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Article

Intra-Body Variations of Stable Isotope Ratios (δ13C, δ15N) and Influence of Storage Methods in Aquatic and Post-Aquatic Stages of the Common Toad, Bufo bufo

A.N. Severtsov Institute of Ecology and Evolution, Russian Academy of Sciences, Leninskiy pr. 33, 119071 Moscow, Russia
*
Author to whom correspondence should be addressed.
Water 2021, 13(18), 2544; https://doi.org/10.3390/w13182544
Received: 12 July 2021 / Revised: 8 September 2021 / Accepted: 14 September 2021 / Published: 16 September 2021
(This article belongs to the Section Biodiversity and Functionality of Aquatic Ecosystems)

Abstract

:
Isotopic signatures of carbon and nitrogen are widely used for analysis of the structure of food webs in aquatic ecosystems. The study of animals raises a number of methodological questions, including choice of representative tissues and organs for sampling as well as storage of the studied organisms. Furthermore, the impacts of preservation methods can be tissue-specific, age-specific, and even taxon-specific; thus, studies of these impacts on particular taxa are necessary. We focused on the C and N isotope composition of the common toad (Bufo bufo), one of the most widespread European anuran amphibians. We hypothesized that its different tissues and organs may vary in isotopic composition, and ethanol and freezing may have different effects on isotopic values. Our results showed that both “tissue” and “storage method” factors significantly affected the δ13C values of tadpoles and postmetamorphic juveniles, whereas only the “tissue” factor had a significant effect on the δ15N values. The two stages, tadpoles and postmetamorphs, should be analyzed separately despite the brief postmetamorphic period of the juveniles. The skin, legs, muscles, and tail in tadpoles and legs, muscles and heart in juveniles can be used for δ13C and δ15N analysis regardless of the method of storage. The results will serve for the optimization of future study designs in isotopic ecology.

1. Introduction

Isotopic signatures of carbon and nitrogen are widely used for analysis of the structure of food webs in both aquatic and terrestrial ecosystems [1,2,3]. The carbon 13C/12C ratio (usually expressed as δ13C) allows tracing basal carbon sources [4,5], whereas 15N/14N ratio (δ15N) increases as nitrogen passes from one consumer to another, indicating the trophic level of an animal in the trophic chain [6]. These phenomena make stable isotope analysis (SIA) a suitable tool for solving a wide range of questions in trophic ecology [7,8]. The unique isotope composition (“isotopic signature”) of an animal reflects its diet over time with a certain (sometimes significant) delay. In this case, carbon and nitrogen of different tissues and functional systems of the body are updated at different rates [9,10,11].
In small-sized animals of terrestrial and aquatic taxa (e.g., collembolans, mites, microcrustaceans), the isotopic composition can be measured in the whole organism without selection of a representative tissue or organ [12,13,14]. The use of a whole organism of large-sized animals is not possible and raises a number of methodological questions. The isotopic composition of different organic compounds that make up the tissues of organisms is not the same. For example, lipids are depleted in 13C and 15N in comparison with proteins and sugars [15,16]. Different tissues and body parts of animals differ significantly in the mass content of proteins, lipids, and may contain bones (vertebrates) or chitin (invertebrates). These differences determine the variation in the isotopic composition of various tissues and organs [14,17]. Indeed, a significant variation in the isotopic composition of carbon (δ13C) and nitrogen (δ15N) within one organism was shown for a number of invertebrates and vertebrates, including diplopods [18], orthopterans [19], aphids [20], fishes [21], reptiles [22,23], birds [24] and mammals [25,26,27]. However, we know little about such differences between tissues in amphibian [28,29,30], which play an important role in trophic webs of both aquatic (in larval stage) and terrestrial (as adults) ecosystems.
Muscles are considered the most convenient tissue for stable isotope study of an animal because structural elements are assumed to have lower rates of isotopic incorporation than splanchnic tissues and organs [31]. Muscles have been often used in studies on vertebrates [21], including amphibians [29,32]. However, the extraction of muscle tissue from the legs of tadpoles is possible only in the final developmental stages. The extraction of muscles from the legs of juveniles can also be difficult due to the small sizes of the specimens. However, the use of a homogenized individual for isotopic analysis may result in an incorrect assessment due to the possible chemical differences between tissues and organs.
Another important question relates to the methods of material preservation prior to isotopic analysis. Freeze drying and subsequent dry storage is considered the most convenient conservation method [33], which does not affect the isotopic composition of the material [34,35,36]. However, the use of a freeze-dryer for the preservation of samples during field studies is very difficult, since this is a piece of stationary laboratory equipment that is difficult to transport to the field. Under these conditions, samples can be frozen in the freezer of a household or mobile refrigerator (usually at −18 °C) or fixed with preservative liquids, among which ethanol is most frequently used [37]. However, ethanol is known to alter stable isotope ratios in samples [27,36,38]. Furthermore, the impacts of storage methods are taxon-specific, age-specific (see e.g., [33,39,40]) and can even be tissue-specific. Thus, studies on the impacts of these storage methods on particular taxa are necessary.
The influence of preservation methodology on stable isotope ratios has also been investigated in various taxa, such as terrestrial invertebrates [33,41], birds [42] and mammals [27], as well as aquatic microorganisms [39,43], invertebrates [36,44], fishes [45], and mammals [46]. Nevertheless, the influence of the storage method on the ratio of stable isotopes of such an important taxon as amphibians is not clear. Amphibian larvae occupy basal trophic niches, can reach significant seasonal biomass in fresh water bodies, and, after metamorphosis, move a great volume of organic matter to land, linking aquatic and terrestrial ecosystems [47,48]. After metamorphosis, anuran amphibians change their feeding habits from omnivorous to predatory [48]. The same is true for toads (Bufonidae) which are periodically the object of ecological studies with stable isotope analysis [49,50]. Moreover, there is practically no data on the effect of the storage method on different tissues of toads, as fixation with ethanol can alter the isotopic values of one tissue, but not affect the other [27]. Thus, in addition to choosing a particular tissue for isotope analysis, it is necessary to compare the methods of its preservation.
In our study, we focused on isotope composition of the common toad (Bufo bufo), one of the most widespread European anuran species. We hypothesized that different tissues may differ in isotopic composition, and ethanol may affect the isotopic values of C and N, and formulated the following tasks: (i) to assess the differences in the C and N isotopic composition of tissues and organs within the same developmental stages for tadpoles and separately for juveniles; (ii) to test the impact of the most common preservation method (ethanol) on isotope composition of different tissues. Additionally, we compared isotopic values of the same tissues and organs of larval (“tadpoles”) and newly-metamorphosed juvenile (hereafter referred to as “juveniles”) toads.

