Litter preparation

The decomposed litter used in the rearing experiment was collected in August 2020 from a deciduous broadleaf forest located in the upper reaches of the Ota River in Minamisōma City, Fukushima Prefecture. The site is approximately 30 km away from Tokyo Electric Power Company’s Fukushima Daiichi Nuclear Power Plant. This site is the forest where we have been studying the transfer of ¹³⁷Cs from aquatic and terrestrial insects to fish that consume them [18]. The litter was collected from relatively flat areas alongside headwater stream, where broadleaf litter accumulated. Dry streams or slope area were excluded to minimize the effects of wash-off due to water flow. The decomposed litter was obtained from the partially decomposed organic layer (F-H layer, including fermentation and humus sublayers) after removing fresh litter from the surface. Sampling of approximately 1.5 kg of litter was performed at 10 different locations and the collected litter was mixed. The litter was brought to the laboratory, dried in an oven at 60 °C for 7 d, ground and homogenized using an impact mill (IFM-800DG; Iwatani Ltd., Tokyo, Japan), and sieved through a 1-mm mesh sieve. Because there is a restriction on the quantity of radiocesium material that can be brought into the laboratory according to the safety management regulations for sample handling personnel established by the institute, the litter was used after mixing with ¹³⁷Cs-free commercially available substrate, which was made from powdered decayed oak wood (spawning first; Fortech Ltd., Wakayama, Japan) at a ratio of 1:1.

Preparation of flower chafer

We collected flower chafer imagines from May to September 2020 using traps at the institution property in Miharu Town, Fukushima Prefecture (Fig 1A). Protaetia orientalis is a common and known pest species [19]. It is not listed as a nationally endangered species under the Law for the Conservation of Endangered Species of Wild Flora and Fauna of Japan (https://www.env.go.jp/en/nature/biodiv/law.html). Therefore, permissions are not required for its collection and experimentation in the study area. The collected imagines were placed in a cage with a commercial ¹³⁷Cs-free substrate for oviposition and their offspring were used for the breeding experiment. The larvae were reared for 3–5 months at 20 °C. Subsequently, the third and last instar larvae were used for the rearing experiment (Fig 1B).

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Fig 1. Rearing experiment.

Pictures of (A) Imago and (B) larva of flower chafer, and (C) the U-8 container used for the rearing experiment.

https://doi.org/10.1371/journal.pone.0310088.g001

Breeding experiment

The design of the breeding experiment is shown in Fig 2. Before starting the breeding experiment, 250 g of litter moistened with water was placed in a plastic container (Fig 1C, U8 container, d = 50 mm; h = 62 mm) and the ¹³⁷Cs activity concentration of the litter was measured. Twenty third-instar larvae were reared for 30 d, with one larva in each U8 container kept in a laboratory under dark conditions at 20 °C. After one month, the larvae were removed from the U8 containers and fixed in 70% ethanol. The fixed larvae were cleaned by removing body surface contaminants using a brush. Subsequently, their muscle, skin, and digestive tract were dissected using dissecting scissors and the dissected body parts were dried at 60 °C for approximately 2 d. We prepared samples of each individual larvae, obtaining a total of 20 samples of each body part. After the dry weights were measured using a balance (XSE205 DualRange; Mettler Toledo Ltd., Tokyo, Japan), samples were cut into small pieces using dissecting scissors and stored in flat-bottom test tubes (diameter = 15 mm and height = 57 mm) until subsequent germanium (Ge) γ-ray spectrometry analyses. The larval feces left in pellet form in the U8 container were separated from the remaining leaf litter using a sieve with a mesh size of 1 mm. The feces and remaining leaf litter were dried at 60 °C for approximately 2 d, the dry weights were measured and homogenized, and the samples were stored in U8 containers until Ge γ-ray spectrometry analyses. Some larvae were reared until they reached the imago stage in the U8 containers by using the same litter; however, many larvae died before reaching the imago stage, resulting in four samples for prepupae/pupae and three samples of imagines.

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Fig 2. Flow chart of the breeding experiment.

