are more adequate for assessing effects at the population level [20, 25]. The primary standard methods for laboratory chronic toxicity tests have been established in ISO [26–27] and OECD [28–31] guidelines. The objective of these standardized tests is similar for different groups of inverte‐ brates, although they do present differences, especially in test duration, as a result of the different reproductive characteristics of different species. In these tests, adults are exposed to a range of sublethal concentrations of the test substance, with the concentrations defined

Behavioral tests with soil invertebrates are also becoming common because they provide a preliminary evaluation of responses to soil pollution over a shorter period relative to that of toxicity tests [13]. In addition to providing ecologically relevant results because of the sensi‐ tivity of species in detecting polluting substances in soils, these tests can be performed more quickly (2 days on average) and at a reduced cost [13, 32]. Avoidance tests, for example, can be used as triage tools to assess the habitat suitability of soils because they are based on the ability of animals (e.g., earthworms) to avoid potentially toxic substances upon exposure to contaminated soils [33] because of the presence of chemoreceptors that are highly sensitive to

Although behavioral assays offer alternative endpoints for assessing the impact of contami‐ nants on soil invertebrates, it is recommended to use such evaluations along with acute and/or chronic toxicity tests [35] because in certain cases, the substances can cause 100%

according to preliminary tests (range finding) or results from the literature.

(multispecies) to test dose–response relations [22].

38 Invertebrates - Experimental Models in Toxicity Screening

mites, mollusks, and few other insects [20].

chemicals in the environment [34].

Ideally, the toxic effects of all chemicals introduced to terrestrial ecosystems, such as agroindustrial and urban wastes, should be tested on all species inhabiting the ecosystem before commercialization (xenobiotics) or direct application to the soil [13]. Because these measures cannot be achieved in the laboratory, edaphic invertebrate species that have known sensitiv‐ ities to anthropogenic changes and provide the main representative ecosystem services of the fauna have been chosen as indicators of the ecotoxicological risk to terrestrial ecosystems. The review presented in Ref. [20] provides a list of the main invertebrates used in terrestrial ecotoxicological assays. This list includes earthworms (*Eisenia andrei* and *Eisenia fetida*), enchytraeids (*Enchytraeus albidus* and *Enchytraeus crypticus*), mollusks (*Helix aspersa*), mites (*Hypoaspis aculeifer*, *Platynothrus peltifer*, and *Oppia nitens*), isopods (*Porcellio scabere* and *Porcellionides pruinosis*), collembolans (*Folsomia candida* and *Folsomia fimetaria*), insects of the family Carabidae (*Pterostichus oblongpunctatus* and *Poecilus cupreus*), *Oxythyrea funesta*, and other organisms used in the methods standardized by international guidelines for ecological risk assessments of soil contaminants.

The methods described by the ISO and OECD international guidelines for ecotoxicity tests with terrestrial invertebrates are designed to standardize the tests so that similar results can be obtained in different laboratories regardless of the region. Such standardization facilitates comparisons and increases the reliability of the established toxicity levels. To develop stand‐ ardized tests, standard species must be selected based on ecological relevance, ease of maintenance in the laboratory, and short-generation time [37–38]. In addition, the selected species should have well-known biological parameters so that a large number of healthy, homogeneous (same size and biomass), and age-synchronized individuals can be obtained. The number of species that meet all of these requirements is small.

In general, toxicity assays with standard species are performed individually; therefore, each species is tested separately to exclude the effects of interactions among species present in soil. However, the use of several species, even when tested separately, increases the ecological relevance of laboratory analyses because different organisms respond differently to pollutants, and the potential risk to the ecosystem varies [20]. The performance of a balanced battery of tests using organisms of different functional and taxonomic groups and different routes of exposure is therefore necessary to improve the reliability of ecological risk assessments determined via laboratory tests [39].

The main standard invertebrates used in batteries of soil ecotoxicological tests are earthworms [29, 40–42], collembolans [26, 31], mites [39], and enchytraeids [22, 28, 43]. Because of different morphological (e.g., epidermis structure) and physiological (e.g., water and oxygen uptake pathways) characteristics as well as feeding and behavioral habits (e.g., movement within the soil or at the soil surface and digging habits), these taxonomic groups encompass different pollutant uptake routes and encounter pollutants through the exposure to water and air present in the soil pores and ingestion of food and soil particles [44].

Earthworms are important components of the soil biota because they aid in the formation and maintenance of soil structure and fertility. Although they are not numerically dominant, their large size makes them one of the main contributors to invertebrate biomass in soil. These organisms are important indicators of soil life and quality because their populations are affected by common agricultural practices, especially by the use of pesticides and fertilizers and application of waste [45]. Because they are soft-bodied organisms, earthworms absorb water mainly through their skin; therefore, they can accumulate chemicals during water absorption [44]. Another important route of contaminant absorption is through ingestion because these organisms ingest large amounts of soil with adsorbed substances along with their food (soil organic material, which may also be contaminated).

Although several earthworm species have been used in terrestrial ecotoxicological tests [22], only *E. fetida* and *E. andrei* were included in the ISO and OECD guidelines. These species are preferred because they have worldwide distribution, are naturally tolerant to various organic substrates, are easily handled in single species or mixed cultures [45], and may be easily acquired commercially (adults, juveniles, or cocoons) or obtained from other soil ecotoxicology laboratories.

*E. fetida* and *E. andrei* were initially differentiated into two forms, var. tipica and var. unicolor (lightly stripped and uniformly pigmented, respectively), by Bouché (1972), who considered them as subspecies (*E. fetida fetida* and *E. fetida andrei*). These organisms were thought to belong to the same species for a long time because of their similar appearances, ecological demands, and frequent associations. However, recent investigations have determined that their crossing does not result in viable descendants, and biochemical methods were used to confirm that they constitute different species [46]. This differentiation is important for their use in ecotoxicolog‐ ical tests because the effects of contaminants on the two species may be different.

Neither of the two species is typical of most agricultural soils, and they only occur in soils rich in organic matter. Under ideal conditions, their life cycle until maturity is relatively short (varying between 45 and 51 days) compared with that of other species and extends from the recently deposited cocoon until the adult stage, when the worms are sexually mature (with the presence of clitellum) and ready to produce the next generation. The time for juveniles (recently hatched) to reach sexual maturity varies between 21 and 30 days. Both species are prolific, and between two and five juveniles are generated from each viable cocoon. Depending on the rearing temperature and substrate, their maximum life span ranges from four and a half to five years [45].

and the potential risk to the ecosystem varies [20]. The performance of a balanced battery of tests using organisms of different functional and taxonomic groups and different routes of exposure is therefore necessary to improve the reliability of ecological risk assessments

The main standard invertebrates used in batteries of soil ecotoxicological tests are earthworms [29, 40–42], collembolans [26, 31], mites [39], and enchytraeids [22, 28, 43]. Because of different morphological (e.g., epidermis structure) and physiological (e.g., water and oxygen uptake pathways) characteristics as well as feeding and behavioral habits (e.g., movement within the soil or at the soil surface and digging habits), these taxonomic groups encompass different pollutant uptake routes and encounter pollutants through the exposure to water and air

Earthworms are important components of the soil biota because they aid in the formation and maintenance of soil structure and fertility. Although they are not numerically dominant, their large size makes them one of the main contributors to invertebrate biomass in soil. These organisms are important indicators of soil life and quality because their populations are affected by common agricultural practices, especially by the use of pesticides and fertilizers and application of waste [45]. Because they are soft-bodied organisms, earthworms absorb water mainly through their skin; therefore, they can accumulate chemicals during water absorption [44]. Another important route of contaminant absorption is through ingestion because these organisms ingest large amounts of soil with adsorbed substances along with

Although several earthworm species have been used in terrestrial ecotoxicological tests [22], only *E. fetida* and *E. andrei* were included in the ISO and OECD guidelines. These species are preferred because they have worldwide distribution, are naturally tolerant to various organic substrates, are easily handled in single species or mixed cultures [45], and may be easily acquired commercially (adults, juveniles, or cocoons) or obtained from other soil ecotoxicology

*E. fetida* and *E. andrei* were initially differentiated into two forms, var. tipica and var. unicolor (lightly stripped and uniformly pigmented, respectively), by Bouché (1972), who considered them as subspecies (*E. fetida fetida* and *E. fetida andrei*). These organisms were thought to belong to the same species for a long time because of their similar appearances, ecological demands, and frequent associations. However, recent investigations have determined that their crossing does not result in viable descendants, and biochemical methods were used to confirm that they constitute different species [46]. This differentiation is important for their use in ecotoxicolog‐

Neither of the two species is typical of most agricultural soils, and they only occur in soils rich in organic matter. Under ideal conditions, their life cycle until maturity is relatively short (varying between 45 and 51 days) compared with that of other species and extends from the recently deposited cocoon until the adult stage, when the worms are sexually mature (with the presence of clitellum) and ready to produce the next generation. The time for juveniles (recently hatched) to reach sexual maturity varies between 21 and 30 days. Both species are

ical tests because the effects of contaminants on the two species may be different.

present in the soil pores and ingestion of food and soil particles [44].

their food (soil organic material, which may also be contaminated).

determined via laboratory tests [39].

40 Invertebrates - Experimental Models in Toxicity Screening

laboratories.

Enchytraeids (family Enchytraeidae) belong to the same class as earthworms (Oligochaeta) and can live in both water and soil. However, they are mostly found in soil, where they perform important ecosystem services, such as increasing the rate of organic matter decomposition, maintaining the soil structure (creation of biopores), and dispersing microbes on a local scale [47–48]. These services are especially important in acidic, sandy, and nutrient-poor soils, where enchytraeids are the dominant soil fauna group (up to 75% of the biomass) [48–49]. In these environments, the role of enchytraeids in organic matter decomposition may not be performed by other fauna groups [50].

The family Enchytraeidae has over 600 described species [48]. Known as white worms or potworms because of their pale color and small size (many are only a few millimeters long, although some may reach up to 5 cm), most enchytraeids are hermaphroditic (capable of selffertilization), although certain species are parthenogenetic or reproduce through fragmenta‐ tion [51]. Enchytraeids have a limited capacity for movement inside the soil; therefore, they live in the most superficial soil layers (0–10 cm) where the organic material and biological activity are concentrated. These organisms are found from arctic to tropical regions, and they are more abundant in forest soils (or soils rich in organic material) and less abundant in pastures and agricultural fields [52]. Their main food source is fungal mycelium; however, they also feed on organic matter that has been predigested by fungi as well as on other microorganisms [48].

These oligochaetes are sensitive to potentially toxic substances abundant in many soils where earthworms are not present or are not well represented. In addition, these animals are easy to handle and rear and have a significantly shorter life cycle than other worms, which is con‐ venient for standardized toxicity assays [28, 43, 53]. These organisms live in close contact with the soil pore water and are exposed to soil contaminants through dermal, intestinal (through feeding), and respiratory routes [44]. Although their use in laboratory ecotoxicological tests was reported for the first time approximately 40 years ago, enchytraeids were selected for use in standardized ecotoxicological laboratory tests only 10 years ago as reported in the guidelines ISO 16387 [43] and OECD 220 [28].

Enchytraeus is the only enchytraeid genus with species selected for ecotoxicological tests standardized by ISO and OECD guidelines (e.g., *E. albidus* and *E. crypticus*) because this genus is considered typical of environmental stress indicator organisms and can be easily reared in the laboratory. *E. albidus* is the best known species for soil ecotoxicology [54] because it can be reared in many different substrates (it is widely distributed in terrestrial ecosystems) with different types of food, and it has a proven sensitivity to soil contaminants [55]. When these animals reach maturity (approximately 21 days at 18°C), the size of *E. albidus* adults can vary between 15 and 40 mm. Variations are observed in the total developmental cycle (33–74 days), embryonic development period (12–18 days), eggs per cocoon (7–10), and viable cocoon percent (40–50%) according to environmental conditions, especially in response to changes in culture temperature [43, 54]. The ideal reproduction temperature for the species is 15°C, although they can reproduce at temperatures between 12 and 22°C. Temperatures above 25°C should be avoided because they can suppress reproduction [54].

Although *E. crypticus* has lower ecological relevance because its prevalence in the field is unknown [56], the species is adequate for ecotoxicological laboratory tests, and its use in current standardized tests appears to be increasing [20]. Compared with *E. albidus*, *E. crypti‐ cus* has the advantage of being able to grow in agar medium, and it also has a higher repro‐ duction rate, has a shorter generation time, tolerates a wider range of soil properties, and presents other characteristics considered methodologically advantageous [53, 57]. The adults of this species vary in sizes between 3 and 12 mm, and they have a generation time of approx‐ imately 18 days (at 21°C) in agar medium. The mean number of eggs per cocoon can vary between 1 and 35 with a mean of 4.6 eggs produced per day [58]. Studies have indicated that the number of juveniles may also vary according to the type of soil used, with apparently higher numbers in standard LUFA 2.2 natural soil [59] than in artificial OECD soil [60].

The order Collembola is one of the most diverse and abundant terrestrial arthropod orders, with 21 families and 20,000 described species [61]. In general, collembolans are small, varying from a few to approximately 10 mm in length [62]. The body can exhibit colorful pigmenta‐ tion,althoughthe inhabitantsofdeeper soillayersare typicallynotpigmented[63].Most species feed on fungal hyphae and decomposing plant material; thus, they have a significant effect on microbial ecologyandsoilfertilityandcancontrol certainplantdiseases causedbyfungi[62,64].