2. Materials and Methods

2.1. Experimental Design and Sampling

We collected B. bufo samples at the Lake Glubokoe Hydrobiological Station (N 55°45′; E 36°30′) of the A.N. Severtsov Ecology and Evolution Institute, the Russian Academy of Sciences, located 70 km west from the center of Moscow [51]. Toads of two ages (tadpoles and juveniles; hereafter the factor is referred to as “stage”) were caught simultaneously on 19 June 2019. The Glubokoe lake has a 60 m wide nearshore shallow waters, overgrown by macrophytes with a domination of water horsetail Equisetum fluviatile L., 1753, common reed Phragmites australis (Cav.) Trin. ex Steud., 1871, and broadleaf cattail Typha latifolia L. (1753). The lake and surroundings belong to Glubokoe Lake Nature Reserve. There are no significant anthropogenic local pollutants, such as sewage, which can potentially affect isotopic composition of basal food of tadpoles in the studied lake. The region is characterized by warm summers and moderately cold winters, with the mean monthly temperatures ranging from −10 °C (January) to +17 °C (July) and an average annual precipitation of about 700 mm [52].
Tadpoles were collected close to the shoreline and newly-metamorphosed juveniles were collected on the bank near the shoreline. The collected tadpoles were at Gosner’s (1960) stage 37.4 ± 0.9 (mean ± standard error is presented here and below) with sizes: L = 33.6 ± 0.3 mm; m = 441.6 ± 10.1 mg; n = 10. All the collected juveniles (L = 12.3 ± 0.3 mm; m = 181.6 ± 9.0 mg; n = 10) completed metamorphosis and had resorbed their tail. After collection, tadpoles and juveniles were left starving in separate containers without food for two days to evacuate the digestive tract. The studied specimens were sacrificed by immersion in ethyl 3-aminobenzoate methanesulfonate (Sigma-Aldrich). This anesthetic does not have significant influence on isotope ratios of tissues [30]. Half of the caught specimens were placed in a freezer and stored at −18 °C, other half of the samples were fixed in 96% ethanol (hereafter the factor is referred to as “storage method”). We used the same quality and quantity of ethanol for all the samples. The samples were processed in February 2020, with a duration of storage of 8 months. Furthermore, the tissues and organs were extracted from each specimen (hereafter the factor is referred to as “tissue”). The replicate of each tissue or organ was five (from five different specimens). Thus, we used 10 larval individuals (5 for analysis of frozen and 5 for analysis of ethanol-stored tissues) as well as 10 juveniles (5 for analysis of frozen and 5 for ethanol-stored tissues). Additionally, 10 larval specimens were homogenized as a whole for analysis of freezing and ethanol storage effects (5 and 5, respectively) and 10 juvenile specimens were used as a whole to study freezing and ethanol storage effects (5 and 5, respectively). The following tissues and organs were analyzed in tadpoles: muscles from back, hind legs, skin, heart, tail, digestive tract including intestines, and larval stomach, the manicotto glandulare, (hereafter referred to as “muscles”, “legs”, “skin”, “heart”, “tail” and “guts”, respectively) and the whole homogenized specimens (hereafter referred to as “whole organism”). Unfortunately, we were not able to analyze heart samples from tadpoles after freezing because of destruction of this organ upon thawing. Thus, heart was excluded from some pairwise comparisons. The following tissues and organs were analyzed in juveniles after freezing and separated after ethanol fixation: muscles, legs, skin, heart, digestive tract (including guts and well-developed stomach) and whole homogenized specimens (hereafter referred to as “muscles”, “legs”, “skin”, “heart”, “guts” and “whole organism”, respectively). The collection and processing of the organisms are in line with all current regulations of the Russian Federation and approved by the Animal Care and Use Committee of the A.N. Severtsov Ecology and Evolution Institute, the Russian Academy of Sciences (permit # 12 from 14 May 2018).

2.2. Stable Isotope Analysis

All tissue samples were separately oven-dried at 50 °C for 3 days. To assess the mean isotopic values of whole specimen, we homogenized samples after drying using a ball mill (Retsch MM200, Retsch GmbH, Haan, Germany). Then, samples were weighed (approx. 300 µg) and wrapped in tin foil. Their isotopic compositions were determined using a Thermo-Finnigan Delta V Plus continuous-flow mass spectrometer (Thermo Electron GmbH, Bremen, Germany) coupled with an elemental analyzer (Thermo Flash 1112, Thermo Electron) at the Joint Usage Center “Instrumental Methods in Ecology” at the IEE RAS. The isotopic composition of N and C was expressed in the δ-notation relative to the international standards (atmospheric nitrogen and VPDB): δX(‰) = [(Rsample/Rstandard) − 1] × 1000, where R is the molar ratio of the heavier isotope to the lighter isotope. Samples were analyzed with reference gases calibrated against IAEA (Vienna, Austria), reference materials USGS 40 and USGS 41. The drift was corrected using an internal laboratory standard (casein). The standard deviation of δ15N and δ13C values in the reference material was <0.15‰ (n = 8). Along with stable isotope composition, the average mass values of nitrogen and carbon (%) were determined. A total of 125 samples and 20 internal laboratory standards (casein) were analyzed for δ13C, δ15N, C and N.