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Estimating the biological half-life of ¹³⁷Cs

Approximately 60 third-instar larvae were reared for one month using the same litter containing ¹³⁷Cs as used in the breeding experiment in three cages (33 × 23 × 19 cm, 20 larvae per cage). They were then moved to a commercial ¹³⁷Cs-free substrate in three cages (33 × 23 × 19 cm, 20 larvae per cage) to evaluate the elimination of ¹³⁷Cs from the larvae. Subsequently, the larvae of six individuals were preserved in 70% ethanol for 0, 1, 3, 6, 12, 24, 48, 96, and 192 h. The larvae were then dissected, dried, weighed, homogenized, and stored in flat-bottom test tubes until Ge γ-ray spectrometry analyses, following the same procedures as those used for the breeding experiments.

The biological half-lives of ¹³⁷Cs in flower chafer larvae were estimated for the digestive tract, skin, and muscle tissues separately because different ¹³⁷Cs elimination rates were assumed for each part [13, 20, 21]. ¹³⁷Cs elimination was described using the following single- and two-component exponential models: (1) where A₀ = initial ¹³⁷Cs activity concentration (Bq/g), A_t = ¹³⁷Cs activity concentration at time t (days from the initial date of the larvae moving to ¹³⁷Cs-free litter) (Bq/g), and λ = elimination rate constants (d^-1). (2) where A_1, A₂ = initial ¹³⁷Cs activity concentration of the first and second components, respectively (Bq/g), A_t = ¹³⁷Cs activity concentration at time t (Bq/g), and λ₁ and λ₂ = elimination rate constants (d^-1) for each component. A two-component model has been used to model cases wherein the contaminant initially declines rapidly, followed by a gradual rate of decline [13, 20–22]. The parameters for these equations were estimated by performing a non-linear least-squares method using the “nls” function of R ver. 4. 3. 3, a programming language for statistical computing supported by the R Core Team and the R Foundation for Statistical Computing [23]. The AIC was compared between the estimated single and two-component model and the two-component model was selected because it had a smaller AIC value in both cases: digestive tract: AIC_{single-component model} = 104.3 and AIC_{two-component model} = 59.4 and skin and muscle tissues: AIC_{single-component model} = 48.0 and AIC_{two-component model} = 43.4). Using the rapid and slow decreasing rate constants, λ₁ and λ₂, of the two-component model, the biological half-lives (T_b) were calculated.

Sequential extraction of ¹³⁷Cs

After larval consumption, the sequential extraction method was adopted for the litter and fecal pellets. Twelve replicates of litter were subsampled from the litter prepared for the rearing experiment and 12 replicates of fecal pellets were prepared from 12 of the 20 individuals used for the rearing experiments. The homogenized litter and fecal samples after Ge γ-ray spectrometry analyses were dried at 60 °C for approximately 1 d before subsample.

Sequential extraction has been applied to evaluate the chemical forms of ¹³⁷Cs in soil, sediments, and litter [11, 12, 17, 24]. We conducted a ¹³⁷Cs extraction experiment based on Takechi et al. (2020); however, we added a water-soluble extraction step as a first step. We sequentially extracted ¹³⁷Cs, which exists in a (1) water-soluble form, (2) ion-exchangeable form (¹³⁷Cs that can be exchanged by ammonium ions), (3) oxide form (¹³⁷Cs that is bound to iron and manganese oxides and extracted by reductive dissolution), (4) organic form (¹³⁷Cs that is bound to organic matter and extracted by oxidizing and decomposing organic matter), and (5) fixed form (¹³⁷Cs that is trapped in minerals and is scarcely dissolved). The litter and fecal pellet samples were used for the following extraction procedure after determining ¹³⁷Cs activity concentration by Ge γ-ray spectrometry analyses. The steps can be described in detail below:

1. Step (1): Water-soluble form. Three grams (dry weight) of each sample were placed in a centrifuge tube and 30 mL of pure water was added. Three fecal pellet samples weighed less than 3 g (2.87–2.96 g) owing to insufficient sample volume. In these cases, the solvent was added at a ratio of 1 (sample):10 (solvent). The tube was stirred (150 rpm) at room temperature (25 °C) for 16 h using shaking incubators (PRAS, 12-R-FF, Preci Co., Ltd.) and then centrifuged at 7,931 rpm for 30 min (7000, KUBOTA) to separate the supernatant and residue. The supernatant was spontaneously filtered through a filter paper (Whatman 42, Global Life Science Technologies Japan K.K., pore size 2.5 μm) and the extract acquired after filtering was transferred to a U8 container. The residue was rinsed with 30 mL pure water, centrifuged at 7,931 rpm for 30 min, and the resultant supernatant was discarded. In each subsequent extraction, the residue was rinsed with the same volume of pure water as the extraction solution added in each step.
2. Step (2): Ion-exchangeable form. The residue from the previous step was mixed with 30 mL of 1 mol/L ammonium nitrate solution (1 mol/L NH₄NO₃). The resultant suspension was stirred, centrifuged, and filtered, and the acquired extract was separated under the same conditions as described in Step (1).
3. Step (3): Oxide form. The residue of the previous step was mixed with 60 mL of 0.5 mol/L hydroxyl amine hydrochloride (HONH₂-HCl, containing 2.5% v/v 2.0 mol/L HNO₃). The suspension was stirred, centrifuged, and filtered, and the extract was separated under the same conditions as described in Step (1).
4. Step (4): Organic form. The residue of the previous step was mixed with 12 mL of hydrogen peroxide (H₂O₂ 36%), adjusted to pH 2.0 using 1% HNO_3, and left at room temperature (20°C) for approximately 1 h until the reaction settled. The container (with a loose lid) containing the sample was placed in a thermostatic bath (Thermominder, SX-10N, TAITEC) at 85 °C and heated for 1 h while shaking it occasionally. Thereafter, the liquid was heated without a lid until the volume was less than 4 mL. This process was repeated two times. After the sample had cooled to room temperature, 36 mL of 1.0 mol/L NH₄NO₃ adjusted to pH 2.0 using concentrated nitric acid was added. The suspension was stirred, centrifuged, and filtered, and the extract was separated under the same conditions as described in Step (1).
5. Step (5): Residuals. Finally, the residue was transferred to a U8 container and thoroughly dried at 105 °C, and the dry weights were measured. The extracts obtained from Steps (1)–(4) were transferred to a U8 container and weighed.

¹³⁷Cs measurements

The radioactivity of the flower chafer samples was measured using a high-purity germanium (HPGe) coaxial detector (Canberra GCW6023; Mirion Technologies, Canberra Ltd., Tokyo, Japan). The leaf litter and fecal samples from the breeding experiment and the extract and residue from the sequential extraction experiment were stored in U8 containers, and ¹³⁷Cs activity concentrations were measured using HPGe (Canberra GCW6023; Mirion Technologies, Canberra Ltd., Tokyo, Japan, and SEG-EMS GEM 30–70; SEIKO EG&G Ltd., Tokyo, Japan). Most samples were measured for <10% of counting error, except for small quantities of dissected flower chafer samples in flat-bottomed test tubes, which were measured for <15% of counting error. The minimum detectable amount of ¹³⁷Cs above the detection limit was about 0.07 Bq in the well-type HPGe coaxial detector. The standardized mixed sources (¹³⁴Cs and ¹³⁷Cs) for calibrating the detectors were MX035 (Japan Radioisotope Association, Tokyo, Japan) for the well-type HPGe coaxial detectors and MX033U8PP (Japan Radioisotope Association, Tokyo, Japan) for the U8 container. The standardized mixed sources were measured with varying filling quantities. Measurement efficiencies for ¹³⁴Cs (604.66 and 795.76 keV) and ¹³⁷Cs (661.66 keV) were obtained, and three efficiency calibration curves were created for different filling heights. The concentration of ¹³⁷Cs was then calculated using the comparative method based on these curves. Gamma Studio software (SEIKO EG&G, Tokyo, Japan) and Spectrum Explorer (Mirion Technologies, Canberra Ltd., Tokyo, Japan) were used to analyze the γ-ray spectra. The sample activities were corrected for radioactive decay until sample collection. ¹³⁷Cs activity concentrations were expressed as ¹³⁷Cs concentrations on a dry weight basis. S1 and S2 Tables show the ¹³⁷Cs activity concentrations (Bq/kg) and total ¹³⁷Cs (Bq) of the breeding experiment and sequential extraction experiment, respectively.