Collembolans are vulnerable to the presence of potentially toxic contaminants in soil [3] because they are exposed through water ingestion or absorption from wet/moist surfaces, food (living or dead) consumption, and soil pore air inhalation [44]. The responses obtained in tests using these arthropods may indicate environmental stress levels and the ecological risk of substances; therefore, these organisms supply information that can serve as a basis for legislation [65]. Collembolans have been used to estimate the effects of pesticides and other environmental pollutants on nontarget soil arthropods for approximately four decades, with *F. candida –* being typically used in standardized ecotoxicological tests [26, 31, 66]. *F. candida* has great importance for terrestrial ecosystem services because of its high sensitivity, short generation time, high reproduction rate, and easy culturing in the laboratory using a diet of granulated dry yeast [67].

*F. candida* (Willem 1902) is an arthropod of the family Isotomidae, and it is distributed in soils worldwide, although it is not common in most agricultural soils. This animal has a high occurrence rate in sites rich in organic matter [68]. This species has no pigmentation or eyes [38], and it reproduces exclusively through parthenogenetic females, which are approximately 2 mm long and sexually mature at 21–24 days of age (at 20°C). The optimal temperature for egg incubation and production is 21°C, and under these conditions, females lay 30–50 eggs, which take 7–10 days to hatch. This species has strong feeding preferences for certain species of fungi, and they are classified as microsaprophagous [62] and can be fed dry yeast in the laboratory [26, 31].

Although used in a fewer number of studies, *F. fimetaria* has been used as complementary or alternative species to *F. candida*, and the choice to use *F. fimetaria* is related to its higher ecological relevance because it is present in many natural and agricultural habitats where *F. candida* is not found. In addition, *F. fimetaria* meets all of the necessary requirements of a standard species for ecotoxicological laboratory tests [69]. In 2009, OECD guideline no. 232 [31] established *F. fimetaria* as a standard species for standardized laboratory assays. In addition to *F. fimetaria*, other species can be used to increase the ecological relevance of tests, although they may not be included in guidelines [21, 68, 70–71].

although they can reproduce at temperatures between 12 and 22°C. Temperatures above 25°C

Although *E. crypticus* has lower ecological relevance because its prevalence in the field is unknown [56], the species is adequate for ecotoxicological laboratory tests, and its use in current standardized tests appears to be increasing [20]. Compared with *E. albidus*, *E. crypti‐ cus* has the advantage of being able to grow in agar medium, and it also has a higher repro‐ duction rate, has a shorter generation time, tolerates a wider range of soil properties, and presents other characteristics considered methodologically advantageous [53, 57]. The adults of this species vary in sizes between 3 and 12 mm, and they have a generation time of approx‐ imately 18 days (at 21°C) in agar medium. The mean number of eggs per cocoon can vary between 1 and 35 with a mean of 4.6 eggs produced per day [58]. Studies have indicated that the number of juveniles may also vary according to the type of soil used, with apparently higher numbers in standard LUFA 2.2 natural soil [59] than in artificial OECD soil [60].

The order Collembola is one of the most diverse and abundant terrestrial arthropod orders, with 21 families and 20,000 described species [61]. In general, collembolans are small, varying from a few to approximately 10 mm in length [62]. The body can exhibit colorful pigmenta‐ tion,althoughthe inhabitantsofdeeper soillayersare typicallynotpigmented[63].Most species feed on fungal hyphae and decomposing plant material; thus, they have a significant effect on microbial ecologyandsoilfertilityandcancontrol certainplantdiseases causedbyfungi[62,64].

Collembolans are vulnerable to the presence of potentially toxic contaminants in soil [3] because they are exposed through water ingestion or absorption from wet/moist surfaces, food (living or dead) consumption, and soil pore air inhalation [44]. The responses obtained in tests using these arthropods may indicate environmental stress levels and the ecological risk of substances; therefore, these organisms supply information that can serve as a basis for legislation [65]. Collembolans have been used to estimate the effects of pesticides and other environmental pollutants on nontarget soil arthropods for approximately four decades, with *F. candida –* being typically used in standardized ecotoxicological tests [26, 31, 66]. *F. candida* has great importance for terrestrial ecosystem services because of its high sensitivity, short generation time, high reproduction rate, and easy culturing in the laboratory using a diet of

*F. candida* (Willem 1902) is an arthropod of the family Isotomidae, and it is distributed in soils worldwide, although it is not common in most agricultural soils. This animal has a high occurrence rate in sites rich in organic matter [68]. This species has no pigmentation or eyes [38], and it reproduces exclusively through parthenogenetic females, which are approximately 2 mm long and sexually mature at 21–24 days of age (at 20°C). The optimal temperature for egg incubation and production is 21°C, and under these conditions, females lay 30–50 eggs, which take 7–10 days to hatch. This species has strong feeding preferences for certain species of fungi, and they are classified as microsaprophagous [62] and can be fed dry yeast in the

Although used in a fewer number of studies, *F. fimetaria* has been used as complementary or alternative species to *F. candida*, and the choice to use *F. fimetaria* is related to its higher

should be avoided because they can suppress reproduction [54].

42 Invertebrates - Experimental Models in Toxicity Screening

granulated dry yeast [67].

laboratory [26, 31].

Mites are arthropods belonging to class Arachnida and subclass Acari, and they have a small size and unsegmented bodies [61]. In total, mites are ordered in 1,200 families and approxi‐ mately 500,000 species [72], of which many are the most abundant mesofauna inhabitants in many types of soil and litter [73]. Suborder Gamasida (order Mesostigmata) includes the main species of predatory mites inhabiting soil pores [74]. The community structure and abundance of predatory mites are strongly dependent on the nature and availability of their prey [22]. Most mites feed on enchytraeids, nematodes, and microarthropods, although certain groups are considered fungivorous, bacteriophagous, facultative phytophagous, or have unknown feeding habits [73].

The ecosystem services provided by soil predatory mites include the biological control of pests and other species with abundant populations; thus, they significantly contribute to the flow of energy and matter in terrestrial ecosystems as well as to the maintenance of food chains [22]. In addition, several genera have been isolated for over 30 years from soil and tested as quality bioindicators [72]. Currently, reports are available for different toxicity tests using these arthropods; however, only the reproduction test using *H. aculeifer* has been standardized by guidelines for the evaluation of soil quality [20, 22, 30]. *H. aculeifer* has been considered the most adequate mite species for ecotoxicological assays because it has an acceptable generation time (approximately 1 month at 20°C), is a generalist predator, and can be easily handled in the laboratory [20, 72]. Because it represents a different trophic level from the other inverte‐ brates used in standardized tests and is exposed to contaminants through different routes [44], *H. aculeifer* has been included on the EU community's list of nontarget organisms considered in the assessment of environmental risk of pesticides in soil [16, 72, 75].

Specimens of *H. aculeifer* are brown and have a light-brown dorsal shield. Although the size of both sexes is rather small, females are larger (0.8–0.9 mm) than males (0.55–0.65 mm) [22, 76]. Under temperatures between 20 and 23°C, these organisms become adults in approxi‐ mately 16 (females) and 18 (males) days after going through larval, protonymph, and deuto‐ nymph developmental stages. However, their development time can be strongly affected by the temperature [76]. Usually, reproduction is sexual, although arrhenotokous parthenogen‐ esis may occur in the absence of males, with this process only generating males [77]. However, females generally occur at a higher frequency because the sex ratio can also be controlled through selective cannibalism [78]. Each female lays approximately 100 eggs during its reproductive life [79]. These eggs are white, elliptical, and laid at the soil or culture substrate surface. Although it is a polyphagous predator, it is usually fed with the mites *Tyrophagus putrescentiae* or *Rhizoglyphus* sp. in ecotoxicological tests [30]. In the case of food scarcity, these organisms can survive cannibalistically [78].

In Ref. [22], the main invertebrates used in standardized terrestrial ecotoxicological tests were compared and it was concluded that the standard species of earthworms (*E. andrei* and *E. fetida*), enchytraeids (*E. albidus* and *E. crypticus*), collembolans (*F. candida* and *F. fimetaria*), and predatory mites (*H. aculeifer*) are adequate for the performance of tests on several soils from temperate and tropical regions. However, earthworms present limitations in acidic or basic soils. Enchytraeids are more tolerant to changes in soil pH and organic-matter concentrations, although they grow better in sandier soils. Mites and collembolans appear to be adequate for tests in most soil types and are considered less sensitive than oligochaetes (earthworms and enchytraeids) to soil properties. In tests that use soils with extreme characteristics (acidic or sandy soils), more than one species as well as alternative species should be used [22]. In addition, *H. aculeifer* generally appears to be less sensitive to certain substances compared with the remaining species. However, because it is the only predatory standard species, its inclusion in routine assessments of ecotoxicity is supported [80].

### **3. Summary of the standard procedures for bioassays with soil invertebrates**

Performing ecotoxicological laboratory tests requires a series of steps, including planning and material preparation, animal rearing and maintenance in the laboratory, contamination of artificial/natural soils (or preparation of previously contaminated soil samples), experimental procedures for the initial test conditions as well as the maintenance and evaluation of toxicity tests using organisms, and data analysis and interpretation. The steps of the standardized tests using the species *E. andrei*/*E. fetida*, *E. albidus*/*E. crypticus*, *F. candida*, and *H. aculeifer* described in the guidelines ISO no. 11268-2 [27], no. 16387 [43], no. 11267 [26] and OECD no. 226 [30], respectively, will be summarized in this section to describe the main methods established by the guidelines and the adaptations that are used in regions with tropical climates.

Most standardized tests using soil invertebrates were developed to quantify the impact of chemical exposure on organisms in artificial soils [42]. However, although studies using artificial soils supply information that can be internationally compared, natural soils may provide information on local problems. Although the practice is still not described by the guidelines, the use of natural soils in standardized ecotoxicological tests has been increasing.

OECD soil is a standard artificial substrate recommended by ISO/OECD guidelines for most terrestrial ecotoxicology studies [42]. This substrate consists of a mixture of 70% industrial sand (with more than 50% particles between 0.05 and 0.2 mm), 20% kaolinite clay, and 10% peat (ground and dry). However, for assays in tropical regions, studies have used a modified version of this substrate [81–83] known as tropical artificial soil (TAS). TAS uses powdered coconut husks as replacement for peat because of its higher availability in tropical regions. In both cases, after the materials are mixed, the pH (1 M KCL 1:5 weight:volume ratio) of the artificial soil should be adjusted to 6.0 ± 0.5 through the addition of CaCO3. In addition, the soil water-holding capacity should be determined for moisture adjustments [26].

To use natural soils in standardized tests, the soils should offer minimum conditions for the survival and reproduction of the test species without causing morphological or behavioral changes in the absence of the contaminant. To test chemicals for regulation purposes (e.g., ecological risk of pesticides), natural soils are artificially contaminated, and the results should be comparable between laboratories. Therefore, the soils should have similar characteristics, as is the case for LUFA soils [84–85], EURO soils [86], SIM soils [87], and other natural soils selected as standard soils for specific regions, for example, Polish [88] and Mediterranean soils [89]. In the case of natural soils from contaminated areas, the use of a control soil with the same characteristics (texture, pH, organic matter concentration, and C:N ratio) but without contam‐ inants is recommended [87].

In Ref. [22], the main invertebrates used in standardized terrestrial ecotoxicological tests were compared and it was concluded that the standard species of earthworms (*E. andrei* and *E. fetida*), enchytraeids (*E. albidus* and *E. crypticus*), collembolans (*F. candida* and *F. fimetaria*), and predatory mites (*H. aculeifer*) are adequate for the performance of tests on several soils from temperate and tropical regions. However, earthworms present limitations in acidic or basic soils. Enchytraeids are more tolerant to changes in soil pH and organic-matter concentrations, although they grow better in sandier soils. Mites and collembolans appear to be adequate for tests in most soil types and are considered less sensitive than oligochaetes (earthworms and enchytraeids) to soil properties. In tests that use soils with extreme characteristics (acidic or sandy soils), more than one species as well as alternative species should be used [22]. In addition, *H. aculeifer* generally appears to be less sensitive to certain substances compared with the remaining species. However, because it is the only predatory standard species, its inclusion

in routine assessments of ecotoxicity is supported [80].

44 Invertebrates - Experimental Models in Toxicity Screening

**invertebrates**

**3. Summary of the standard procedures for bioassays with soil**

the guidelines and the adaptations that are used in regions with tropical climates.

soil water-holding capacity should be determined for moisture adjustments [26].

Performing ecotoxicological laboratory tests requires a series of steps, including planning and material preparation, animal rearing and maintenance in the laboratory, contamination of artificial/natural soils (or preparation of previously contaminated soil samples), experimental procedures for the initial test conditions as well as the maintenance and evaluation of toxicity tests using organisms, and data analysis and interpretation. The steps of the standardized tests using the species *E. andrei*/*E. fetida*, *E. albidus*/*E. crypticus*, *F. candida*, and *H. aculeifer* described in the guidelines ISO no. 11268-2 [27], no. 16387 [43], no. 11267 [26] and OECD no. 226 [30], respectively, will be summarized in this section to describe the main methods established by

Most standardized tests using soil invertebrates were developed to quantify the impact of chemical exposure on organisms in artificial soils [42]. However, although studies using artificial soils supply information that can be internationally compared, natural soils may provide information on local problems. Although the practice is still not described by the guidelines, the use of natural soils in standardized ecotoxicological tests has been increasing.

OECD soil is a standard artificial substrate recommended by ISO/OECD guidelines for most terrestrial ecotoxicology studies [42]. This substrate consists of a mixture of 70% industrial sand (with more than 50% particles between 0.05 and 0.2 mm), 20% kaolinite clay, and 10% peat (ground and dry). However, for assays in tropical regions, studies have used a modified version of this substrate [81–83] known as tropical artificial soil (TAS). TAS uses powdered coconut husks as replacement for peat because of its higher availability in tropical regions. In both cases, after the materials are mixed, the pH (1 M KCL 1:5 weight:volume ratio) of the artificial soil should be adjusted to 6.0 ± 0.5 through the addition of CaCO3. In addition, the

Natural soils should be dried, sieved (5 or 2 mm, preferentially), and chemically and physically characterized before use, and at least the texture, pH, WHC, and moisture content should be determined. In addition, the organic matter concentration (or organic C), CEC, C:N ratio, and metal (or other element) concentration may also be measured [87]. Regardless of the type of soil (artificial or natural), defauning is recommended (by soil freezing at −20°C followed by thawing to room temperature) to eliminate the original soil fauna organisms [90].