2.3. Data Analysis

Thus, the experiment had a factorial design with the three following treatments: the factors “stage”, “tissue”, and “storage method”, the influence of which was tested on carbon and nitrogen isotopic values, as well as on C/N ratios. To assess the effect of the factors “tissue”, “storage method”, and their interaction (shown as “Storage method × Tissue”) on the δ13C, δ15N and C/N values of tadpoles and juveniles separately (n = 60 with exception of 5 frozen heart samples and n = 60 samples, respectively), we applied general linear modelling (GLM) with the forward stepwise predictor selection method which permutes combinations of variables to find the best model built from variables with significant effects [53].
To estimate differences between δ13C and δ15N values of B. bufo tadpoles and juveniles, we applied GLM, forward stepwise predictor selection method, with factors “stage”, “tissue”, and their interaction (shown as “Stage × Tissue”). For this aim we used 100 samples in total, of which 50 were frozen and 50 were ethanol-fixed. The analyses were provided separately for frozen (n = 50) and ethanol-fixed (n = 50) samples. The comparison was made for samples of muscles, skin, guts, and legs taken from amphibians of both stages and whole specimens were used.
Before analyses, for assessing homogeneity and normality of the selected model, the data were tested using the Statistica residuals tool. All dependent variables were also checked for normal distribution with the Shapiro-Wilk test and showed to be normally distributed and homogenous and required no transformation, which allows us to apply GLM. After GLM, the significance of differences between means was assessed using Tukey’s HSD test. All statistical hypotheses were tested at the p < 0.05 significance level. Data processing was performed using Statistica 13.3 (TIBCO Software, Palo Alto, CA, USA).

3. Results

3.1. “Tissue” and “Storage Method” Factors in Tadpoles

Both “tissue” and “storage method” factors significantly affected the δ13C values of tadpoles (Table 1), but not the interaction of these factors. Fixation in ethanol increased δ13C values as compared to frozen tadpoles by an average of 0.4‰ (Figure 1A). However, pairwise comparison of different tissues and organs showed significant differences only for comparisons of the digestive tract. Fixation with ethanol of other tissues and organs did not affect δ13C values (Figure 1A). The range of δ13C mean values between different tadpole organs and tissues averaged about 1.5‰. The digestive tract was significantly depleted in 13C relative to the other organs (Figure 1A).
Only the “tissue” factor had a significant effect on the δ15N values of tadpoles (Table 1). The highest δ15N values were observed in tail, muscles, and the whole body (Figure 1B). The lowest δ15N values were in the digestive tract, which on average differed from the tail, muscles and the whole homogenized organism by 1.2‰, 0.7‰, and 0.7‰, respectively.
Storage of the sample in ethanol significantly reduced the C/N ratio in the tissues of the digestive tract, muscles, legs, tail, and the whole organism, but not in the skin (Figure 1C).

3.2. “Tissue” and “Storage Method” Factors in Juveniles

Both factors “tissue” and “storage method” significantly influenced the δ13C values in juveniles (Table 1), but not the interaction of the factors. Storage of samples in ethanol on average increased the δ13C values by 0.68‰. The highest difference between frozen and ethanol-fixed samples was in the digestive tract (Figure 1D). Mean δ13C values of different tissues varied in the range of 1‰, but only δ13C values of digestive tract (about −24.1‰) and legs (about −23.1‰) differed significantly (Figure 1D).
The δ15N values were influenced only by the “tissue” factor (Table 1). The mean δ15N values were significantly lower in the skin than in the digestive tract, legs, muscles, and the whole organism (Figure 1E), with the spread of mean values about 0.97‰.
The juvenile C/N ratio significantly depended on the interaction of factors “tissue” and “storage method” (Table 1). As in tadpole tissues fixed with ethanol, the C/N ratios of ethanol-fixing tissues of juveniles were significantly lower compared to frozen ones, including skin but not legs and whole homogenized organism (Figure 1F).

3.3. The Tissue Isotope Composition and Effect of Ethanol: Tadpoles vs. Juveniles

The δ13C values significantly differed (factor “stage”) in both frozen and ethanol-fixed tadpoles and juveniles (Table 2). In general, juveniles had higher δ13C values as compared to tadpoles (Figure S1A,C). The muscles, legs, and whole organisms showed lower δ13C values in frozen tadpoles compared to frozen juveniles (Figure S1A). The digestive tract and whole organism of individuals fixed with ethanol were also significantly different (Figure S1C). The skin did not differ significantly between stages for any of the fixation methods.
The δ15N values significantly differed (factor “stage”) in both frozen and ethanol-fixed tadpoles and juveniles (Table 2). The δ15N difference between the tissues of tadpoles and juveniles showed a similar trend in both frozen and ethanol-fixed animals (Figure S1B,D). Significant δ15N differences between frozen tadpoles and juveniles were found in the digestive tracts and legs (Figure S1B).

4. Discussion

4.1. Differences in Stable Isotope Signatures of Tissues/Organs and Influence of Storage Methods

4.1.1. Tissues/Organs

The results show differences in stable isotope signatures of some tissues/organs of the common toad at both age stages. The digestive tract of tadpoles was significantly depleted in 13C relative to other organs and depleted in 15N relative to the muscles and tail; the digestive tract of juveniles was significantly depleted in 13C relative to the legs and enriched in 15N relative to the skin (Figure 1A,B,D,E). These differences in isotope compositions can most likely be caused by the remains of food or presence of parasites and symbionts in the digestive tract (as special cleaning procedures were not performed for this isotope analysis). Indeed, we noted that the digestive tract of all tadpole and some juvenile individuals contained the remains of food despite the 2-day forced starving period. Tadpoles could repeatedly consume faeces due to caprophagy [48] from the bottom of containers and juveniles could most likely digest slowly or hunt small flying invertebrates. In amphibian studies, gut contents are often analyzed separately to assess the isotopic composition of their food [54]. The differences between consumer’s tissues and its food are common phenomenon conditioned by diet-tissue discrimination factors [55]. Moreover, there is no comprehensive information about the possible influence of symbiotic intestinal macro- and microorganisms and microbial fermentation processes in tadpole intestines [56,57,58,59] on isotope assimilation by an organism and on whole gut content chemical composition. Theoretically, parasites, symbionts and microbial intestinal components may affect the resulting isotope signature of gut content.
Earlier, Cloyed et al. [29] assessed tissue-specific trophic discrimination factors in the adult green frog Lithobates clamitans (Latreille, 1801) and found that bone collagen is the most affected tissue, compared to the skin and whole blood. However, we could not compare our results with those of Cloyed et al. [29] because taking of blood and bone collagen samples is rather difficult in small-sized amphibians.