Statistical analysis and calculation of assimilation rate and concentration ratio

For the breeding experiment, the distribution of both ¹³⁷Cs concentrations and log-transformed ¹³⁷Cs concentrations were not normal by the Shapiro-Wilk normality test and not equivalent across samples by the Levene’s Test for Homogeneity of Variance. Therefore, multiple comparisons were made using pairwise Wilcoxon rank-sum tests for ¹³⁷Cs concentrations in litter before breeding, feces, dissected tissues of flower chafers (digestive tract, skin, and muscle), and whole-body larvae with Bonferroni adjustments for p values. The ¹³⁷Cs concentrations in whole-body larvae were calculated using the weight and concentration of each tissue in the dissection experiment assuming that the total ¹³⁷Cs in whole-body larvae is equivalent to the sum of ¹³⁷Cs in each body part. The absorption rate of ¹³⁷Cs in flower chafer larvae during the breeding experiment was calculated for each individual (Fig 2). The total ¹³⁷Cs in the litter consumed by the larvae was calculated by subtracting the total ¹³⁷Cs in the remaining litter after breeding from the total ¹³⁷Cs in the provided litter (A). The total ¹³⁷Cs assimilated in the larval tissue was calculated as the sum of ¹³⁷Cs in the larval skin and muscle (B). The absorption rate at which ¹³⁷Cs in the diet was transferred to flower chafer larval tissue at the end of the experiment was calculated as (B/A) × 100. ¹³⁷Cs concentrations in whole-body larvae, prepupae/pupae, and imagines were compared using the pairwise Wilcoxon rank-sum test. Additionally, the concentration ratio (CR) was calculated as follows: CR = ¹³⁷Cs concentration in flower chafer (Bq/g) / ¹³⁷Cs concentration in litter (Bq/g) for whole-body larvae, prepupae/pupae, and imagines.

Based on the results of the sequential extraction experiment, the bioavailable fraction of ¹³⁷Cs (percentage of the sum of water-soluble, exchangeable, oxide, and organic forms to the total ¹³⁷Cs contained in the sample) between the litter and feces excreted from larvae was compared using pairwise Wilcoxon rank-sum tests. The quantity of extracted ¹³⁷Cs from the sample per gram in water-soluble, exchangeable, oxide, and organic forms and the remaining ¹³⁷Cs were also compared between the litter and feces. Because the distributions of the log-transformed total quantities of the extract were not normal and did not have equivalence variance across samples, we applied pairwise Wilcoxon rank-sum tests. All statistical analyses and calculations were performed using R ver. 4.3.1. [23].

Uptake and absorption of ¹³⁷Cs into flower chafer larvae

The results of the dissection experiment showed that the ¹³⁷Cs concentration in the digestive tract of the larvae was significantly higher than that in the skin and muscle tissues (p < 0.001, Fig 3A). The calculated distribution of ¹³⁷Cs in larvae showed that most ¹³⁷Cs was localized in the digestive tract of the larvae (85.1 ± 9.7% of the total ¹³⁷Cs), whereas the muscle and skin included only 10.9 ± 6.9% and 3.9 ± 3.1% of the total ¹³⁷Cs, respectively (Fig 3B). This ¹³⁷Cs distribution was comparable to that of earthworms showing 95% of the total ¹³⁷Cs in the digestive tract [13]. Most ¹³⁷Cs concentrations in detritivorous aquatic insects also have been described by their digestive tract contents, the removal of which has been observed to significantly reduce ¹³⁷Cs concentrations [20, 25].

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Fig 3. Concentrations of ¹³⁷Cs concentrations in flower chafer larvae.

(A)¹³⁷Cs concentrations in flower chafer larvae after the breeding experiment. Litter (provided as diet), and feces, digestive tract, skin, muscle, and whole-body of larvae are shown (N = 20). Different letters beside each box indicate significant differences in the ¹³⁷Cs concentrations based on the Wilcoxon rank-sum tests. (B) Percentage of ¹³⁷Cs distributed in each organ of larvae for total ¹³⁷Cs in flower chafer larvae (N = 20). Each violin represents the distribution of an individual measure, and the dots represent the means.

https://doi.org/10.1371/journal.pone.0310088.g003

No statistically significant changes in ¹³⁷Cs concentrations were detected between the litter and feces (p = 0.25, Fig 3A), possibly because the ¹³⁷Cs decrease due to transfer from litter to larvae was marginal. Additionally, fecal samples may have been contaminated with the surrounding litter powders when the larvae were discharged, which is indicated by the ¹³⁷Cs concentrations in feces being significantly higher than those in the digestive tract contents obtained by dissection (p = 0.018).