According to the guidelines, incorporating test substances directly into soil (artificial contam‐ ination) varies with the water solubility of the contaminant, and there are three main methods: (a) for water-soluble substances; (b) for water-insoluble but organic solvent-soluble substan‐ ces; and (c) for water-insoluble and organic solvent-insoluble substances. In all cases, the concentration gradients of the test substances should be prepared immediately before the beginning of the assay in the volume necessary to maintain the soil moisture between 40 and 60% of its WHC. In the case of water-insoluble substances, after applying the solutions in increasing concentrations, the soil moisture should be adjusted through the addition of pure water. It is recommended that the concentrations be prepared in a geometric series separated by a factor of 1.8 or lower. If effects are not observed for the tested substance (e.g., active ingredient of a pesticide) at the highest concentration (1,000 mg kg−1) in the preliminary tests (acute toxicity assays), then a limit test should be performed to evaluate the toxicity using only the control treatment at a concentration of 1,000 mg kg−1.

The environmental conditions of the standardized ecotoxicological tests and laboratory cultures should be controlled. Controlled temperatures and light conditions in the test/culture chamber/room are fundamental for obtaining homogeneous cultures (with the same age and size) with development cycles that occur within the time predicted for the tests. The main recommended protocols are a mean temperature of 20 ± 2°C and constant light intensity between 400 and 800 lux on the culture containers [26, 31]. In addition, it is recommended that cultures be kept under controlled light/dark cycles, preferably 12 hours light/12 hours dark or 16 hours light/8 hours dark. However, the environmental test and culture conditions have been adapted for regions with predominantly tropical climates to better reflect the influence of local climate conditions. To simulate tropical conditions, temperatures varying between 23 and 27°C have been used [83, 91–93].

The earthworms *E. andrei* and *E. fetida* should be reared in substrates composed of a mix of horse or cow manure (defauned following the same process described for soils) and peat (1:1, dry weight) [27], and the culture medium should be moistened weekly with pure water. Similar to artificial soils, peat is usually replaced with powdered coconut husks in tropical climate regions [13, 18, 81]. Earthworms should be fed weekly with a mixture of oat flakes and water, and the rearing substrate should be periodically replaced.

*E. crypticus* can be cultured in natural soil, although recent studies have opted for culturing in Petri dishes with agar medium [83, 94]. This culture medium is composed of a mixture (1:1, v:v) of a salt solution (calcium chloride, magnesium sulfate, potassium chloride, and sodium bicarbonate) and a bacto-agar solution (e.g., Oxoid - Agar No. 1). Approximately 50 mg of ground oat flakes should be supplied as food once a week, and the organisms should be transferred to new substrate every 2 months.

Laboratory cultures of collembolans and mites should be performed using a substrate composed of a mixture of activated charcoal, plaster of Paris (calcium sulfate), and deionized water, with a recommended 1:8 charcoal:plaster of Paris ratio for *F. candida* and 1:9 ratio for *H. aculeifer* (w:w) [26, 30]. The volume of deionized water should be 60–100 mL for each 100 g of mixture, although the water content varies with the type of plaster of Paris. The bottom of plastic containers should be filled with the mixture to a height of approximately 1 cm. Collembolans should be fed dry yeast once a week, and cultures of *T. putrescentiae* or *Calogly‐ phus* sp. (cheese mites) should be simultaneously maintained with *H. aculeifer* cultures to serve as food (prey) for the predator. Small quantities of cheese mites should be supplied to *H. aculeifer* twice a week and the cheese mites should be fed once a week with powdered brewer's yeast [30].

Avoidance assays with *E. andrei*/*E. fetida* or *F. candida*, which are described by the guidelines ISO:17512-1 and ISO:17512-2 [41, 95], are performed using rectangular (earthworms) or round (collembolans) plastic boxes that are divided into two equal compartments by a plastic divider vertically introduced and filled with the test soil (amounts depend on the size of the container). Contaminated soil is added to one of the compartments, and the same amount of the respective control soil is added to the other compartment. Ten *E. andrei* or *E. fetida* adults (with developed clitellum) or 20 *F. candida* individuals at 10–12 days of age (originating from synchronized cultures) are then placed on the separation line between the two compartments, and the plastic divider is removed. The containers are then closed with perforated lids to allow air circulation. The animals are not fed during the test. After 48 hours, the number of individuals present in each compartment is recorded according to the specific method for each species [18, 96]. A double control test is also performed with control soil in both compartments to determine whether the organisms are randomly distributed between the two compartments in the absence of contaminants.

Chronic toxicity tests with *E. andrei*/*E. fetida* should be performed according to ISO:11268-2 [27]. Round plastic containers are filled with approximately 500 g soil (dry weight) treated with solutions with increasing contaminant concentrations (or dilutions of the naturally contami‐ nated soils) or the respective control soils. Ten adult earthworms (with visible clitellum) with individual weights between 300 and 600 mg that had been previously incubated in control soil for at least 24 hours are selected for each experimental unit. The containers are closed with perforated lids, and the earthworms are fed horse manure (≈5 g per replicate) at the beginning of the assay and then once a week until the end of the assay. The assay lasts for a total of 56 days. Adult earthworms are removed after 28 days, washed with water, and weighed. The final percentage of body biomass (after 28 days) relative to the initial weight can be calculated to assess the effects of contaminants on organismal growth. After 56 days from the beginning of the assay, the number of juveniles is counted to verify the treatment effect on species reproduction (for additional details, see Ref. [27], and/or Ref. [18]).

dry weight) [27], and the culture medium should be moistened weekly with pure water. Similar to artificial soils, peat is usually replaced with powdered coconut husks in tropical climate regions [13, 18, 81]. Earthworms should be fed weekly with a mixture of oat flakes and water,

*E. crypticus* can be cultured in natural soil, although recent studies have opted for culturing in Petri dishes with agar medium [83, 94]. This culture medium is composed of a mixture (1:1, v:v) of a salt solution (calcium chloride, magnesium sulfate, potassium chloride, and sodium bicarbonate) and a bacto-agar solution (e.g., Oxoid - Agar No. 1). Approximately 50 mg of ground oat flakes should be supplied as food once a week, and the organisms should be

Laboratory cultures of collembolans and mites should be performed using a substrate composed of a mixture of activated charcoal, plaster of Paris (calcium sulfate), and deionized water, with a recommended 1:8 charcoal:plaster of Paris ratio for *F. candida* and 1:9 ratio for *H. aculeifer* (w:w) [26, 30]. The volume of deionized water should be 60–100 mL for each 100 g of mixture, although the water content varies with the type of plaster of Paris. The bottom of plastic containers should be filled with the mixture to a height of approximately 1 cm. Collembolans should be fed dry yeast once a week, and cultures of *T. putrescentiae* or *Calogly‐ phus* sp. (cheese mites) should be simultaneously maintained with *H. aculeifer* cultures to serve as food (prey) for the predator. Small quantities of cheese mites should be supplied to *H. aculeifer* twice a week and the cheese mites should be fed once a week with powdered brewer's

Avoidance assays with *E. andrei*/*E. fetida* or *F. candida*, which are described by the guidelines ISO:17512-1 and ISO:17512-2 [41, 95], are performed using rectangular (earthworms) or round (collembolans) plastic boxes that are divided into two equal compartments by a plastic divider vertically introduced and filled with the test soil (amounts depend on the size of the container). Contaminated soil is added to one of the compartments, and the same amount of the respective control soil is added to the other compartment. Ten *E. andrei* or *E. fetida* adults (with developed clitellum) or 20 *F. candida* individuals at 10–12 days of age (originating from synchronized cultures) are then placed on the separation line between the two compartments, and the plastic divider is removed. The containers are then closed with perforated lids to allow air circulation. The animals are not fed during the test. After 48 hours, the number of individuals present in each compartment is recorded according to the specific method for each species [18, 96]. A double control test is also performed with control soil in both compartments to determine whether the organisms are randomly distributed between the two compartments in the

Chronic toxicity tests with *E. andrei*/*E. fetida* should be performed according to ISO:11268-2 [27]. Round plastic containers are filled with approximately 500 g soil (dry weight) treated with solutions with increasing contaminant concentrations (or dilutions of the naturally contami‐ nated soils) or the respective control soils. Ten adult earthworms (with visible clitellum) with individual weights between 300 and 600 mg that had been previously incubated in control soil for at least 24 hours are selected for each experimental unit. The containers are closed with perforated lids, and the earthworms are fed horse manure (≈5 g per replicate) at the beginning

and the rearing substrate should be periodically replaced.

transferred to new substrate every 2 months.

46 Invertebrates - Experimental Models in Toxicity Screening

yeast [30].

absence of contaminants.

Reproduction tests with *E. crypticus* are described in ISO:16387 [43]. Ten similar-sized adults (with visible clitellum) are placed in cylindrical containers containing 20 g soil (dry weight) that had been treated with the test substance or the respective control soil. Finely ground oat flakes can be supplied as food (≈2 mg per replicate), and the containers should be hermetically closed [83]. The containers are opened weekly to allow for gas exchange, and food and soil moisture are replenished as needed. Twenty-eight days following the beginning of the assay, the total number of enchytraeids are counted using a stereoscopic microscope following fixation in 80% ethanol, staining with rose bengal (1% in ethanol), and wet sieving of the organisms [94]. For *E. albidus*, the experimental procedures are somewhat different, especially with regard to the assay duration.

Tests evaluating the impact of contaminants on the reproduction of the collembolan *F. candida* are described in ISO:11267 [26]. Thirty grams of contaminated or control soil (fresh weight) are added to cylindrical containers (approximately 100 mL). Ten adult collembolans aged between 10 and 12 days (originating from synchronized cultures) are then placed into each experimental unit, and the containers are then hermetically closed. Food (dry yeast) is supplied at the beginning of the assay and on the 14th day, and the containers are opened weekly to allow for gas exchange. On the 28th day after the beginning of the assay, the soil of each replicate is submersed in water to force the survivors to float to the surface, and the juveniles are counted following the addition of several drops of black ink (for increased contrast) [93].

Reproduction tests with *H. aculeifer* are described in guideline OECD no. 226 [30]. The containers are filled with 20 g (fresh weight) of soil treated with the test substance or control soil. Ten females with ages between 28 and 35 days and originating from the synchronized cultures are then placed in each container. The animals receive small amounts of food (cheese mites) at the beginning of the assay and then twice a week until the end of the assay. The containers are then hermetically closed and opened weekly for airing and soil moisture adjustments. Fourteen days after the beginning of the assay, the mites are removed from the soil using a MacFadyen extractor with a gradient of increasing temperatures for 48 hours (12 hours at 25°C, 12 hours at 35°C, and 24 hours at 45°C). Adults and newly emerged juveniles in each replicate should be fixed in 70% ethanol and counted using a stereoscopic microscope.

The following parameters are used to evaluate the critical values: NOEC (no observed effect concentration) and LOEC (lowest observed effect concentration), LC10 and LC50 (lethal concentration to 10 and 50% of the population, respectively), EC10 and EC50 (concentration at which 10 and 50% of the contaminant's maximal effect is observed on the growth or number of juveniles, respectively), and AC50 (concentration causing the avoidance of 50% of the organisms from the contaminated soils). These toxicity parameters are designed to detect the ecological risk of the chemicals in soil and used to derive protection limits for terrestrial ecosystems.

The significance of avoidance responses (LOEC and NOEC) is tested with Fisher's exact test using a two-tailed test for the double control conditions and a one-tailed test for the contami‐ nated soil combination conditions [97]. The AC50 values can be obtained using regression analyses [98]. The significance of the effects (LOEC and NOEC) on the body biomass of the earthworms and number of earthworms, enchytraeids, mites, and collembolan juveniles following exposure to the contaminated soils should be tested using a one-way analysis of variance (ANOVA). When differences are detected (*p* ≤ 0.05), the treatment means should be compared with the results from the respective controls using a post hoc test such as Dunnett's test. The EC10 and EC50 values should be estimated by nonlinear regression using pre-defined exponential, logistic, Gompertz, or hormesis models [71]. The normality and homogeneity of variance should be tested prior to the ANOVAs.

#### **4. Future prospects in soil ecotoxicology**

The future prospects for soil ecotoxicology refer to the challenge of increasing the realism of the analyses in the terrestrial environment and to reduce the uncertainties about the real degree of ecological risk obtained by laboratory tests. To improve the ecological relevance of labora‐ tory assays, it needs a transition from research based on artificial soil to the use of natural soils, which consider the real relationship between contaminants and test organisms in the exposure scenarios. Moreover, it is necessary to increase the list of standard organisms for the tests, considering the practicality in assays, and especially the inclusion of species that represent the geographical and ecological conditions of the test site [99]. As an example, Ref. [20] suggests the standardization of the sublethal toxicity tests with isopods, since they represent an ecologically relevant group of soil fauna, and the effects on these arthropods can be determined at biochemical, genomic, individual (growth, behavior), and ecological (feeding activity) levels.