4.1.2. Storage Method

We compared the two common storage methods that may be useful in absence of the freeze-drying device. As a whole, preservation of samples with ethanol can increase δ13C values but not δ15N values relative to freezing samples. Despite that freezing is the more preferable method for investigation of δ13C values [41], the choice of tissue preservation method is less important for analysis of δ15N signature in tadpoles and juveniles: both methods provide similar results and are well applicable. The greater impact of ethanol on the 13C value may be conditioned by presence, in this preservative, of carbon with its own specific isotopic signature, which can alter the isotope ratios of the analyzed tissues by dissolving and extracting lipids, which have lower 13C content relative to carbohydrates and proteins [33].
Importantly, we studied samples after a comparatively long storage period (8 months). Earlier, investigations of samples after a brief (5 days) storage did not reveal any impact of the ethanol method upon neither δ13C nor δ15N of tissues (e.g., [41]). Additionally, the effect of preservation methods on stable isotope composition may depend on storage duration [38,60].
In our study, storage of samples in ethanol significantly reduced the C/N ratio in the tissues of separate organs and the whole organism, but not in the skin of tadpoles and legs of juveniles (Figure 1C,D). Obviously, these organs contain a minimal amount of fat [61] resulting in a non-significant impact of ethanol. Similar to the above-discussed impact to δ13C value, this is apparently associated with the destruction of lipids and adsorption of C from fat-containing tissues [33].

4.2. Practical Recommendations Concerning Choice of Tissues/Organs and Preservation Methods (Freezing vs. Ethanol) for Stable Isotope Analysis

In tadpoles, any organ other than the digestive tract can be used for δ13C analysis, regardless of the method of storage, because the digestive tract of tadpoles and whole homogenized specimens have reduced δ13C relative to muscles, which are most commonly used in isotopic analysis. Excluding the digestive tract from the stable isotope analyses avoids the potential problems associated with cleaning the digestive tract from food residues, parasites, symbionts, and microbes. When analyzing tadpoles for δ15N values, we can disregard the method of fixation. However, deleting the digestive tract is necessary because it may significantly differ in 15N relative to other organs and, in particular, muscles. Additionally, the heart is not a convenient organ for analysis of samples after freezing because of post-freezing decomposition. The most popular tadpole tissues in stable isotope studies are muscles from the tail [62,63,64], but small-sized tadpoles are often used as a whole but without the digestive tract [62] or after a gut clearing period [65]. Our results showed that the above-listed approaches are appropriate for stable isotope analysis, however special attention should be given to gut clearing procedures because caprophagy is typical for many larval amphibians [48] and symbionts, parasites and microbes may have the potential to impact the resulting isotope signature.
In juveniles, any organs other than the digestive tract or whole organism may be used for δ13C analysis regardless of the method of fixation. Any organs of juveniles may be also used for δ15N analysis regardless of the method of storage, but it is better to avoid analysis of the skin, which is depleted in 15N relative to other organs. Restriction concerning usage of the digestive tract for isotope analysis in tadpoles is also important for juveniles. Our results suggest that the traditional usage of muscle tissue in stable isotope studies of amphibian juveniles (e.g., [37]) is quite a convenient method.
The results also showed that different developmental stages have to be analyzed separately as these stages differ significantly in both studied isotope values. Additionally, significant δ15N differences between frozen tadpoles and juveniles were found in the digestive tracts and legs. The guts of tadpoles and juveniles could contain remains of principally different food objects, as tadpoles are omnivorous, whereas juveniles are obligatory predators [48,66]. This also provides some restrictions for using the whole organism samples. Legs present as rudiments in larval amphibians but rapidly develop and grow during metamorphosis and in early juveniles [48]; so these organs undergo a faster incorporation of chemical elements and their isotopic composition does not reflect averaged food assimilation during entire larval period.
In general, we do not recommend the usage of the digestive tract in the analysis of both tadpoles and juveniles, even after a starvation period. The usage of whole homogenized organisms should be avoided, as the digestive tract or internal fat can significantly affect mean δ13C values. The use of muscle tissues may be most accurate for analysis in tadpoles and juveniles but some non-muscle organs are also well suited for analysis. Our experience suggests that the most convenient or easiest method would be using the muscle tissues from back part (closely to vertebral column) of tadpoles or juveniles because this body part is not transformed drastically during metamorphosis [61].
Importantly, non-lethal sampling of tissues for stable isotope analysis (small pieces of fins for larvae or fingers for adults) is possible and preferable for large-sized specimens of amphibians [30,54] because this group of vertebrates is recognized to be declining since the second half of 20th century [67]. However, when sampling small-sized larvae or juveniles it is difficult to adhere to this guidance. Therefore, we hope our findings will serve for optimization of future study designs in trophic ecology and hence will decrease the number of used amphibian individuals.

5. Conclusions

In tadpoles, skin, legs, muscle and tail can be used for joint 13C and 15N analysis regardless of the method of storage. Whole homogenized individuals may also somewhat differ in 13C and 15N from muscle tissues due to the large proportion of the digestive tract relative to the whole body. It is likely that, for small-sized larvae and newly-metamorphosed juveniles of anuran amphibians, one of the optimal approaches is to remove the digestive tract (or all viscera), homogenize the remaining organism and then use it to estimate δ13C and δ15N isotope values.
In the absence of a freeze-dryer, the most convenient method is freezing of amphibian samples for storage before stable isotope analysis. Ethanol is also an appropriate storage method for some of the above-listed tissues and organs, or a whole organism after deletion of the digestive tract, however, long-term storage in ethanol may slightly affect δ13C.

Supplementary Materials

The following are available online at https://www.mdpi.com/article/10.3390/w13182544/s1, Figure S1: Stable isotope composition (δ13C and δ15N values) of B. bufo tadpoles and juveniles after freezing (A and B) and ethanol-fixation (C and D). Asterisks indicate significant differences between tadpoles and juveniles according to the Tukey HSD test after GLM, p < 0.05.