The localization of ¹³⁷Cs in the digestive tract indicates that most of the ¹³⁷Cs ingested by flower chafer larvae may be excreted in feces, with little ¹³⁷Cs absorbed into the body tissues. Moreover, the ¹³⁷Cs absorption rate estimated in this study was extremely low. The total ¹³⁷Cs in the litter consumed during one month of breeding (A) was 132.0 ± 49.8 Bq and that transferred to the body tissues of muscle and skin (B) was 0.42 ± 0.1 Bq; thus, the absorption rate was 0.39 ± 0.2%. Few comparable studies have investigated ¹³⁷Cs absorption by soil macroinvertebrates [13]. However, ¹³⁷Cs absorption in fish has been estimated via feeding experiments, in which fish were fed with radiocesium-labeled diets; notably, ¹³⁷Cs absorption varied greatly depending on the food items. ¹³⁷Cs absorption in brown trout was 55% from Chironormidae sp. Larvae, 48% from freshwater amphipods, 76% from freshwater snails, 23% from Ephemeroptera sp. Larvae, 82% from zooplankton, and 66% from brown trout muscle [26]. Much lower ¹³⁷Cs absorption in bluegill was reported from detritus (3%) and sediment-fed Chironomus larvae (7%–16%), indicating that fish can absorb most of the ¹³⁷Cs associated with tissues and absorb marginal quantities of ¹³⁷Cs from detritus and clay in Chironomus larvae [27]. ¹³⁷Cs absorption in flower chafer larvae in this study was lower than that in fish, indicating significantly lower absorption rates for soil macroinvertebrates from detritus containing clay minerals. ¹³⁷Cs absorption can be affected by the physicochemical forms of ¹³⁷Cs in the litter and the quantity of clay minerals; however, future studies are needed to clarify this relationship.

By rearing flower chafers to the imago stage, ¹³⁷Cs concentrations were substantially reduced throughout the developmental stages (Table 1). Compared to the concentrations in whole-body larvae, ¹³⁷Cs concentrations were reduced by 50% in prepupae/pupae (p = 0.06, marginal due to small sample size) and 72% in imagines (p < 0.01). The larva discharges most of its digestive tract content by purging the gut before pupation. Subsequently, the digestive tract structure of scarab beetles completely changes during the pupal stage, and the cuticle of the digestive tract and the remaining contents are excreted during eclosion [28]. These metamorphosis processes may eliminate ¹³⁷Cs from the digestive tract. The ¹³⁷Cs concentrations in imagines were approximately equal to those assimilated in the skin and muscle tissues of the larvae, suggesting that the imago ¹³⁷Cs concentration was determined by the ¹³⁷Cs assimilated into the tissues during the larval stage (Table 1). The calculated whole body ¹³⁷Cs CRs for larvae, prepupa/pupa, and imago were 0.27 ± 0.11, 0.13 ± 0.03, and 0.06 ± 0.02, respectively (Table 1). These CR values were almost consistent with the CR value (0.13 ± 0.10) determined previously for forest detritivorous insects (all CRs were calculated for the imago stage) [15], thus, confirming that the transferred ¹³⁷Cs concentration evaluated in our breeding experiment was within the range observed in the field.

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Table 1. ¹³⁷Cs concentrations and concentration ratios for flower chafer larvae at each developmental stage.

Standard deviations are shown in parentheses.

https://doi.org/10.1371/journal.pone.0310088.t001

Estimation of the biological half-life of ¹³⁷Cs

In the elimination experiment, ¹³⁷Cs concentrations decreased at different rates in the digestive tract, muscles, and skin tissues (Fig 4; Table 2). ¹³⁷Cs in the digestive tract was rapidly eliminated during the first 2 d. This result was consistent with a previous finding that reported that the contents were completely excreted from the digestive tract within 47 h in third-instar scarab larvae (Trypoxylus dichotomus) [29]. After 2 d, when the digestive tract contents were assumed to have been replaced by clean litter, ¹³⁷Cs in the digestive tract remained and decayed slowly. This can be attributed to litter powder trapped by the walls of the digestive tract. Although ¹³⁷Cs in the digestive tract gradually decreased, it was expected to remain in the tract until it was completely excreted by pupation and eclosion, as has been shown in breeding experiments up to the imago stage. The estimated biological half-life of ¹³⁷Cs in the fast component by excretion of digestive tract contents (0.26–0.64 d; T_b1 for digestive tract in Table 2) was slightly slower than those estimated for earthworms (0.10–0.33 d; summarized in Table 1 in Tanaka et al. 2018) and Trichopteran larvae (0.22–0.37 d; Fujino et al. 2018).