It is still necessary to move forward in the assessments of long-term sublethal effects; besides, there is a need for better understanding of exposure, absorption, and metabolism of substances in individuals, and the identification of the responses at different levels of biological organi‐ zation (e.g., communities) [20]. Based on this assumption, one of the apparent possibilities to evaluate the long-term impact in standard laboratory tests would be the use of multigenera‐ tional assays, where toxic effects such as delayed reproductive failures, transmission of the bioaccumulation to offspring, or accumulation of DNA damage could be identified [100]. For a better understanding of the relationships between pollutants and species, more studies using chemical, biochemical, and molecular prospects (ecotoxicogenomics) are needed, particularly assessments of bioavailability, bioaccumulation, and molecular biomarkers [20, 101]. The assessment of impacts at higher levels of biological organization can be accomplished through multispecies assays, which consider the relationships between species, contaminants, and soil properties. In addition, the semi-field and field tests may offer a better understanding of the contaminant's impacts on soil communities, although they are more complex, especially when there is the involvement of comparisons between different ecosystems [13, 20].

Finally, it is necessary that future adjustments be performed in the standard assays available, in order to enable them to address the new and emerging needs of the current ecotoxicology, such as the case of evaluation of the toxicity of nanoparticles and mixtures of contaminants, among others [20]. These adjustments also extend to the studies performed in tropical regions, where there is a need for a revision of the methods, especially in terms of soils, species, and climatic conditions, in order to increase the ecological relevance of the analyses at local level.

#### **Author details**

ecological risk of the chemicals in soil and used to derive protection limits for terrestrial

The significance of avoidance responses (LOEC and NOEC) is tested with Fisher's exact test using a two-tailed test for the double control conditions and a one-tailed test for the contami‐ nated soil combination conditions [97]. The AC50 values can be obtained using regression analyses [98]. The significance of the effects (LOEC and NOEC) on the body biomass of the earthworms and number of earthworms, enchytraeids, mites, and collembolan juveniles following exposure to the contaminated soils should be tested using a one-way analysis of variance (ANOVA). When differences are detected (*p* ≤ 0.05), the treatment means should be compared with the results from the respective controls using a post hoc test such as Dunnett's test. The EC10 and EC50 values should be estimated by nonlinear regression using pre-defined exponential, logistic, Gompertz, or hormesis models [71]. The normality and homogeneity of

The future prospects for soil ecotoxicology refer to the challenge of increasing the realism of the analyses in the terrestrial environment and to reduce the uncertainties about the real degree of ecological risk obtained by laboratory tests. To improve the ecological relevance of labora‐ tory assays, it needs a transition from research based on artificial soil to the use of natural soils, which consider the real relationship between contaminants and test organisms in the exposure scenarios. Moreover, it is necessary to increase the list of standard organisms for the tests, considering the practicality in assays, and especially the inclusion of species that represent the geographical and ecological conditions of the test site [99]. As an example, Ref. [20] suggests the standardization of the sublethal toxicity tests with isopods, since they represent an ecologically relevant group of soil fauna, and the effects on these arthropods can be determined at biochemical, genomic, individual (growth, behavior), and ecological (feeding activity)

It is still necessary to move forward in the assessments of long-term sublethal effects; besides, there is a need for better understanding of exposure, absorption, and metabolism of substances in individuals, and the identification of the responses at different levels of biological organi‐ zation (e.g., communities) [20]. Based on this assumption, one of the apparent possibilities to evaluate the long-term impact in standard laboratory tests would be the use of multigenera‐ tional assays, where toxic effects such as delayed reproductive failures, transmission of the bioaccumulation to offspring, or accumulation of DNA damage could be identified [100]. For a better understanding of the relationships between pollutants and species, more studies using chemical, biochemical, and molecular prospects (ecotoxicogenomics) are needed, particularly assessments of bioavailability, bioaccumulation, and molecular biomarkers [20, 101]. The assessment of impacts at higher levels of biological organization can be accomplished through multispecies assays, which consider the relationships between species, contaminants, and soil properties. In addition, the semi-field and field tests may offer a better understanding of the contaminant's impacts on soil communities, although they are more complex, especially when

there is the involvement of comparisons between different ecosystems [13, 20].

ecosystems.

levels.

variance should be tested prior to the ANOVAs.

48 Invertebrates - Experimental Models in Toxicity Screening

**4. Future prospects in soil ecotoxicology**

Paulo Roger Lopes Alves1\* and Elke Jurandy Bran Nogueira Cardoso2

\*Address all correspondence to<EMAIL_ADDRESS>
1 Federal University of South Border, Chapecó, Brazil

2 Luiz de Queiroz College of Agriculture -University of São Paulo, Piracicaba, Brazil

#### **References**

[21] Kapanen A, Itavaara M. Ecotoxicity tests for compost applications. Ecotoxicology and Environmental Safety. 2001;49:1–16.

[7] Moser H, Römbke J, editors. Ecotoxicological Characterization of Waste – Results and Experiences of an International Ring Test. Chichester: Wiley. 2009;p. 308 p.

[8] Parisi V, Menta C, Gardi C, Jacomini C, Mozzanica E. Microarthropod communities as a tool to assess soil quality and biodiversity: A new approach in Italy. Agriculture,

[9] Cardoso EJBN, Vasconcellos LR, Bini D, Yumi MM, Santos CA, Alves PRL, de Paula AM, Nakatani AS, Pereira JM, Nogueira MA. Soil health: Looking for suitable indica‐ tors. What should be considered to assess the effects of use and management on soil

[10] Schoenholtz SH, Vam Miegroet H, Burger JA. A review of chemical and physical properties as indicators of forest soil quality: Challenges and opportunities. Forest

[11] Everts JW, Aukema B, Hengeveld R, Koeman JH. Side-effects of pesticides on ground-dwelling predatory arthropods in arable ecosystems. Environmental Pollu‐

[12] Krogh PH. Microarthropods as bioindicators [thesis]. Arhus: University of Arhus;

[13] Cardoso EJBN, Alves PRL. Soil ecotoxicology. In: Begum G, editor. Ecotoxicology. Ri‐

[14] Melo LCA. Phytotoxicity of cadmium and barium and derivation of critical limits in

[15] Shugart LR, editor. Emerging Topics in Ecotoxicology: Principles, Approaches and

[16] EC. 284/2013 of 1 March 2013 Setting Out the Data Requirements for Plant Protection Products, in Accordance with Regulation (EC) No 1107/2009 of the European Parlia‐ ment and of the Council Concerning the Placing of Plant Protection Products on the

[17] EC. Directive 2008/98/EC: Directive of the European Parliament and of the Council of 19 November 2008 on Waste and Repealing Certain Directives. Official Journal of the

[18] Alves PRL, Cardoso EJBN, Martines AM, Sousa JP, Pasini A. Earthworm ecotoxico‐ logical assessments of pesticides used to treat seeds under tropical conditions. Che‐

[19] Méhu J, Bazin C, Grelier-Volatier L. Classification réglementaire et écocompatibilité

[20] Van Gestel CAM. Soil ecotoxicology: State of the art and future directions. ZooKeys.

Ecosystems & Environment. 2005;105:323–333.

health? Scientia Agricola. 2013;70:274–289.

Ecology and Management. 2000;138:335–356.

jeka: InTech – Open Access Publisher. 2012; pp. 27–50.

soils [thesis]. Piracicaba: University of São Paulo; 2010.

Perspectives. Oak Ridge: Springer Science; 2009. 400p.

des déchets. Techniques de l'ingénieur. 2004;G2-030:1–10.

tion. 1989;59:203–225.

50 Invertebrates - Experimental Models in Toxicity Screening

Market. J.O.L. 2013;93:1–84.

European Union. 2008;312:3–30.

mosphere. 2013;90:2674–2682.

2012;176:275–296.

1994.

[49] Dash MC. Oligochaeta: Enchytraeidae. In: Dindal DL, editor. Soil Biology Guide. New York: John Wiley & Sons; 1990. pp. 311–40.

[35] Heupel K. Avoidance response of different collembolan species to Betanal. European

[36] Langdon CJ, Hodson ME, Arnold RE, Black S. Survival, Pb-uptake and behavior of three species of earthworm in Pb treated soils determined using an OECD-style tox‐

icity test and a soil avoidance test. Environmental Pollution. 2005;138:368–375. [37] Ronday R, Houx NWH. Suitability of seven species of soil-inhabiting invertebrates for testing toxicity of pesticides in soil pore water. Pedobiologia. 1996;40:106–112. [38] Fountain MT, Hopkin SP. *Folsomia candida* (Collembola): A "Standard" soil arthro‐

[39] Van Gestel CAM, Léon CD, Van Straalen NM. Evaluation of soil fauna ecotoxicity tests regarding their use in risk assessment. In: Tarradellas J, Bitton G, Rossel D, edi‐

[40] ISO. Soil quality No. 11268-1: Effects of pollutants on earthworms (*Eisenia fetida*) – Part 1: Determination of acute toxicity using artificial soil substrate. Geneva: Interna‐

[41] ISO. Soil quality No. 17512-1: Avoidance Test for Determining the Quality of Soils and Effects of Chemicals on Behavior – Part 1: Test with Earthworms (*Eisenia fetida*

and *Eisenia andrei*). Geneva: International Standardization Organization; 2008.

[42] OECD. Guidelines for the Testing of Chemicals No. 207: Earthworm, Acute Toxicity Tests. Paris: Organization for Economic Co-operation and Development; 1984. [43] ISO. Soil quality No. 16387: Effects of Pollutants on Enchytraeidae (*Enchytraeus sp.*) – Determination of Effects on Reproduction and Survival. Geneva: International Stand‐

[44] Peijnenburg W, Capri E, Kula C, Liess M, Luttik R, Montforts M, Nienstedt K, Rombke J, Sousa JP, Jensen J. Evaluation of exposure metrics for effect assessment of soil invertebrates. Critical Reviews in Environmental Science and Technology.

[45] Edwards CA, editor. Earthworm Ecology. V. 3. Boca Raton: CRC Press LLC; 2004.

[46] Domínguez J, Ferreiro A, Velando A. Are *Eisenia fetida* (Savigny, 1826) and *Eisenia an‐ drei* Bouché, 1972 (Oligochaeta, Lumbricidae) different biological species? Pedobiolo‐

[47] Hassal L, Turner JG, Rands MRW. Effects of terrestrial isopods on the decomposition

[48] Didden WAM. Ecology of terrestrial Enchytraeidae. Pedobiologia. 1993;37:2–29

of woodland leaf litter. Oecologia. 1987;72:23–36.

tors. Soil Ecotoxicology. Boca Raton: CRC Press; 1997. pp. 291–317.

Journal of Soil Biology. 2002;38:273−276.

52 Invertebrates - Experimental Models in Toxicity Screening

pod. Annual Review of Entomology. 2005;50:201–22.

tional Standardization Organization; 1993.

ardization Organization; 2004.

2012;42:1862–1893.

gia. 2005;49:81–87.

438 p.

 Propensity towards cannibalism among *Hypo‐ aspis aculeifer* and *H. miles*, two soil-dwelling predatory mite species. Experimental and Applied Acarology. 2003;31(1–2):1–14.

[64] Culik, MP, Zeppelini D. Diversity and distribution of Collembola (Arthropoda: Hex‐

[65] Van Straalen NM. Assessment of soil contamination – a functional perspective. Bio‐

[66] Krogh PH. Toxicity Testing with the Collembolans *Folsomia fimetaria* and *Folsomia candida* and the Results of a Ring Test (trans: DMU/AU DoTE). Miljoministeriet; 2008.

[67] Crouau Y, Gisclard C, Perotti P. The use of *Folsomia candida* (Collembola, Isotomidae)

[68] Wiles JA, Krogh PH. Testing with the collembolans *I. viridis*, *F. candida* and *F. fimeta‐ ria*. In: Løkke H, Van Gestel CAM, editors. Handbook of Soil Invertebrate Toxicity

[69] Van Gestel CAM. The developmental status of ecotoxicity tests. Løkke H, Van Gestel CAM, editors. Handbook of Soil Invertebrate Toxicity Tests. Chichester: John Wiley

[70] Greenslade P, Vaughan GT. A comparison of Collembola species for toxicity testing

[71] Environment-Canada. Guidance Document on Statistical Methods for Environmental Toxicity Tests. Environmental Protection Series, EPS 1/RM/46, 2005 with 2007 up‐

[72] Huguier P, Manier N, Owojori OJ, Bauda P, Pandard P, Römbke J. The use of soil

[73] Mineiro JC, Moraes GJ. Gamasida (Arachnida: Acari) Edáficos de Piracicaba, Estado

[74] RömbkeJ, Beck L, Dreher P, Hund-Rinke K, Jänsch S, Kratz W, Pieper S, Ruf A, Spel‐ da J, Woas S. Entwicklung von bodenbiologischen Bodengüteklassen für Acker- und

[75] Smit CE, Moser T, Rombke J. A new OECD test guideline for the predatory soil mite *Hypoaspis aculeifer*: Results of an international ring test. Ecotoxicology and Environ‐

[76] Helyer N, Brown K, Cattlin ND. A Colour Handbook of Biological Control in Plant

[77] Krogh PH, Axelsen JA. Test on the predatory mite *Hypoaspis aculeifer* preying on the collembolan *Folsomia fimetaria*. In: Løkke H, Van Gestel CAM, editors. Handbook of Soil Invertebrate Toxicity Tests. Chichester: John Wiley & Sons; 1998. pp. 239–251.

mites in ecotoxicology: A review. Ecotoxicology. 2015;24(1):1–18.

de São Paulo. Neotropical Entomology. 2001;30(3):379–385.

Grünlandstandorte. UBA-Texte 20/02; 2002. 273 p.

Protection. London: Manson Publishing; 2003. 128 p.

mental Safety. 2012;82:56–62.

apoda) of Brazil. Biodiversity and Conservation. 2003;12:1119–1143.

in bioassays of waste. Applied Soil Ecology. 2002;19:65–70.