Author Contributions

Conceptualization, A.N.R.; Methodology, Writing—Original Draft Preparation, Writing—Review and Editing, D.I.K., A.N.R.; Funding Acquisition, A.N.R., D.I.K. All authors have read and agreed to the published version of the manuscript.

Funding

The study was supported by the IEE RAS initiative for development of investigations at biological stations. Isotopic analyses were partly supported by the Russian Science Foundation (project 19-74-10104).

Institutional Review Board Statement

The study was conducted according to the guidelines of the Declaration of Helsinki, and approved by the Ethics Committee of the A.N. Severtsov Institute of Ecology and Evolution, the Russian Academy of Sciences (protocol # 12 from 14 May 2018).

Informed Consent Statement

Not applicable.

Data Availability Statement

The primary data are available upon request to the authors.

Acknowledgments

We thank M.G. Zibrova, E.A. Noskova and A.V. Popova for processing samples for isotopic analyses and J.A. Titova for linguistic corrections. The isotopic composition of C and N was processed at the Joint Usage Center «Instrumental Methods in Ecology» at the A.N. Severtsov Institute of Ecology and Evolution of the Russian Academy of Sciences, Moscow.

Conflicts of Interest

The authors have no conflicts of interest/competing interests.

References

  1. Peterson, B.J.; Fry, B. Stable isotopes in ecosystem studies. Annu. Rev. Ecol. Evol. Syst. 1987, 18, 293–320. [Google Scholar] [CrossRef]
  2. Layman, C.A.; Araujo, M.S.; Boucek, R.; Hammerschlag-Peyer, C.M.; Harrison, E.; Jud, Z.R.; Matich, P.; Rosenblatt, A.E.; Vaudo, J.J.; Yeager, L.A.; et al. Applying stable isotopes to examine food-web structure: An overview of analytical tools. Biol. Rev. 2011, 87, 545–562. [Google Scholar] [CrossRef]
  3. Boecklen, W.J.; Yarnes, C.T.; Cook, B.A.; James, A.C. On the use of stable isotopes in trophic ecology. Annu. Rev. Ecol. Evol. Syst. 2011, 42, 411–440. [Google Scholar] [CrossRef][Green Version]
  4. Araujo-Lima, C.; Forsberg, B.R.; Victoria, R.; Martinelli, L. Energy Sources for Detrivorous Fishes in the Amazon. Science 1986, 234, 1256–1258. [Google Scholar] [CrossRef] [PubMed]
  5. Anderson, W.B.; Polis, G.A. Marine subsidies of island communities in the Gulf of California: Evidence from stable carbon and nitrogen isotopes. Oikos 1998, 81, 75–80. [Google Scholar] [CrossRef]
  6. Post, D.M. Using stable isotopes to estimate trophic position: Models, methods, and assumptions. Ecology 2002, 83, 703–718. [Google Scholar] [CrossRef]
  7. Hobson, K.A. Tracing origins and migration of wildlife using stable isotopes: A review. Oecologia 1999, 120, 314–326. [Google Scholar] [CrossRef]
  8. Newsome, S.D.; Yeakel, J.D.; Wheatley, P.V.; Tinker, M.T. Tools for quantifying isotopic niche space and dietary variation at the individual and population level. J. Mammal. 2012, 93, 329–341. [Google Scholar] [CrossRef][Green Version]
  9. Schmidt, O.; Scrimgeour, C.M.; Curry, J.P. Carbon and nitrogen stable isotope ratios in body tissue and mucus of feeding and fasting earthworms (Lumbricus festivus). Oecologia 1999, 118, 9–15. [Google Scholar] [CrossRef] [PubMed]
  10. Phillips, D.L.; Eldridge, P.M. Estimating the timing of diet shifts using stable isotopes. Oecologia 2006, 147, 195–203. [Google Scholar] [CrossRef]
  11. Gratton, C.; Forbes, A. Changes in δ13C stable isotopes in multiple tissues of insect predators fed isotopically distinct prey. Oecologia 2006, 147, 615–624. [Google Scholar] [CrossRef]
  12. Klarner, B.; Maraun, M.; Scheu, S. Trophic diversity and niche partitioning in a species rich predator guild—Natural variations in stable isotope ratios (13C/12C, 15N/14N) of mesostigmatid mites (Acari, Mesostigmata) from Central European beech forests. Soil Biol. Biochem. 2013, 57, 327–333. [Google Scholar] [CrossRef]
  13. Schilder, J.; Tellenbach, C.; Möst, M.; Spaak, P.; van Hardenbroek, M.; Wooller, M.J.; Heiri, O. The stable isotopic composition of Daphnia ephippia reflects changes in δ13C and δ18O values of food and water. Biogeosciences 2015, 12, 3819–3830. [Google Scholar] [CrossRef][Green Version]
  14. Potapov, A.M.; Tiunov, A.V.; Scheu, S. Uncovering trophic positions and food resources of soil animals using bulk natural stable isotope composition. Biol. Rev. 2019, 94, 37–59. [Google Scholar] [CrossRef]
  15. Deniro, M.J.; Epstein, S. Influence of diet on distribution of carbon isotopes in animals. Geochim. Cosmochim. Acta 1978, 42, 495–506. [Google Scholar] [CrossRef]
  16. Webb, S.C.; Hedges, R.E.M.; Simpson, S.J. Diet quality influences the δ13C and δ15N of locusts and their biochemical components. J. Exp. Biol. 1998, 201, 2903–2911. [Google Scholar] [CrossRef]
  17. Shipley, O.N.; Matich, P. Studying animal niches using bulk stable isotope ratios: An updated synthesis. Oecologia 2020, 193, 27–51. [Google Scholar] [CrossRef] [PubMed]