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Fig 4. Elimination of ¹³⁷Cs from flower chafer larvae.

The lines represent the prediction of the fitted two-component model (Table 2).

https://doi.org/10.1371/journal.pone.0310088.g004

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Table 2. Estimated parameters for the two-component exponential model of ¹³⁷Cs concentrations for flower chafer larvae.

For biological half-life, T_b, 95% confidence intervals are shown in parentheses.

https://doi.org/10.1371/journal.pone.0310088.t002

Contrastingly, ¹³⁷Cs in skin and muscle tissues decreased rapidly during the first day and a relatively constant decrease of ¹³⁷Cs followed until Day 8. The two-component model indicated that different pools of ¹³⁷Cs were likely involved in ¹³⁷Cs elimination [21, 30]. The fast component may be small pools of ¹³⁷Cs that are easily discharged within 1 d and the slow component may be ¹³⁷Cs that is assimilated into tissues and gradually discharged. The estimated biological half-life of the slow component (4.0–17.6 d; T_b2 for muscle and skin in Table 2) was relatively faster than that estimated in earthworms (15–54 d; summarized in Table 1 in Tanaka et al. 2018), although the direct comparisons were difficult due to the short experimental period of this study.

¹³⁷Cs whole-body burdens have been used to estimate the half-life in earthworms [13] and invertebrates [20, 21, 31]. In these estimated two-component models, short-lived and long-lived components were interpreted to represent gut clearance and physiological clearance of assimilated ¹³⁷Cs, respectively. However, our dissection experiment revealed ¹³⁷Cs kinetics more comprehensively; particularly, digestive tract and physiological clearance were represented by a two-component model respectively consisting of multiple ¹³⁷Cs pools.

Bioavailable ¹³⁷Cs by sequential extractions

We calculated the recovery rate by dividing the sum of ¹³⁷Cs extracted in each fraction by the total ¹³⁷Cs present in the pre-extraction samples. This yielded a recovery rate of 94.7 ± 7.9%, confirming the validity of the sequential extraction analysis of this study. Fig 5 shows the results of the sequential extraction of litter and feces after conducting the breeding experiment. Because ¹³⁷Cs dissolution primarily occurs by ion exchange, reductive dissolution of Fe and Mn oxides, and microbial degradation of organic matter, these fractions (water-soluble, exchangeable, oxide, and organic forms) were considered as the bioavailable fraction [17].

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Fig 5. Proportion of different forms of ¹³⁷Cs in litter (L) and feces (F).

The proportions shown are the means of 12 samples for litter and feces.

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The previous study conducting the sequential extraction experiments of litter in Fukushima reported that relatively mobile ¹³⁷Cs remained in constant proportions from 2012 to 2019, which were assumed to be co-extracted with soluble fats, waxes, oils, holocellulose, and acid-soluble lignin [11]. These forms of ¹³⁷Cs may be extracted as bioavailable ¹³⁷Cs in this experiment, although direct comparison is not possible due to differences in extraction solvents. Litter digestion by flower chafer significantly reduced bioavailable fraction from 23.2 ± 2.3% in litter to 17.7 ± 2.5% in feces (p < 0.001). Additionally, the quantity of extracted ¹³⁷Cs was significantly lower in the feces than in the litter for all forms of bioavailable ¹³⁷Cs (Fig 6A); the reduction was 45%, 22%, 29%, and 24% for water-soluble, exchangeable, oxidized, and organic forms, respectively. This suggested that litter feeding by flower chafer larvae may absorb ¹³⁷Cs through the digestive process not only in an easy-elution form (water soluble and exchangeable) but also in oxide and organic forms.

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Fig 6. Extracted bioavailable fraction of ¹³⁷Cs.