Tests. Chichester: John Wiley & Sons; 1998. pp. 131–156.

of Australian soils. Pedobiologia. 2003;47:171–179.

dates. Ottawa: Environment Canada; 2007.

degradation. 2002;13:41–52.

54 Invertebrates - Experimental Models in Toxicity Screening

& Sons; 1998.pp. 57–67.

44 p.

## **Rotifers as Models in Toxicity Screening of Chemicals and Environmental Samples**

Roberto Rico-Martínez, Mario Alberto Arzate-Cárdenas, Daniel Robles-Vargas, Ignacio Alejandro Pérez-Legaspi, Alvarado-Flores Jesús and Gustavo Emilio Santos-Medrano

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/61771

#### **Abstract**

[90] Pesaro M, Widmer F, Nicollier G, Zeyer J. Effects of freeze-thaw stress during soil storage on microbial communities and methidathion degradation. Soil Biology and

[91] Niemeyer JC, Moreira-Santos M, Ribeiro R, Da Silva EM, Sousa JP. Environmental risk assessment of a metal contaminated area in the tropics. Tier I: Screening phase.

[92] Natal-Da-Luz T, Moreira-Santos M, Ruepert C, Castillo LE, Ribeiro R, Sousa JP. Eco‐ toxicological characterization of a tropical soil after diazinon spraying. Ecotoxicolo‐

[93] Alves PRL, Cardoso EJBN, Martines AM, Sousa JP, Pasini A. Seed dressing pesticides on springtails in two ecotoxicological laboratory tests. Ecotoxicology and Environ‐

[94] Chelinho S, Domene X, Campana P, Andrés P, Römbke J, Sousa JP. Toxicity of phen‐ medipham and carbendazim to *Enchytraeus crypticus* and *Eisenia andrei* (Oligochaeta)

[95] ISO. Soil quality No. 17512-2: Avoidance Test for Determining the Quality of Soils and Effects of Chemicals on Behaviour – Part 2: Test with Collembolans (*Folsomia*

[96] Natal-Da-Luz T, Tidona S, Jesus B, Morais PV, Sousa JP. The use of sewage sludge as soil amendment. The need for an ecotoxicological evaluation. Journal of Soils and

[98] Sakuma M. Probit analysis of preference data. Applied Entomology and Zoology.

[99] Kuperman RG, Checkai RT, Vinicius M, Garcia B, Römbke J. State of the science and the way forward for the ecotoxicological assessment of contaminated land. Pesquisa

[100] Paumen ML, Steenbergen E, Kraak MHS, Van Straalen NM, Van Gestel, CAM. Multi‐ generation exposure of the springtail *Folsomia candida* to phenanthrene: From doseresponse relationships to threshold concentrations. Environmental Science and

[101] Filser J, Wiegmann S, Schröder B. Collembola in ecotoxicology — Any news or just

[97] Zar JH. Biostatistical Analysis. London: Prentice Hall International; 1999. 663 p.

in Mediterranean soils. Journal of Soils and Sediments. 2014;14:584–599.

*candida*). Geneva: International Standardization Organization; 2011.

Biochemistry. 2003;35:1049–1061.

56 Invertebrates - Experimental Models in Toxicity Screening

gy. 2012;21:2163–2176.

mental Safety. 2014;105:65–71.

Sediments. 2009;9(3):246–260.

Agropecuária Brasileira. 2009;1:811–824.

boring routine? Applied Soil Ecology. 2014;83:193–199.

Technology. 2008;42:6985–6990.

1998;33:339–347.

Journal of Soils and Sediments. 2010;10:1557–1571.

An important objective of aquatic ecotoxicology is to determine the effects of toxic com‐ pounds in organisms that play a central role in aquatic communities where rotifers have a large impact on several important ecological processes. The contribution of the rotifers to secondary production in many aquatic communities is substantial as they are often the larger fraction of zooplankton biomass at certain times of the year. In addition to the im‐ portance of their ecological roles in aquatic communities, the rotifers are attractive organ‐ isms for ecotoxicological studies by its short life cycles and rapid reproduction, their small size, and little volumes needed for culture and toxicity assays. The main end points used in ecotoxicological studies are mortality, reproduction, behavior, and biomarkers. Such parameters are included in international regulations from all over the world, where different species are used to evaluate the effect of environmental samples or chemical compounds. The high diversity of rotifers is an important issue because it can modify their relative susceptibility to toxicants. Thus, more studies are needed to know the rela‐ tions and mechanisms involved in clonal variation, sensitivity, and development, which can be all assessed by state-of-the-art procedures.

**Keywords:** aquatic toxicology, ecotoxicology, metal toxicity, acute toxicity, endocrine dis‐ ruption

#### **1. Introduction**

The analytical equipment can identify and quantify a chemical substance but not its toxicity in the organisms or the environment, which can be evaluated only in life organisms [1]. Toxicity testing in water samples assesses the concentration and exposure time of the chemical

© 2016 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

substances that produce an adverse effect in aquatic organisms, generating useful data for risk assessment. A toxicity test can be accepted by the scientific community if (a) it is capable to predict adverse effects for a variety of compounds in different organisms, (b) it must be replicable with statistical-based analysis, (c) its data must include adverse effects in a range of concentrations in real exposure times, (d) it must be useful to evaluate a risk, (e) it is economic and easy to perform, and (f) it is sensitive and realistic [2].

A model organism for a toxicity test must be abundant, native, or representative of the ecosystem that would be impacted, with ecological and economic importance; there must be a good knowledge of its basic biology that helps interpretation of data and be available for routine maintenance in the laboratory [2].

Toxicity tests have been developed for rotifers to assess many and diverse end points like (a) mortality, (b) reproduction, (c) production of amictic females, (d) cyst production, (e) proba‐ bility of extinction, (f) behavior, (g) ingestion rates, (h) swimming activity, (i) *in vivo* enzymatic activity, and (j) genetic expression, among others [3–5].

Most of the toxicity tests in rotifers are lethal tests that rely in mortality as the end point to calculate the LC50 value typically at 24 or 48 h without feeding [5]. These tests once standar‐ dized are the base to develop monitoring protocols to assess water quality [4, 5]. On the other hand, chronic toxicity tests assess sublethal parameters like behavioral, physiological, or reproductive alterations as the first responses to toxic substances [5, 6], which show high sensitivity in shorter periods of time. The rotifer species *Brachionus calyciflorus* and *Brachionus plicatilis* are among the species most frequently used for both lethal and chronic tests [7, 8].

#### **1.1. Phylum Rotifera**

Rotifers are aquatic or semiaquatic microscopic invertebrates with nearly 1850 species; they are unsegmented, pseudocoelomate, and bilaterally symmetric [4, 9, 10]. Nowadays, two classes are recognized: Pararotatoria (with the single order Seisonacea) and Eurotatoria, with two subclasses, Bdelloidea and Monogononta. The size of rotifers ranges from 50 to 2000 µm in length. Most of the species are free swimmers, but some species are fixed to some substratum [4]. Their morphology is of saccate type that is cylindrical with three easily recognizable regions: corona, trunk, and foot (Figure 1) [11].

#### **1.2. Ecological relevance**

Rotifers are cosmopolitans. They inhabit aquatic environments of the three types: marine, freshwater, and estuarine [9]. Most rotifers are freshwater, littoral with a few species truly planktonic. The few species that comprise the order Seisonacea and nearly 100 species of the subclass Monogononta are exclusively marine [4]. Species of the subclass Bdelloidea are found in freshwater ecosystems, lakes, temporary pools, interstitial water, soil, moss, and lichens [4, 12]. Rotifers are important in freshwater environments due to having one of the highest reproductive rate among metazoans, thus obtaining high population densities in short times, being dominant in many zooplanktonic communities. They act as links between the microbial community and the higher trophic levels. Rotifers colonize habitats quickly and convert Rotifers as Models in Toxicity Screening of Chemicals and Environmental Samples http://dx.doi.org/10.5772/61771 59

**Figure 1.** Diagram of the freshwater rotifer *Brachionuscalyciflorus* (Monogononta) [11].

primary production (algae and cyanobacteria) in a usable form for secondary consumers, making energy available for the next trophic levels. In interstitial water from swampy soils, they contribute to nutrient recycling [4]. Some bdelloids and Monogononta are abundant in wastewater treatment plants as part of activated sludge, in filtration systems, or residual lagoons feeding on the bacterial biomass [2, 13]. The population dynamics of rotifers is well characterized both in field as in the laboratory; for this reason, they are useful to investigate ecological and demographic principles. Besides, they are frequently used to assess aquatic toxicity from a population point of view. The intrinsic growth rate (*r*) is an end point commonly used and has proved to be highly sensitive [5, 14].

#### **1.3. Culture**

substances that produce an adverse effect in aquatic organisms, generating useful data for risk assessment. A toxicity test can be accepted by the scientific community if (a) it is capable to predict adverse effects for a variety of compounds in different organisms, (b) it must be replicable with statistical-based analysis, (c) its data must include adverse effects in a range of concentrations in real exposure times, (d) it must be useful to evaluate a risk, (e) it is economic

A model organism for a toxicity test must be abundant, native, or representative of the ecosystem that would be impacted, with ecological and economic importance; there must be a good knowledge of its basic biology that helps interpretation of data and be available for

Toxicity tests have been developed for rotifers to assess many and diverse end points like (a) mortality, (b) reproduction, (c) production of amictic females, (d) cyst production, (e) proba‐ bility of extinction, (f) behavior, (g) ingestion rates, (h) swimming activity, (i) *in vivo* enzymatic

Most of the toxicity tests in rotifers are lethal tests that rely in mortality as the end point to calculate the LC50 value typically at 24 or 48 h without feeding [5]. These tests once standar‐ dized are the base to develop monitoring protocols to assess water quality [4, 5]. On the other hand, chronic toxicity tests assess sublethal parameters like behavioral, physiological, or reproductive alterations as the first responses to toxic substances [5, 6], which show high sensitivity in shorter periods of time. The rotifer species *Brachionus calyciflorus* and *Brachionus plicatilis* are among the species most frequently used for both lethal and chronic tests [7, 8].

Rotifers are aquatic or semiaquatic microscopic invertebrates with nearly 1850 species; they are unsegmented, pseudocoelomate, and bilaterally symmetric [4, 9, 10]. Nowadays, two classes are recognized: Pararotatoria (with the single order Seisonacea) and Eurotatoria, with two subclasses, Bdelloidea and Monogononta. The size of rotifers ranges from 50 to 2000 µm in length. Most of the species are free swimmers, but some species are fixed to some substratum [4]. Their morphology is of saccate type that is cylindrical with three easily recognizable

Rotifers are cosmopolitans. They inhabit aquatic environments of the three types: marine, freshwater, and estuarine [9]. Most rotifers are freshwater, littoral with a few species truly planktonic. The few species that comprise the order Seisonacea and nearly 100 species of the subclass Monogononta are exclusively marine [4]. Species of the subclass Bdelloidea are found in freshwater ecosystems, lakes, temporary pools, interstitial water, soil, moss, and lichens [4, 12]. Rotifers are important in freshwater environments due to having one of the highest reproductive rate among metazoans, thus obtaining high population densities in short times, being dominant in many zooplanktonic communities. They act as links between the microbial community and the higher trophic levels. Rotifers colonize habitats quickly and convert

and easy to perform, and (f) it is sensitive and realistic [2].

activity, and (j) genetic expression, among others [3–5].

regions: corona, trunk, and foot (Figure 1) [11].

routine maintenance in the laboratory [2].

58 Invertebrates - Experimental Models in Toxicity Screening

**1.1. Phylum Rotifera**

**1.2. Ecological relevance**

Rotifers can be obtained directly from a natural aquatic system or from laboratory cultures. Rotifers can be hatched from cysts, and today there are toxicity kits of two species: *B. calyci‐ florus* and *B. plicatilis* (Rotoxkits). The culture of rotifers has been developed for freshwater, estuarine, and marine organisms taken directly from the natural environment after a previous filtration, sterilization, and neutralization are performed. There is also synthetic hard like EPA medium [15] or synthetic marine water (Instant Ocean) that can be adjusted to the desired salinity. In freshwater species, usually the pH level is maintained in the physiological range (6.5–8) and temperature oscillates between 20 and 30°C. Marine species are maintained at salinities of 10 to 20 psu depending on a particular strain [16]. Rotifers are mainly filter feeders on microalgae, bacteria, or detritus; a few species are predatory. The supply of fresh and optimal food is the main problem for culturing rotifers. However, numerous rotifer species have been kept in the laboratory routinely [4]. Among the main marine microalgae used to feed rotifers are as follows: *Nannochloropsis* sp., *Chaetoceros* sp., *Dunaliella* sp., *Pyramimonas*, *Isochrysis* sp., and *Tetraselmis* sp.; for freshwater: *Nannochloris* sp., *Nannochloropsis* sp., and *Chlorella* sp. [16]. Culture media have to be renewed 2–3 times a week, and the supply of food must be frequent. Some species are maintained on diluted suspensions of commercial fish food, grain extracts or infusions, manure, and soil [4]. Among the most cultured species are found those of the genus *Brachionus*: *B. plicatilis*, *B. rotundiformis*, and *B. calyciflorus* [17, 18]. Some species of the genus *Lecane* can be easily cultured [19]. The economic importance of these rotifer species that are mass cultured are mainly based in their use as live feed for fish larvae and crustaceans in aquaculture [18].