  18. Semenyuk, I.I.; Tiunov, A.V. Intraspecific dispersion and age changes of isotopic composition (15N/14N, 13C/12C) of diplopod tissues (Myriapoda, Diplopoda). Izv. Penz. Gos. Pedagog. Univ. Im. V.G. Belinskogo 2011, 25, 428–436. (In Russian) [Google Scholar]
  19. Wehi, P.; Hicks, B. Isotopic fractionation in a large herbivorous insect, the Auckland tree weta. J. Insect Physiol. 2010, 56, 1877–1882. [Google Scholar] [CrossRef] [PubMed]
  20. Perkins, M.J.; McDonald, R.A.; van Veen, F.J.F.; Kelly, S.D.; Rees, G.; Bearhop, S. Important impacts of tissue selection and lipid extraction on ecological parameters derived from stable isotope ratios. Methods Ecol. Evol. 2013, 4, 944–953. [Google Scholar] [CrossRef]
  21. Pinnegar, J.K.; Polunin, N.V.C. Differential fractionation of δ13C and δ15N among fish tissues: Implications for the study of trophic interactions. Funct. Ecol. 1999, 13, 225–231. [Google Scholar] [CrossRef]
  22. Murray, I.W.; Wolf, B.O. Diet and growth influence carbon incorporation rates and discrimination factors (Δ13C) in desert box turtles, Terrapene ornata luteola. Herpetol. Conserv. Biol. 2013, 8, 149–162. [Google Scholar]
  23. Warne, R.W.; Wolf, B.O. Nitrogen stable isotope turnover and discrimination in lizards. Rapid Commun. Mass Spectrom. 2021, 35, e9030. [Google Scholar] [CrossRef]
  24. Hobson, K.A.; Clark, R.G. Assessing avian diets using stable isotopes 2. Factors influencing diet-tissue fractionation. Condor 1992, 94, 189–197. [Google Scholar] [CrossRef]
  25. Lesage, V.; Hammill, M.O.; Kovacs, K.M. Diet-tissue fractionation of stable carbon and nitrogen isotopes in phocid seals. Mar. Mam. Sci. 2002, 18, 182–193. [Google Scholar] [CrossRef]
  26. Ehrich, D.; Tarroux, A.; Stien, J.; Lecomte, N.; Killengreen, S.; Berteaux, D.; Yoccoz, N.G. Stable isotope analysis: Modelling lipid normalization for muscle and eggs from arctic mammals and birds. Methods Ecol. Evol. 2011, 2, 66–76. [Google Scholar] [CrossRef]
  27. Javornik, J.; Hopkins, J.B.; Zavadlav, S.; Levanič, T.; Lojen, S.; Polak, T.; Jerina, K. Effects of ethanol storage and lipids on stable isotope values in a large mammalian omnivore. J. Mammal. 2019, 100, 150–157. [Google Scholar] [CrossRef]
  28. Caut, S.; Angulo, E.; Courchamp, F. Variation in discrimination factors (δ15N and δ13C): The effect of diet isotopic values and applications for diet reconstruction. J. Appl. Ecol. 2009, 46, 443–453. [Google Scholar] [CrossRef]
  29. Cloyed, C.S.; Newsome, S.D.; Eason, P.K. Trophic discrimination factors and incorporation rates of carbon- and nitrogen-stable isotopes in adult green frogs, Lithobates clamitans. Physiol. Biochem. Zool. 2015, 88, 576–585. [Google Scholar] [CrossRef] [PubMed][Green Version]
  30. Bélouard, N.; Petit, E.J.; Huteau, D.; Oger, A.; Paillisson, J.-M. Fins are relevant non-lethal surrogates for muscle to measure stable isotopes in amphibians. Knowl. Manag. Aquat. Ecosyst. 2019, 420, 2–8. [Google Scholar] [CrossRef]
  31. Martinez del Rio, C.; Wolf, N.; Carleton, S.A.; Gannes, L.Z. Isotopic ecology ten years after a call for more laboratory experiments. Biol. Rev. 2009, 84, 91–111. [Google Scholar] [CrossRef]
  32. Smith, R.L.; Beard, K.H.; Shiels, A.B. Different prey resources suggest little competition between non-native frogs and insectivorous birds despite isotopic niche overlap. Biol. Invasions 2017, 19, 1001–1013. [Google Scholar] [CrossRef][Green Version]
  33. Krab, E.J.; Van Logtestijn, R.S.P.; Cornelissen, J.H.C.; Berg, M.P. Reservations about preservations: Storage methods affect delta C-13 signatures differently even in closely related soil fauna. Methods Ecol. Evol. 2012, 3, 138–144. [Google Scholar] [CrossRef]
  34. Barrow, L.M.; Bjorndal, K.A.; Reich, K.J. Effects of preservation method on stable carbon and nitrogen isotope values. Physiol. Biochem. Zool. 2008, 81, 688–693. [Google Scholar] [CrossRef][Green Version]
  35. Fleming, N.E.C.; Houghton, J.D.R.; Magill, C.L.; Harrod, C. Preservation methods alter stable isotope values in gelatinous zooplankton: Implications for interpreting trophic ecology. Mar. Biol. 2011, 158, 2141–2146. [Google Scholar] [CrossRef][Green Version]
  36. Le Bourg, B.; Lepoint, G.; Michel, L.N. Effects of preservation methodology on stable isotope compositions of sea stars. Rapid Commun. Mass Spectrom. 2020, 34, e8589. [Google Scholar] [CrossRef] [PubMed]
  37. Bissattini, A.M.; Buono, V.; Vignoli, L. Disentangling the trophic interactions between American bullfrogs and native anurans: Complications resulting from post metamorphic ontogenetic niche shifts. Aquat. Conserv. Mar. Freshw. Ecosyst. 2019, 29, 270–281. [Google Scholar] [CrossRef]
  38. Jesus, F.M.; Pereira, M.R.; Rosa, C.S.; Moreira, M.Z.; Sperber, C.F. Preservation methods alter carbon and nitrogen stable isotope values in crickets (Orthoptera: Grylloidea). PLoS ONE 2015, 10, e0137650. [Google Scholar] [CrossRef] [PubMed][Green Version]
  39. Carabel, S.; Verísimo, P.; Freire, J. Effects of preservatives on stable isotope analyses of four marine species. Estuar. Coast. Shelf. Sci. 2009, 82, 348–350. [Google Scholar] [CrossRef]