Extracted bioavailable fraction of ¹³⁷Cs from (A) water-soluble (WS), exchangeable (EX), oxidized (OX), and OR (organic) forms from litter (L) and feces (F) (N = 12). (B) ¹³⁷Cs remaining in residuals. Each violin represents the distribution of an individual measure, and the dots show the means. The p-values show the result of the Wilcoxon rank-sum tests.

https://doi.org/10.1371/journal.pone.0310088.g006

Scarab larvae consume cellulose from various sources, including plant roots, soil organic matter, and decaying wood, and extract nutrients and energy from these sources [32]. The degradation mechanism of these components in the digestive tract has been well-studied, especially in the scarab larvae of Pachnoda ephippiata [32, 33]. Particularly, their midgut is highly alkaline (pH 12) and the most easily digestible proteins are mobilized and absorbed by endogenous proteinases. The digestive tract contents are then passed into a hindgut fermentation chamber containing diverse microorganisms, which degrade cellulose under anaerobic conditions. The digestive system of scarab larvae may promote ¹³⁷Cs elution and absorption in oxide and organic forms present in the litter. In contrast, in our study, digestion did not reduce the quantity of residual ¹³⁷Cs in the feces compared to that in the litter (Fig 6B). More than 70% of the ¹³⁷Cs remained as residuals, which may include ¹³⁷Cs fixed in clay minerals [6, 7, 34]. In addition, the distribution of radiocesium-bearing microparticles (CsMPs) has been reported for the Ota River watershed where the litter was collected; therefore, CsMPs can also contribute to ¹³⁷Cs in the residuals [18]. The ¹³⁷Cs in clay minerals and CsMPs is assumed to be unavailable for biological uptake during digestion. Further investigation for the quantity of clay minerals and CsMPs present in the litter is needed because it is likely to have a significant impact on both the results of the sequential extraction and the ¹³⁷Cs absorption rate in flower chafer larvae.

The digestion process by detritivores potentially influences the bioavailability of ¹³⁷Cs in the litter via decomposition [14, 35, 36]. Our results showed that digestion by larvae reduced the bioavailable fraction of ¹³⁷Cs in the litter. This was contrary to the findings of previous studies that have suggested that digestion of litter by scarab larvae (Trypoxylus dichotomus) significantly increases the exchangeable fraction of KCl in the litter [14]. This inconsistency could be attributed to differences in the litter provided. Ishii et al. (2018) fed larvae with intact litter and concluded that physical decomposition increased ¹³⁷Cs elution in feces. However, the larvae in our experiment were fed with finely crushed litter; thus, the effect of physical decomposition was marginal, but the chemical digestion process may have reduced the bioavailable ¹³⁷Cs in the feces. The effect of the ingestion of flower chafer larvae on the bioavailability of ¹³⁷Cs in intact litter under natural conditions remains unclear. Therefore, further studies are required to evaluate the impact of feeding by detritivores on the bioavailability of ¹³⁷Cs in forest floor litter.

The low bioavailability of ¹³⁷Cs in the litter observed in this study was consistent with earlier observations that reported low absorption ¹³⁷Cs in the detritus of earthworms [13]. However, detritus feeders may play an important role in ¹³⁷Cs input into forest ecosystems by extracting the bioavailable fraction of ¹³⁷Cs stock that is found in abandance on the forest floor. Furthermore,¹³⁷Cs in the skin and muscle tissues of detritivores is probably present in a highly bioavailable form that is susceptible to absorption by predators, such as fish. Therefore, it is likely that more bioavailable ¹³⁷Cs accumulates at higher trophic levels in the food web. In the aquatic food web, the sequential extraction results indicated that the ratio of easy-elution forms (exchangeable and carbonate forms) was higher in planktivorous fish (almost 100%) than in plankton (40%–80%) [37]. Thus, estimating the bioavailability of ¹³⁷Cs in detritivore and higher trophic organisms would be useful for understanding ¹³⁷Cs dynamics through the detrital food chain.

Furthermore, future work to estimate the quantity of ¹³⁷Cs uptake by detritivores against stock in the forest floor is needed to elucidate the contribution of detritivores to the contamination of organisms in the forest ecosystem. In addition, the bioavailability and absorption rates of ¹³⁷Cs in forest litter may be influenced by factors such as tree species and the quantity of fixed ¹³⁷Cs in clay minerals. Therefore, the ¹³⁷Cs availability and absorption rates estimated in this study may not fully represent the broad contaminated area across Fukushima Prefecture. Estimating these parameters among different regions with varying tree species and other detritivorous insect species, would contribute to a more comprehensive understanding of the role of detritivores in the dynamics of ¹³⁷Cs in forest ecosystems.