#### **1.4. Reproduction**

The life cycles of rotifers are shorter than many other animals [14]. The Bdelloidea are repro‐ duced by exclusive parthenogenesis and males are unknown. In the subclass Monogononta, the life cycle is haplodiploid with cyclic parthenogenesis as the dominant phase (amictic females), whereas the sexual reproduction (mixis) involves mictic diploid females that produce haploid eggs that if fertilized produce diploid embryos or cysts (in old literature they are called "resting eggs"), which go through diapause before hatching into amictic females that go back to the asexual phase of the cycle (Figure 2). Mixis is triggered by specific environmental stimuli like high population density or photoperiod [18, 20]. Parthenogenesis eliminates the high cost of producing males resulting in a rapid growth population [18], which allows for the high production of clonal individuals that can be used for aquaculture or toxicity tests.

**Figure 2.** Reproductive cycle of *Brachionus calyciflorus*. Modified from Alvarado-Flores [21].

#### **1.5. Cysts**

on microalgae, bacteria, or detritus; a few species are predatory. The supply of fresh and optimal food is the main problem for culturing rotifers. However, numerous rotifer species have been kept in the laboratory routinely [4]. Among the main marine microalgae used to feed rotifers are as follows: *Nannochloropsis* sp., *Chaetoceros* sp., *Dunaliella* sp., *Pyramimonas*, *Isochrysis* sp., and *Tetraselmis* sp.; for freshwater: *Nannochloris* sp., *Nannochloropsis* sp., and *Chlorella* sp. [16]. Culture media have to be renewed 2–3 times a week, and the supply of food must be frequent. Some species are maintained on diluted suspensions of commercial fish food, grain extracts or infusions, manure, and soil [4]. Among the most cultured species are found those of the genus *Brachionus*: *B. plicatilis*, *B. rotundiformis*, and *B. calyciflorus* [17, 18]. Some species of the genus *Lecane* can be easily cultured [19]. The economic importance of these rotifer species that are mass cultured are mainly based in their use as live feed for fish larvae and

The life cycles of rotifers are shorter than many other animals [14]. The Bdelloidea are repro‐ duced by exclusive parthenogenesis and males are unknown. In the subclass Monogononta, the life cycle is haplodiploid with cyclic parthenogenesis as the dominant phase (amictic females), whereas the sexual reproduction (mixis) involves mictic diploid females that produce haploid eggs that if fertilized produce diploid embryos or cysts (in old literature they are called "resting eggs"), which go through diapause before hatching into amictic females that go back to the asexual phase of the cycle (Figure 2). Mixis is triggered by specific environmental stimuli like high population density or photoperiod [18, 20]. Parthenogenesis eliminates the high cost of producing males resulting in a rapid growth population [18], which allows for the high

production of clonal individuals that can be used for aquaculture or toxicity tests.

**Figure 2.** Reproductive cycle of *Brachionus calyciflorus*. Modified from Alvarado-Flores [21].

crustaceans in aquaculture [18].

60 Invertebrates - Experimental Models in Toxicity Screening

**1.4. Reproduction**

Once the eggs are produced, they fall into the bottom and are deposited in the sediments [18]. The cysts are diploid and have a thick cover and can be viable for many years in dormancy. Cysts are very resistant to harsh environmental conditions like draught and freezing. Cysts can be dispersed through wind, water, or migratory animals. Under favorable conditions in a specific habitat, the cysts respond to a specific environmental clue (photoperiod, temperature, and salinity) and starts hatching producing an amictic diploid female [4]. Cysts can be stored for great periods of time without losing viability having rotifers in the desired period of time [22]. The use of rotifers hatched from cysts to develop toxicity tests was introduced by Snell and Persoone [23]. Nowadays, it is possible to obtain cysts from marine (*B. plicatilis*) and freshwater (*B. calyciflorus*) species in the market.

#### **1.6. Rotifers as sentinels or bioindicators**

Aquatic invertebrates are attractive model organisms in aquatic toxicology due to their short generation time compared with fishes besides their small size require small test volumes [14, 24]. Their small size, the fact that they are filter feeders, and the permeability of the integument made rotifers quite susceptible to chemical and physical changes [19, 25]. Due to the impor‐ tance of rotifers in the aquatic trophic webs, they are useful as sentinel species to indicate toxicant exposure in affected ecosystems [4].

The knowledge of the (a) basic biology of rotifers, (b) sensitivity to contaminants at the physiological and demographic levels, (c) cosmopolitan distribution, (d) great ecological relevance, (e) high growth rate, (f) availability of neonates, (g) high ingestion rates, (h) ease of culture and handling, (i) transparency, (j) short life cycles, (k) ease to obtain clonal individuals, and (l) cyst production makes rotifers useful tools for assessment of aquatic ecosystems [4, 5, 8, 17, 18, 22, 24, 26].

#### **2. Rotifers as model organism among invertebrates**

Invertebrates are the most widely distributed organisms on the Earth and consist of a large and very diverse group, consisting in more than 30 phyla, several of which include more than 1000 different species [27]. The largest phylum within invertebrates is the Arthropoda, with more than one million species, in which insects and crustaceans are the two largest groups [28]. Aquatic crustaceans is a larger group in comparison to the aquatic insects, which a few of them have aquatic larvae [29]. Consequently, crustaceans are the most numerous and ecologically important group of invertebrates in marine and freshwater ecosystems, playing an important role in regulatory toxicity testing, whether in field or laboratory conditions [30, 31]. In addition, these methods are cost-effective because of the facility to obtain amictic eggs or in some cases resting eggs, which have been used to produce commercial kits for toxicity evaluation [32].

Current international protocols include several invertebrates' taxa for toxicity assessment (Table 1), in which cladocerans are the most used organisms and have more approved protocols for both acute (lethal) and (sub)chronic (sublethal) water toxicity assays than other taxa. The genus *Daphnia* is the most studied taxon within cladocerans. For the acute toxicity test, three replicates for at least five toxicant or effluent concentrations are required, every one of them with 100 mL, 48 h for exposure, without feeding [33]. Moreover, the chronic toxicity assay needs almost the same conditions but with feeding and daily medium renewal for 21 days [34]. In comparison, the *Brachionus* or other rotifer bioassays need only 1 mL per replicate [23], and the results can be obtained within 2 days for acute toxicity evaluation, and in 5 days for the chronic ones. Hence, rotifers tests involve similar conditions to the cladocerans, but when time is a matter of concern, the former is the best option to know the toxicity of chemicals or field samples.

Rotifers are even more versatile than daphnids. In April 2010, after the Macondo oil spill in the Gulf of Mexico, the Environmental Protection Agency of the United States (USEPA) in order to assess the impact of petroleum and oil dispersants on the aquatic biota recommended using *B. plicatilis,* a species complex that included *B manjavacas*, *B. plicatilis sensu stricto*, and *B. rotundiformis* among other that can exist [35]. This cost-effective test can be replicated hundreds of times in very short periods of time when necessary [36].

The Organization for the Economic Cooperation and Development (OECD) encourages developing test protocols that include crustaceans [37]. In this concern, the copepod *Eucyclops serrulatus* has been proposed as a model organism, with acute toxicity evaluations in 96 h [38] and chronic toxicity assessment in about 60 days [39]. These periods differ due to intrinsic characteristics of the species; copepods last longer to reach sexual maturity after hatching and require mating to obtain eggs for assays, while rotifer eggs are produced by parthenogenesis [40–42]. Nevertheless, with regard to the importance of copepods in aquatic ecosystems, there is not yet any approved protocol for toxicity evaluation as in rotifers [43, 44].

The cnidarian *Hydra attenuta* is another model organism included in standard methods [45]. This one is particularly interesting since lethal and sublethal end points are assessed in the same assay by quantifying the number of death animals and abnormalities found in the survivor hydras during 96 h for acute procedures, which are performed in 12-well microplates, whit a test volume of 4 mL [46]. Clubbed tentacles appearance is the first sign of abnormalities due to toxicants exposure. From this point, abnormalities follow shortening tentacles and the tulip stage (all tentacles have disappeared), and finally the disintegration of animal bodies occurs. Therefore, lethal effects should be registered from the tulip stage [47].

It is worth to compare with some rotifers in which it is also possible to determine sublethal effects in the same time period. Alvarado-Flores et al. [48] found abnormal body shapes in *B. calyciflorus* after vinclozolin exposure, demonstrating this pesticide also acts as an endocrine disruptor. As result, both models can be used for sublethal toxicity screening in short periods of time, but the difference lies on the apparent ease to determine such effects, while in *Hydra attenuata* are very obvious. In *B. calyciflorus*, more than one thousand organisms were analyzed to find statistically abnormalities in this rotifer species.

Bivalves in ecotoxicology procedures refer the use of species such as *Lampsilis siliquoidea*, *Corbicula fluminea, Leptodea fragilis*, *Ligumia subrostrata*, and *Megalonaias nervosa*, among others [49, 50]. In the scientific literature, most of the bivalve specimens used in toxicological research and evaluations have been collected from environmental samples or commercial producers [51–54]. Glochida are dissected from adult bivalves and need a fish host to develop into juveniles [55–57]. Both immature stages, glochida and juveniles, are used for water quality assessment or chemicals released into aquatic systems [58]. In contrast, rotifer experimental organisms are produced in laboratory-controlled conditions, and animals are ready to use a few hours after hatching instead for several weeks to have juvenile unionids. Hence, rotifer bioassays are less time consuming than bivalves protocols.

protocols for both acute (lethal) and (sub)chronic (sublethal) water toxicity assays than other taxa. The genus *Daphnia* is the most studied taxon within cladocerans. For the acute toxicity test, three replicates for at least five toxicant or effluent concentrations are required, every one of them with 100 mL, 48 h for exposure, without feeding [33]. Moreover, the chronic toxicity assay needs almost the same conditions but with feeding and daily medium renewal for 21 days [34]. In comparison, the *Brachionus* or other rotifer bioassays need only 1 mL per replicate [23], and the results can be obtained within 2 days for acute toxicity evaluation, and in 5 days for the chronic ones. Hence, rotifers tests involve similar conditions to the cladocerans, but when time is a matter of concern, the former is the best option to know the toxicity of chemicals

Rotifers are even more versatile than daphnids. In April 2010, after the Macondo oil spill in the Gulf of Mexico, the Environmental Protection Agency of the United States (USEPA) in order to assess the impact of petroleum and oil dispersants on the aquatic biota recommended using *B. plicatilis,* a species complex that included *B manjavacas*, *B. plicatilis sensu stricto*, and *B. rotundiformis* among other that can exist [35]. This cost-effective test can be replicated hundreds

The Organization for the Economic Cooperation and Development (OECD) encourages developing test protocols that include crustaceans [37]. In this concern, the copepod *Eucyclops serrulatus* has been proposed as a model organism, with acute toxicity evaluations in 96 h [38] and chronic toxicity assessment in about 60 days [39]. These periods differ due to intrinsic characteristics of the species; copepods last longer to reach sexual maturity after hatching and require mating to obtain eggs for assays, while rotifer eggs are produced by parthenogenesis [40–42]. Nevertheless, with regard to the importance of copepods in aquatic ecosystems, there

The cnidarian *Hydra attenuta* is another model organism included in standard methods [45]. This one is particularly interesting since lethal and sublethal end points are assessed in the same assay by quantifying the number of death animals and abnormalities found in the survivor hydras during 96 h for acute procedures, which are performed in 12-well microplates, whit a test volume of 4 mL [46]. Clubbed tentacles appearance is the first sign of abnormalities due to toxicants exposure. From this point, abnormalities follow shortening tentacles and the tulip stage (all tentacles have disappeared), and finally the disintegration of animal bodies

It is worth to compare with some rotifers in which it is also possible to determine sublethal effects in the same time period. Alvarado-Flores et al. [48] found abnormal body shapes in *B. calyciflorus* after vinclozolin exposure, demonstrating this pesticide also acts as an endocrine disruptor. As result, both models can be used for sublethal toxicity screening in short periods of time, but the difference lies on the apparent ease to determine such effects, while in *Hydra attenuata* are very obvious. In *B. calyciflorus*, more than one thousand organisms were analyzed

Bivalves in ecotoxicology procedures refer the use of species such as *Lampsilis siliquoidea*, *Corbicula fluminea, Leptodea fragilis*, *Ligumia subrostrata*, and *Megalonaias nervosa*, among others

is not yet any approved protocol for toxicity evaluation as in rotifers [43, 44].

occurs. Therefore, lethal effects should be registered from the tulip stage [47].

to find statistically abnormalities in this rotifer species.

of times in very short periods of time when necessary [36].

or field samples.

62 Invertebrates - Experimental Models in Toxicity Screening

In freshwater sediment, toxicity screening several protocols are currently carried out by international regulation agencies (Table 1). Nevertheless, rotifers are not included as test species, but it has been proved that the genus *Lecane* is suitable for those evaluations, repre‐ senting a cost-effective and high reliable alternative to the standardized methods.

Note: Numbers in square brackets indicate reference number

**Table 1.** Toxicity evaluations protocols using different invertebrates for freshwater and freshwater sediments, seawater, and seawater sediments.

Differently to the protocols mentioned above, all those for seawater and seawater sediments are found in standardized probes approved by agencies such as USEPA, ASTM, and the OECD (Table 1). As previously described, *B. plicatilis* is a marine rotifer used in seawater toxicology evaluation, an although it is not included in the USEPA guidelines, the agency recommended its used based on the ASTM method [43]. For sediments, the prime option is the polychaetes such as *Arenicola marina*, mainly collected from beach sand in different locations around the world. The protocol involves an exposure time of about 28 days using artificial sediment and field sediments samples [59–61]. Moreover, *B. plicatilis* has been exposed to interstitial water from littoral ecosystems; although it is sensitive to bioavailable toxicants in pore water, it does not respond in the same manner that those organisms that live in the sediment.