  40. Vanderklift, M.A.; Ponsard, S. Sources of variation in consumer-diet δ15N enrichment: A meta-analysis. Oecologia 2003, 136, 169–182. [Google Scholar] [CrossRef]
  41. Butler, O.M.; Rashti, M.R.; Chen, C. Influence of storage and drying methods on invertebrate elemental and isotopic measurements. Comm. Soil Sci. Plant Anal. 2018, 49, 1–7. [Google Scholar] [CrossRef]
  42. Bugoni, L.; Mcgill, R.A.R.; Furness, R.W. Effects of preservation methods on stable isotope signatures in bird tissues. Rapid Commun. Mass Spectrom. 2008, 22, 2457–2462. [Google Scholar] [CrossRef] [PubMed]
  43. Oczkowski, A.; Thornber, C.S.; Markham, E.E.; Rossi, R.; Ziegler, A.; Rinehart, S. Testing sample stability using four storage methods and the macroalgae Ulva and Gracilaria. Limnol. Oceanogr. Methods 2015, 13, 9–14. [Google Scholar] [CrossRef][Green Version]
  44. Fanelli, E.; Cartes, J.E.; Papiol, V.; Rumolo, P.; Sprovieri, M. Effects of preservation on the δ13C and δ15N values of deep sea macrofauna. J. Exp. Mar. Biol. Ecol. 2010, 395, 93–97. [Google Scholar] [CrossRef]
  45. Durante, L.M.; Sabadel, A.J.M.; Frew, R.D.; Ingram, T.; Wing, S.R. Effects of fixatives on stable isotopes of fish muscle tissue: Implications for trophic studies on preserved specimens. Ecol. Appl. 2020, 30, e02080. [Google Scholar] [CrossRef] [PubMed]
  46. Kiszka, J.; Lesage, V.; Ridoux, V. Effect of ethanol preservation on stable carbon and nitrogen isotope values in cetacean epidermis: Implication for using archived biopsy samples. Mar. Mamm. Sci. 2014, 30, 788–795. [Google Scholar] [CrossRef]
  47. Gibbons, J.W.; Winne, C.T.; Scott, D.E.; Willson, J.D.; Glaudas, X.; Andrews, K.M.; Todd, B.D.; Fedewa, L.A.; Wilkinson, L.; Tsaliagos, R.N.; et al. Remarkable amphibian biomass and abundance in an isolated wetland: Implications for wetland conservation. Conserv. Biol. 2006, 20, 1457. [Google Scholar] [CrossRef]
  48. Wells, K.D. The Ecology and Behavior of Amphibians; The University of Chicago Press: Chicago, IL, USA; London, UK, 2007. [Google Scholar]
  49. Glos, J.; Ruthsatz, K.; Schroder, D.; Riemann, J.C. Food source determines stable isotope discrimination factor dN and dC in tadpoles. Amphib. Reptil. 2020, 41, 501–507. [Google Scholar] [CrossRef]
  50. Reshetnikov, A.N.; Korobushkin, D.I.; Gongalskiy, K.B.; Korotkevich, A.Y.; Selskaya, A.N.; Kotov, A.A.; Tiunov, A.V. Trophic positions and niche segregation of two anuran species in the ecosystem of a forest lake. Hydrobiologia 2021. [Google Scholar] [CrossRef]
  51. Smirnov, N.N. Lake Glubokoe (Moscow region, Eastern Europe), general characteristics. Hydrobiologia 1986, 141, 1–6. [Google Scholar] [CrossRef]
  52. Annenskaya, G.N.; Zhichkova, V.K.; Kalinina, V.P.; Mamay, I.I.; Nizovtsev, V.A.; Khrustaleva, M.A.; Tseselchuk, Y.N. Landscapes of Moscow Region and Their Current Condition; SGU: Smolensk, Russia, 1997; 296p. (In Russian) [Google Scholar]
  53. Miller, J.; Haden, P. Statistical Analysis with the General Linear Model; Department of Psychology, University of Otago: Dunedin, New Zealand, 2006; Available online: https://www.otago.ac.nz/psychology/otago039309.pdf (accessed on 1 July 2021).
  54. Trakimas, G.; Jardine, T.D.; Barisevičiūtė, R.; Garbaras, A.; Skipitytė, R.; Remeikis, V. Ontogenetic dietary shifts in European common frog (Rana temporaria) revealed by stable isotopes. Hydrobiologia 2011, 675, 87–95. [Google Scholar] [CrossRef]
  55. Kadye, W.T.; Redelinghuys, S.; Parnell, P.S.; Booth, A.J. Exploring source differences on diet-tissue discrimination factors in the analysis of stable isotope mixing models. Sci. Rep. 2020, 10, 15816. [Google Scholar] [CrossRef] [PubMed]
  56. Beebee, T.J.C.; Wong, L.-C. Leucine uptake by enterobacterial and algal members of larval anuran gut flora. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 1992, 101B, 527–530. [Google Scholar] [CrossRef]
  57. Pryor, G.S.; Bjorndal, K.A. Effects of the nematode Gyrinicola batrachiensis on development, gut morphology, and fermentation in bullfrog tadpoles (Rana catesbeiana): A novel mutualism. J. Exp. Zool. 2005, 303A, 704–712. [Google Scholar] [CrossRef] [PubMed]
  58. Pryor, G.S.; Bjorndal, K.A. Symbiotic fermentation, digestal passage, and gastrointestinal morphology in bullfrog tadpoles (Rana catesbeiana). Physiol. Biochem. Zool. 2005, 78, 201–215. [Google Scholar] [CrossRef]
  59. Vences, M.; Lyra, M.L.; Kueneman, J.G.; Bletz, M.C.; Archer, H.M.; Canitz, J.; Handreck, S.; Randrianiaina, R.-D.; Struck, U.; Bhuju, S.; et al. Gut bacterial communities across tadpole ecomorphs in two diverse tropical anuran faunas. Sci. Nat. 2016, 103, 25. [Google Scholar] [CrossRef][Green Version]
  60. Korotkevich, A.Y.; Kuznetsova, N.A.; Tiunov, A.V. Changes in Carbon and Nitrogen Isotope Ratios (13C/12C and 15N/14N) in Springtails during Long-term Storage of Soil Samples. Russ. J. Ecol. 2016, 47, 572–574. [Google Scholar] [CrossRef]
  61. Terentiev, P.V. Frog; Sovetskaya Nauka: Moscow, Russia, 1950; 345p. [Google Scholar]