Afterward, the use of rotifers in all the above-mentioned matrices is possible due to some advantages, such as their relative small size, their simple organization, their short life cycles that permit multigenerational studies in very short time periods, their reproduction through parthenogenesis, and their genetic homogeneity that leads to almost identical offspring [22]. Despite of these characteristics, there is still a need to continue researching the effect of toxicants and to improve protocols for rotifers in (eco)toxicology to better understand their physiology and how external factors alter rotifers normal responses, since studies with rotifers are not as numerous as other taxa (Figure 3).

**Figure 3.** Number of publications for the groups described in the text. The databases consulted were Elsevier (www.sciencedirect.com), Springer (www.link.springer.com), and Wiley Online Library (http://onlineli‐ brary.wiley.com). Query terms were the genera name and the word "toxicity". Pie chart data represent percentages based on published material.

#### **3. Rotifer sensitivity**

**Class Order Family Species**

Crustaceans Malacostraca Amphipoda Hyalellidae *Hyallela azteca* [109, 112]

Rotifera Bdelloidea Philodinidae *Philodina roseola* [118]

Rotifera Monogononta Ploima Brachionidae *Brachionus plicatilis* [43] Echinodermata Echinoidea Arbacioida Arbaciidae *Arbacia punctulata* [122] Arthropoda Malacostraca Mysida Mysidae *Mysidopsis bahia* [123] Copepoda Maxillopoda Calanoida Acartiidae *Acartia tonsa* [44, 124]

Molluscs Bivalvia Mytiloida Mytilidae *Mytilus edulis* [125, 126]

Annelida Polychaeta Capitellida Arenicolidae *Arenicola marina* [128]

**Table 1.** Toxicity evaluations protocols using different invertebrates for freshwater and freshwater sediments,

not respond in the same manner that those organisms that live in the sediment.

Differently to the protocols mentioned above, all those for seawater and seawater sediments are found in standardized probes approved by agencies such as USEPA, ASTM, and the OECD (Table 1). As previously described, *B. plicatilis* is a marine rotifer used in seawater toxicology evaluation, an although it is not included in the USEPA guidelines, the agency recommended its used based on the ASTM method [43]. For sediments, the prime option is the polychaetes such as *Arenicola marina*, mainly collected from beach sand in different locations around the world. The protocol involves an exposure time of about 28 days using artificial sediment and field sediments samples [59–61]. Moreover, *B. plicatilis* has been exposed to interstitial water from littoral ecosystems; although it is sensitive to bioavailable toxicants in pore water, it does

Note: Numbers in square brackets indicate reference number

64 Invertebrates - Experimental Models in Toxicity Screening

seawater, and seawater sediments.

Artropoda Malocostraca Amphipoda Corophiidae *Leptocheirus plumulosus[110, 129,*

Arthopoda Insecta Diptera Chironomidae *Chironomus dilutus* [109, 110,

Annelida Clitellata Lumbriculida Lumbriculidae *Lumbriculus variegatus* [113-115]

**Freshwater sediments**

**Seawater**

Harpacticoida Miraciidae *Amphiascus tenuiremis* [44]

Ostreoida Osteridae *Crassostrea gigas* [127]

**Seawater sediments**

*Stylodrilus heringianus* [116, 117]

*C. riparius* [109, 110, 119-121]

119-121]

*130]*

The toxicity of a wide variety of chemicals including organic compounds, metals, and phar‐ maceuticals that can be found in water reservoirs as a result of human activity has been tested by performing toxicity tests using rotifers as bioindicators as these organisms play an impor‐ tant ecological role in aquatic environments [3, 131]. Rotifers of the genus *Brachionus* can be found between the most commonly used tests organisms [132]; moreover, several rotifers are included for comparison purposes.

A test organism with high sensitivity to detect adverse effects of xenobiotics is always desirable; nevertheless, differences in sensitivity among species can always be found. Gener‐ ally, these are due to the biological characteristics of test organisms, the type of chemical, and its mode of action; thus, one species might be very sensitive to one toxic but not that sensitive to another compound [133]. This section focuses on the sensitivity of different rotifers, all of them from freshwater ecosystems, with exception of the marine species *B. plicatilis* and *Brachionus koreanus*, when testing the toxicity of pesticides, organic compounds, and metals attending their toxicological importance and the availability of literature.

Some examples of these variations on sensitivity are shown in Table 2. In relation to pesticides, malathion (organophosphorus) displays the lower toxicity and has a toxicity range from 33.72 to 45.5 mg/L, where the lower toxicity level is reported to *B. calyciflorus* and the higher to *B. plicatilis* [134, 135].

Lindane, an organochloride pesticide, and methyl parathion (organophosphorus) show moderate toxicity and have wider toxicity ranges in comparison to malathion. For the orga‐ nochloride, toxicity range is given by *B. koreanus* and *B. plicatilis* (14 and 35.89 mg/L, respec‐ tively). In the case of methyl parathion, which shows higher acute toxicity, a range from 0.607 mg/L for *Euchlanis dilatata* to 29.19 mg/L for *B. calyciflorus* was found. Another organophos‐ phorus of ecotoxicological relevance, chlorpyrifos, tends to affect significantly aquatic invertebrates due the LC50 values reported for *B. koreanus* (3.9 mg/L) and *B. calyciflorus* (11.85 mg/L) [96, 134, 136, 137].

The organochlorides endosulfan and pentachlorophenol (PCP) stand as highly toxic agro‐ chemicals. For endosulfan, *B. koreanus* and *B. plicatilis* correspond to the reference organisms to establish the range because of their 24-h LC50 values found in literature [135, 136]. For PCP, the lowest and highest toxicity levels were noted for *B. calyciflorus* [138, 139].

On literature, little information is available about the toxicity of other organic compounds such as benzene, toluene, xylene, and hexane. Nevertheless, Ferrando and Andreu-Moliner [7] and Pérez-Legaspi et al. [95] have obtained some toxicological data (Table 2). For xylene, hexane, and toluene, *B. calyciflorus* and *B. plicatilis* are the reference organisms, where the first was most sensitive for the three chemicals [7, 95]. For benzene, rotifers from the genus *Lecane* resulted being more sensitive, but also a wide toxicity range is registered from 6.97 for *Lecane luna* to 3762 mg/L for *Lecane hamata* [95].

The lowest LC50 values, and thus, more toxic metals, were reported for zinc, copper, and mercury. Besides, a relatively not wide toxicity range can be observed for these three chemicals (Table 3). The most sensitive test organisms in these cases were *Lecane quadridentata*, *B. plicatilis*, and *B. calyciflorus*, respectively, while the more resistant were *Brachionus havanaen‐ sis* (Zn), *P. acuticornis* (Cu), and *L. hamata* (Hg) [93, 95, 135, 140, 143, 144].

Moreover, it is important to mention that there are physical factors that can alter the sensitivity of a test organism when performing a toxicity test. One example is given by Preston et al. [138],

Note: Numbers in square brackets indicate reference number

\*48h LC50 value

found between the most commonly used tests organisms [132]; moreover, several rotifers are

A test organism with high sensitivity to detect adverse effects of xenobiotics is always desirable; nevertheless, differences in sensitivity among species can always be found. Gener‐ ally, these are due to the biological characteristics of test organisms, the type of chemical, and its mode of action; thus, one species might be very sensitive to one toxic but not that sensitive to another compound [133]. This section focuses on the sensitivity of different rotifers, all of them from freshwater ecosystems, with exception of the marine species *B. plicatilis* and *Brachionus koreanus*, when testing the toxicity of pesticides, organic compounds, and metals

Some examples of these variations on sensitivity are shown in Table 2. In relation to pesticides, malathion (organophosphorus) displays the lower toxicity and has a toxicity range from 33.72 to 45.5 mg/L, where the lower toxicity level is reported to *B. calyciflorus* and the higher to *B.*

Lindane, an organochloride pesticide, and methyl parathion (organophosphorus) show moderate toxicity and have wider toxicity ranges in comparison to malathion. For the orga‐ nochloride, toxicity range is given by *B. koreanus* and *B. plicatilis* (14 and 35.89 mg/L, respec‐ tively). In the case of methyl parathion, which shows higher acute toxicity, a range from 0.607 mg/L for *Euchlanis dilatata* to 29.19 mg/L for *B. calyciflorus* was found. Another organophos‐ phorus of ecotoxicological relevance, chlorpyrifos, tends to affect significantly aquatic invertebrates due the LC50 values reported for *B. koreanus* (3.9 mg/L) and *B. calyciflorus* (11.85

The organochlorides endosulfan and pentachlorophenol (PCP) stand as highly toxic agro‐ chemicals. For endosulfan, *B. koreanus* and *B. plicatilis* correspond to the reference organisms to establish the range because of their 24-h LC50 values found in literature [135, 136]. For PCP,

On literature, little information is available about the toxicity of other organic compounds such as benzene, toluene, xylene, and hexane. Nevertheless, Ferrando and Andreu-Moliner [7] and Pérez-Legaspi et al. [95] have obtained some toxicological data (Table 2). For xylene, hexane, and toluene, *B. calyciflorus* and *B. plicatilis* are the reference organisms, where the first was most sensitive for the three chemicals [7, 95]. For benzene, rotifers from the genus *Lecane* resulted being more sensitive, but also a wide toxicity range is registered from 6.97 for *Lecane luna* to

The lowest LC50 values, and thus, more toxic metals, were reported for zinc, copper, and mercury. Besides, a relatively not wide toxicity range can be observed for these three chemicals (Table 3). The most sensitive test organisms in these cases were *Lecane quadridentata*, *B. plicatilis*, and *B. calyciflorus*, respectively, while the more resistant were *Brachionus havanaen‐*

Moreover, it is important to mention that there are physical factors that can alter the sensitivity of a test organism when performing a toxicity test. One example is given by Preston et al. [138],

the lowest and highest toxicity levels were noted for *B. calyciflorus* [138, 139].

*sis* (Zn), *P. acuticornis* (Cu), and *L. hamata* (Hg) [93, 95, 135, 140, 143, 144].

attending their toxicological importance and the availability of literature.

included for comparison purposes.

66 Invertebrates - Experimental Models in Toxicity Screening

*plicatilis* [134, 135].

mg/L) [96, 134, 136, 137].

3762 mg/L for *Lecane hamata* [95].

In the case of inorganic toxicants (Table 3), lead appears as the "less" toxic and with the wider toxicity range from 0.035 for *E. dilatata* to 56.2 mg/L for *Philodina acuticornis* [97, 140]. For cadmium, *E. dilatata* shows a great sensitivity with a 48 h LC50 of 0.014 mg/L, while *B. plicatilis* is more resistant (39 mg/L) [97, 141]. The most sensitive rotifer to chromium was *Lecane luna* (48-h LC50 of 3.26 mg/L) [95], and the highest 24-h LC50 of 17.4 mg/L was registered for *B. calyciflorus* [142].

**Table 2.** Toxicity range for some pesticides and organic compounds to rotifers used in bioassays.

Note: Numbers in square brackets indicate reference number

**Table 3.** Toxicity range for different metals to rotifers used in bioassays.

where a decrease in LC50 values for PCP and mercury using *B. calyciflorus* as bioindicator was recorded as UV-B exposure time was increased.

Other physical condition that might modify the sensitivity of a test organism is desiccation. Robles-Vargas and Snell [145] found a remarkable resistance of *Philodina sp*. emerged from desiccation to PCP, chlorpyrifos, and mercury in comparison to continuously hydrated rotifers when reproductive tests were implemented [145].

According to the information reviewed and in terms of acute toxicity, rotifers from the family Brachionidae tend to show a great sensitivity to different chemicals, including organic and inorganic compounds. Invertebrates from the family Euchlanidae and Lecanidae appear to be sensitive mainly to inorganic toxicants. Furthermore, rotifers belonging to the family Philodi‐ nidae exhibit a notorious resistance to chemicals.

#### **4. Rotifer database for screening toxicants**

Every chemical product that can reach ecosystems and, as a consequence, the human being needs to be characterized to avoid noxious effects on all organisms exposed. At first instance, toxicological studies were mainly anthropocentrically focused, but that point of view would not have been enough to protect our own existence at all if affecting organisms in different ecosystems would have continued. It is known that rotifers play an important role as secondary producers and in some freshwater ecosystems represent the larger fraction of biomass of zooplankton [146]. Therefore, these organisms are nowadays considered as model organisms for toxicological screening.

Currently, one can find in the scientific literature reports of almost every class of chemical compound used or produced by man, investigating their toxicological properties. These papers are relatively abundant in several websites specialized in gathering such kind of information, which is available to everyone who has access to the published material. For scientist, collecting this material is a daily activity that guarantees their studies to follow the right direction by not working in exactly the same topic and manner, which would represent the duplicity of information but at higher cost because of multiple double efforts and resources investment. Nonetheless, it must be pointed out that differences in susceptibility to toxicants can be found among clones or strains, which is very interesting to assess by multiclonal analyses [22, 147]. However, when it is not possible to access multiple clones or strains, previous reports (published material) from databases become more relevant as a source of comparison to improve the comprehension of the observed results.

The USEPA, in an attempt to ease the search for environmental toxicological data, has created the ECOTOX database, hosted in their website [150]. It provides a user-friendly interface and allows discrimination among taxonomic and chemical groups, aquatic or terrestrial environ‐ ment, tests results or conditions, and type (year) of publication. Although it does not give access to the cited references, it returns a list of publications to be consulted elsewhere. In the ECOTOX results table, the data shown are species name, exposure type, chemical, media type, end points, bioconcentration factor, effects, and statistics, among others (Figure 4). A complete list of all abbreviations used in the ECOTOX database can be directly downloaded for the same host website.