  62. Caut, S.; Angulo, E.; Díaz-Paniagua, C.; Gomez-Mestre, I. Plastic changes in tadpole trophic ecology revealed by stable isotope analysis. Oecologia 2013, 173, 95–105. [Google Scholar] [CrossRef][Green Version]
  63. Arribas, R.; Díaz-Paniagua, C.; Caut, S.; Gomez-Mestre, I. Stable isotopes reveal trophic partitioning and trophic plasticity of a larval amphibian guild. PLoS ONE 2015, 10, e0130897. [Google Scholar] [CrossRef][Green Version]
  64. Vassilieva, A.B.; Sinev, A.Y.; Tiunov, A.V. Trophic segregation of anuran larvae in two temporary tropical ponds in southern Vietnam. Herpetol. J. 2017, 27, 217–229. [Google Scholar]
  65. San Sebastián, O.; Navarro, J.; Llorente, G.A.; Richter-Boix, Á. Trophic strategies of a non-native and a native amphibian species in shared ponds. PLoS ONE 2015, 10, e0130549. [Google Scholar] [CrossRef] [PubMed][Green Version]
  66. Kuzmin, S.L. The Amphibians of the Former Soviet Union; Pensoft: Moscow, Russia; Sofia, Bulgaria, 2013; 383p. [Google Scholar]
  67. Houlahan, J.E.; Findlay, C.S.; Schmidt, B.R.; Meyer, A.H.; Sergius, L.; Kuzmin, S.L. Quantitative evidence for global amphibian population declines. Nature 2000, 404, 752–755. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Stable isotope composition δ13C, δ15N (‰ ± SE) and C/N of B. bufo tadpoles (AC) and juveniles (DF). Asterisks indicate significant differences between frozen and ethanol-fixed tissues according to Tukey HSD test after GLM, p < 0.05. Different letters indicate significant differences between mean values of certain type of tissues by Tukey HSD test. Heart of frozen tadpoles was not analyzed.
Figure 1. Stable isotope composition δ13C, δ15N (‰ ± SE) and C/N of B. bufo tadpoles (AC) and juveniles (DF). Asterisks indicate significant differences between frozen and ethanol-fixed tissues according to Tukey HSD test after GLM, p < 0.05. Different letters indicate significant differences between mean values of certain type of tissues by Tukey HSD test. Heart of frozen tadpoles was not analyzed.
Water 13 02544 g001
Table 1. Results of GLM (forward stepwise selection of factors) for mean δ13C, δ15N and C/N values of Bufo bufo tadpoles and juveniles. Heart values were removed from the “tadpole” dataset. “SS”—sums of the squares, “p”—probability levels, “Error” refers to the total unexplained variance remaining in the model. “n.s.” indicates lack of significant results (p > 0.05).
Table 1. Results of GLM (forward stepwise selection of factors) for mean δ13C, δ15N and C/N values of Bufo bufo tadpoles and juveniles. Heart values were removed from the “tadpole” dataset. “SS”—sums of the squares, “p”—probability levels, “Error” refers to the total unexplained variance remaining in the model. “n.s.” indicates lack of significant results (p > 0.05).
Factorsδ13Cδ15NC/N
SSpSSpSSp
Tadpoles
Storage method2.420.037 n.s.3.28<0.0001
Tissue10.30.004411.78<0.00019.68<0.0001
Storage method × Tissue n.s. n.s.0.800.0084
Error28.05 12.8 2.16
Model R20.31 0.479 0.864
Model p 0.0022 <0.0001 <0.0001
Juveniles
Storage method6.990.0004 n.s.3.2<0.0001
Tissue7.480.0166.30.00012.67<0.0001
Storage method × Tissue n.s. n.s.1.37<0.0001
Error22.5 1.01
Model R20.386 9.2 0.878
Model p 0.000570.4060.0001 <0.0001
Table 2. Results of δ13C and δ15N values comparison between the same tissues of B. bufo tadpoles and juveniles using GLM (forward stepwise selection of factors). “n.s.” indicates lack of significant results (p > 0.05).
Table 2. Results of δ13C and δ15N values comparison between the same tissues of B. bufo tadpoles and juveniles using GLM (forward stepwise selection of factors). “n.s.” indicates lack of significant results (p > 0.05).
FactorFreezingEthanol
δ13Cδ15Nδ13Cδ15N
SSpSSpSSpSSp
Stage9.260.0000152.1650.00029318.490.0000011.2920.044
Tissue8.960.0008074.3780.0000876.790.030980 n.s.
Stage × Tissue4.680.0258433.2580.000784 n.s. n.s.
Error13.84 5.011 25.44 14.54
Model R20.61 0.66 0.498 0.082
Model p 0.00002 0.000002 0.000008 0.044
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Reshetnikov, A.N.; Korobushkin, D.I. Intra-Body Variations of Stable Isotope Ratios (δ13C, δ15N) and Influence of Storage Methods in Aquatic and Post-Aquatic Stages of the Common Toad, Bufo bufo. Water 2021, 13, 2544. https://doi.org/10.3390/w13182544

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Reshetnikov AN, Korobushkin DI. Intra-Body Variations of Stable Isotope Ratios (δ13C, δ15N) and Influence of Storage Methods in Aquatic and Post-Aquatic Stages of the Common Toad, Bufo bufo. Water. 2021; 13(18):2544. https://doi.org/10.3390/w13182544

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Reshetnikov, Andrey N., and Daniil I. Korobushkin. 2021. "Intra-Body Variations of Stable Isotope Ratios (δ13C, δ15N) and Influence of Storage Methods in Aquatic and Post-Aquatic Stages of the Common Toad, Bufo bufo" Water 13, no. 18: 2544. https://doi.org/10.3390/w13182544

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