Rotifers as Models in Toxicity Screening of Chemicals and Environmental Samples http://dx.doi.org/10.5772/61771 69

Other physical condition that might modify the sensitivity of a test organism is desiccation. Robles-Vargas and Snell [145] found a remarkable resistance of *Philodina sp*. emerged from desiccation to PCP, chlorpyrifos, and mercury in comparison to continuously hydrated rotifers

According to the information reviewed and in terms of acute toxicity, rotifers from the family Brachionidae tend to show a great sensitivity to different chemicals, including organic and inorganic compounds. Invertebrates from the family Euchlanidae and Lecanidae appear to be sensitive mainly to inorganic toxicants. Furthermore, rotifers belonging to the family Philodi‐

Every chemical product that can reach ecosystems and, as a consequence, the human being needs to be characterized to avoid noxious effects on all organisms exposed. At first instance, toxicological studies were mainly anthropocentrically focused, but that point of view would not have been enough to protect our own existence at all if affecting organisms in different ecosystems would have continued. It is known that rotifers play an important role as secondary producers and in some freshwater ecosystems represent the larger fraction of biomass of zooplankton [146]. Therefore, these organisms are nowadays considered as model organisms

Currently, one can find in the scientific literature reports of almost every class of chemical compound used or produced by man, investigating their toxicological properties. These papers are relatively abundant in several websites specialized in gathering such kind of information, which is available to everyone who has access to the published material. For scientist, collecting this material is a daily activity that guarantees their studies to follow the right direction by not working in exactly the same topic and manner, which would represent the duplicity of information but at higher cost because of multiple double efforts and resources investment. Nonetheless, it must be pointed out that differences in susceptibility to toxicants can be found among clones or strains, which is very interesting to assess by multiclonal analyses [22, 147]. However, when it is not possible to access multiple clones or strains, previous reports (published material) from databases become more relevant as a source of comparison to

The USEPA, in an attempt to ease the search for environmental toxicological data, has created the ECOTOX database, hosted in their website [150]. It provides a user-friendly interface and allows discrimination among taxonomic and chemical groups, aquatic or terrestrial environ‐ ment, tests results or conditions, and type (year) of publication. Although it does not give access to the cited references, it returns a list of publications to be consulted elsewhere. In the ECOTOX results table, the data shown are species name, exposure type, chemical, media type, end points, bioconcentration factor, effects, and statistics, among others (Figure 4). A complete list of all abbreviations used in the ECOTOX database can be directly downloaded for the same

when reproductive tests were implemented [145].

68 Invertebrates - Experimental Models in Toxicity Screening

nidae exhibit a notorious resistance to chemicals.

**4. Rotifer database for screening toxicants**

improve the comprehension of the observed results.

for toxicological screening.

host website.

**Figure 4.** Print screen of ECOTOX database results. Query terms were *Brachionus* and chromium, selected from the op‐ tions displayed in the database [150].

Nowadays, toxicity assays with rotifers include very diverse toxicants, and a relatively high number of test have been assessed the effect of inorganic (metals and metalloids) and very versatile organic compounds like pesticides, solvents, colorants, detergents, and emergent toxicants, in which health and care products and pharmaceuticals form part of it (Table 4). Some of these toxicants are effective at concentrations as low as some nanograms per liter, altering rotifers population dynamics by different mechanisms like endocrine disruption.

**Species Chemical compound or metal Species Chemical compound or metal**

2,5-Dichloro aniline [150] Bc, Bp, Ed, Lh, Ll,

Bc, Lh, Ll, Lq Acetone [94, [150]] Methanol [150] Bc Acetylsalicylic acid [152, 153] Methoprene [150] Av Acrylamide [154] As, Ba, Bc, Bp, Ed Methyl parathion [97, 150]

Ab, Bc, Li, Lq Aluminum [93, 150, 156] Naphthalenol [150] Bc Amitriptyline [152, 153, 157] Naproxen [150] Bc, Br Ammonia [150] n-Hexane [150] Bc, Pa Ammonium chloride [150] Bc, Pa Nickel [150]

Bc Aroclor 1260 [150] Nitric acid, Cadmium salt [150]

Bc, Pp, Pr, Arsenic [150, 158>] Nonyl phenol [150] Bk Atenolol [159] Ofloxacin [161, 165] Bc, Kq Atrazine [150] Orphenadrine [152, 153, 157] Bc Atropine [152, 153, 157] Oxytetracycline [150] Bc Barium nitrate [150] Bc, Bk Oxytetracycline [159, 165] Bc, Lh, Ll, Lq Benzene [94, 150] Bc Paracetamol [152, 153]

2- [2- [2-(Dodecyloxy)ethoxy]ethoxy]ethanol, Hydrogen sulfate, Sodium salt [150]

Acetaminophen [150]

70 Invertebrates - Experimental Models in Toxicity Screening

Acrylic acid [150]

Bc Amphetamine sulfate [153]

Bc Bezafibrate [160]

Ab, Ai, Bc, Bh, Bp, Br, Bu, Ed, Lh, Ll, Lq, Pa

Bc

2,4-Dichloro aniline [150] Bc Malathion [152] 2,4-Dichlorophenoxyacetic [150] Bc Malathion [153] 2,4-dinitrochlorobenzene [150] Bc, Li, Lq Manganese [150], [166], [169] 2,4-xylenol [150] Bc MARLON A390 [150]

3,4-dichloro aniline [150] Av Methacrylamide [171]

Bc

Bc

Bc

Bc

Cadmium [94, 143, 150, 160] Para-dichlorobenzene [150]

Aldrin [155] N,N,N-Trimethyl-1-octanamonium

Alkyl\* sodium benzene sulfonate [150] N,N-Dimethyl-1-dodecanamine

Alkyl\* trimethyl ammonium chloride [150] N,N-Dimethyl-1-octanamine [150]

Lq, Pa Mercury [91, 94, 97]

Bc Metallic silver [150]

Methacrylic acid [150]

Molinate [150]

chloride [150]

[150]

Nicotine [152, 153]

Para-chlorophenol [150]

Note: superscripts indicate reference number

Notes: Species are abbreviated with the first letters of their genus and species: Ad = *Adineta vaga*, Ab = *Asplanchna brightwellii*, Ag = *Asplanchna girodi*, Ai = *Asplanchna intermedia*, As = *Asplanchna sieboldi*, Ba = *Brachionus angularis*, Bb = *Brachionus bidentata*, Bc = *Brachionus calyciflorus*, Bh = *Brachionus havanaensis*, Bk = *Brachionus koreanus*, Bp = *Brachionus patulus*, Bp = *Brachionus plicatilis*, Bq = *Brachionus quadridentatus*, Br = *Brachionus Rubens*, Bu = *Brachionus urceolaris*, Eb = *Epiphanes brachionus*, Ed = *Euchlanis dilatata*, Ft = *Filinia terminalis*, Hi = *Hexarthra intermedia*, Hm = *Hexarthra mira*, Ka = *Keratella americana*, Kc = *Keratella cochlearis*, Kq = K*eratella quadrata*, Lh = *Lecane hamata*, Li = *Lecane inermis*, Ll = *Lecane luna*, Lq = *Lecane quadridentata*, Mq = *Monostyla quadridentata*, Pa = *Philodina acuticornis*, Pr = *Philodina roséola*, Pp = *Plationus patulus*, Pq = *Platyias quadricornis*, Tp = *Testudinella patina.*

**Table 4.** Relation of toxicants and rotifers species used in ecotoxicological studies

The available data about toxicity to rotifers show that organic compounds, as a complex group, is the most studied (32.4%) followed by the drugs group (24.58%), which could be due to the complexity of both groups, as they include a huge variety of toxicants. Inorganic compounds among other groups represent 17% out of the total number of tests included in this chapter, which can be split up into inorganic (nonmetallic compounds) and metals (Figure 5). The last one comprised those commonly named heavy metals [148]. Other groups might not be as numerous as the former ones, but this should not be misinterpreted as a lack of interest from the environmental toxicologists, but a challenge and a continuous research to find out how these chemicals are affecting the aquatic ecosystems and their inhabitants like rotifers species.

**Species Chemical compound or metal Species Chemical compound or metal**

Lh, Ll, Lq Ethyl acetate [94, 168] Theophylline [152, 153]

Ab Iron [156] Tributylstannane [150]

Bc, Lq Glyphosate [150] Bc Verapamil [152, 153]

Notes: Species are abbreviated with the first letters of their genus and species: Ad = *Adineta vaga*, Ab = *Asplanchna brightwellii*, Ag = *Asplanchna girodi*, Ai = *Asplanchna intermedia*, As = *Asplanchna sieboldi*, Ba = *Brachionus angularis*, Bb = *Brachionus bidentata*, Bc = *Brachionus calyciflorus*, Bh = *Brachionus havanaensis*, Bk = *Brachionus koreanus*, Bp = *Brachionus patulus*, Bp = *Brachionus plicatilis*, Bq = *Brachionus quadridentatus*, Br = *Brachionus Rubens*, Bu = *Brachionus urceolaris*, Eb = *Epiphanes brachionus*, Ed = *Euchlanis dilatata*, Ft = *Filinia terminalis*, Hi = *Hexarthra intermedia*, Hm = *Hexarthra mira*, Ka = *Keratella americana*, Kc = *Keratella cochlearis*, Kq = K*eratella quadrata*, Lh = *Lecane hamata*, Li = *Lecane inermis*, Ll = *Lecane luna*, Lq = *Lecane quadridentata*, Mq = *Monostyla quadridentata*, Pa = *Philodina acuticornis*, Pr = *Philodina roséola*, Pp = *Plationus*

Ab, Li, Lq Iron [96, 156, 166] Xylene [150]

Isoniazid [152, 153] Ab, Bc, Bh Li, Lq,

Flutamide [150] Tributyltin chloride [150] Furosemide [170] Bc, Kq. Pa Trichlorfon [150] Gemfibrozil [160] Bk Trimethoprime [159]

Hexachlorophene [150] Lh, Ll,Lq Vinyl acetate [94]

Ethyl alcohol [150] ThiobencarB[150] Ethyl methacrylate [150] Thioridazine [153] Ethylene glycol [150] Li Tin [166] Fenitrothion [150] Lh, Ll, Lq Titanium [94] Fenofibrate [160] Bc, Lh, Ll, Lq Toluene [94, 150]

Endosulfan [150] Sodium selenate [150] Endothall [150] Sodium tetradecyl sulfate [150] Erythromycin [165] Sulfamethoxazole [159, 161, 165] Estradiol [150] Testosterone [168] Ethinylestradiol [94, 168] Thallium(I) sulfate [150]

Bc

Bc

Bc

Sodium oxalate [150]

Tributyl phosphate [150]

Warfarin [152]

Pa Zinc [93, 143, 150, 156, 166]

Diphenylhydantoin [152]

72 Invertebrates - Experimental Models in Toxicity Screening

Bc

Bc

Bc

Bc

Bc, Pa Ferrous sulfate [150]

Hydroquinone [150]

Isopropyl alcohol [150]

*patulus*, Pq = *Platyias quadricornis*, Tp = *Testudinella patina.*

**Table 4.** Relation of toxicants and rotifers species used in ecotoxicological studies

Note: superscripts indicate reference number

**Figure 5.** Comparison of toxicity tests between chemical groups conducted with different rotifers species.

Several rotifer species have been used to evaluate the effect and toxic mechanisms of the chemical groups aforementioned, where *B. calyciflorus* is the most abundant in the literature, with about 63% out of the total publications (Figure 6). However, there are other species that present advantages over brachionids, such as the littoral genus *Lecane* that might be used to assess the effect of sediment-associated toxicants, or the predator genus *Asplanchna*, used to evaluate toxicants biomagnification among rotifers and other aquatic invertebrates [149]. Therefore, rotifers represent a group of organisms that have not been completely studied and require more attention from toxicologists.

Red bars correspond to left *y*-axis and blue bars to right *y*-axis.

**Figure 6.** Number of ecotoxicological studies through by different rotifer species.

#### **5. Rotifer tests already used worldwide and in certain regions**

The ability to produce cysts has allowed the development of toxicity kits, called Rotoxkits, employed for acute/chronic marine and freshwater toxicity testing [179]. Cyst production is an outstanding characteristic that has enabled the development of several toxicity protocols using rotifers that have been used worldwide [180]. Rotifers are not directly represented in the legislation of several countries (as the cladocerans *Daphnia magna* and *Ceriodaphnia dubia* do). However, toxicity tests with rotifers have been published by official societies [43, 84, 181] Perhaps the most notorious participation of rotifer toxicity tests is when EPA asked BP plc (the former name of the company was British petroleum) to use the acute toxicity test with the euryhaline rotifer *B. plicatilis* to assess the toxicity generated after the 2010´s Gulf of Mexico oil spill [35]. The marine water rotifer toxicity test TK22 was used to analyze thousands of sites in the Gulf of Mexico; toxicity was analyzed for both oil and the oil dispersant used [36].

#### **5.1. Europe**

The Convention for the Protection of the Marine Environment of the North-East Atlantic (the "OSPAR Convention") was established in 1992. The convention started working in 1998. It has been ratified by Belgium, Denmark, Finland, France, Germany, Iceland, Ireland, Luxembourg, Netherlands, Norway, Portugal, Sweden, Switzerland, and the United Kingdom and approved by the European Community and Spain. One of their goals was the development of directives for the analysis of toxicity both in marine and freshwater environments. In their 2007 publi‐ cation "Practical Guidance Document on Whole Effluent Assessment" (OSPAR, 2007), they applied the protocol of determination of the chronic toxicity to *B. calyciflorus* in 48 h [84] and validated its use.

In the Port of Aveiro, Portugal, standardized acute toxicity test using the marine rotifer, *B. plicatilis*, on sediment elutriates revealed higher toxicity levels in cases where the European Union Water Framework Directive suggested that sediment quality was acceptable [183]. Isidori et al. [184] employing *B. plicatilis* in 24-h toxicity tests found that all samples of municipal solid waste landfills in southern Italy expressed acute toxicity.