Skip to main content

Exposure to Batrachochytrium dendrobatidis affects chemical defences in two anuran amphibians, Rana dalmatina and Bufo bufo



Batrachochytrium dendrobatidis (Bd) is the causative agent of chytridiomycosis, one of the major causes of worldwide amphibian biodiversity loss. Many amphibians exhibit skin-based chemical defences, which may play an important role against invading pathogens, but whether the synthesis of these chemical compounds is enhanced or suppressed in the presence of pathogens is largely unknown. Here we investigated direct and indirect effects of larval exposure to the globally distributed and highly virulent Bd-GPL strain on skin secreted chemical defences and life history traits during early ontogeny of agile frogs (Rana dalmatina) and common toads (Bufo bufo).


Exposure to Bd during the larval stage did not result in enhanced synthesis of the antimicrobial peptide Brevinin-1 Da in R. dalmatina tadpoles or in increased production of bufadienolides in B. bufo tadpoles. However, exposure to Bd during the larval stage had a carry-over effect reaching beyond metamorphosis: both R. dalmatina and B. bufo froglets contained smaller quantities of defensive chemicals than their Bd-naïve conspecifics in the control treatment. Prevalence of Bd and infection intensities were very low in both larvae and metamorphs of R. dalmatina, while in B. bufo we observed high Bd prevalence and infection intensities, especially in metamorphs. At the same time, we did not find a significant effect of Bd-exposure on body mass or development rate in larvae or metamorphs in either species.


The lack of detrimental effect of Bd-exposure on life history traits, even parallel with high infection intensities in the case of B. bufo individuals, is surprising and suggests high tolerance of local populations of these two species against Bd. However, the lowered quantity of defensive chemicals may compromise antimicrobial and antipredatory defences of froglets, which may ultimately contribute to population declines also in the absence of conspicuous mass-mortality events.


Amphibians are among the most threatened vertebrate groups, with their populations declining worldwide [1,2,3]. Although there may be no single main cause of declines [3], diseases caused by infection with viral, bacterial or fungal agents are clearly among the most devastating factors [4,5,6]. Studies aiming at uncovering and better understanding the processes that take place during interactions between amphibian hosts and their pathogens are therefore in the focus of current conservation-oriented research [7,8,9].

Chytridiomycosis, a disease affecting amphibians, is caused by the chytrid fungi Batrachochytrium dendrobatidis (Bd) and Batrachochytrium salamandrivorans (Bsal) [10]. Because Bsal has only been discovered recently [11], we know comparatively little about it [12], so that here we concentrate on chytridiomycosis caused by Bd. Chytridiomycosis has already led to the decline or extinction of several hundred species [13] and continues to cause mass mortality events on five continents due to repeated introductions arising from human activities [14]. Bd infects keratinous epidermal layers of the skin [15] and impairs its osmoregulatory function. This effect can cause shifts in electrolyte balance leading to cardiac asystolic death in juveniles and adults [16]. Tadpoles exhibit keratinous elements only in their mouthparts, so that they are less susceptible to Bd infection than later life-stages [17, 18], and can act as reservoirs in natural habitats [19,20,21].

As a part of innate immune defences, amphibians de novo synthesise numerous chemical compounds in their skin, serving as the first line of defence against pathogens and parasites [22]. These compounds can be cytolytic peptides, steroids, alkaloids or biogenic amines [23,24,25,26,27]. The most widespread defences against pathogens are cytolytic antimicrobial peptides (AMPs) which have been reported for species across eleven anuran families [27]. These AMPs are active against viruses, bacteria and microscopic fungi, including Bd [27,28,29,30,31]. The susceptibility of amphibian species and populations to chytridiomycosis is related to differences in AMP profiles [32, 33]. Bufonid toads lack skin-secreted AMPs [22], but may instead produce bufadienolides from early larval development on [34,35,36]. These steroid compounds exhibit antimicrobial, antiprotozoal activity, and may protect toads also against Bd [37, 38].

In this study, our aim was to experimentally investigate whether exposure to Bd resulted in increased production of skin-borne chemical defences as expressions of phenotypic plasticity, or if it caused lowered synthesis of defensive chemicals due to costs of infection or because of immune suppression. Therefore we exposed agile frog (Rana dalmatina; Fitzinger 1838) and common toad (Bufo bufo; Linnaeus, 1758) larvae to a highly virulent Bd isolate and monitored consequences for chemical defences and life history traits in well-developed tadpoles. Potential effects of larval infection reaching beyond metamorphosis on the abovementioned parameters were also tested. Adult R. dalmatina individuals produce at least one skin secreted AMP, Brevinin-1 Da [39], and B. bufo secrete several bufadienolide compounds in the skin [35, 36, 40]. Therefore, these species are suitable for the investigation of interactions between Bd infection and chemical defences.


Rana dalmatina

Control individuals all remained uninfected. The low Bd dose treatment resulted in very low infection prevalence and intensity in both tadpoles and froglets, while the high Bd dose treatment resulted in higher prevalence and intensity values in tadpoles and especially so in froglets (Table 1; Fig. 1A).

Table 1 Prevalence of Bd infection in the studied species after experimental exposure to Bd. Ntot is the total number of Bd-exposed individuals surviving until sampling, Ninf is the number of infected individuals at sampling. Control individuals are not shown because all remained uninfected
Fig. 1

Bd Load as zoospore genomic equivalents in case of Rana dalmatina froglets (A) Bufo bufo tadpoles (B) and B. bufo toadlets (C). Tadpoles of R. dalmatina are not depicted because of extremely low Bd prevalence (one tadpole infected in the low Bd treatment and three tadpoles infected in the high Bd treatment). Note that scales are different. Horizontal lines represent medians, boxes represent interquartiles, bars represent ranges, asterisks indicate outliers (deviating from the boundary of the interquartile range (IQR) by more than 1.5 × IQR)

Treatment had no effect on the relative amount of Brevinin-1 Da in pooled samples of tadpoles (GLM: F2,15 = 2.31, P = 0.13; Fig. 2A). However, older tadpoles tended to exhibit larger amounts of the peptide than younger conspecifics (F1,16 = 4.51, P = 0.05). Infection intensity and body mass were not related to Brevinin-1 Da quantity (infection intensity: F1,16 = 1.14, P = 0.30; body mass: F1,16 = 0.67, P = 0.43). In case of froglets, however, exposure to Bd resulted in significantly reduced relative amounts of Brevinin-1 Da in both Bd treatments as compared to the controls (GLM: F2,13 = 30.60, P < 0.001; Fig. 2B). The other measured variables had no detectable effect on Brevinin-1 Da quantity (length of larval development: F1,12 = 0.37, P = 0.55; infection intensity: F1,12 = 0.35, P = 0.57; body mass: F1,12 = 3.31, P = 0.09).

Fig. 2

Quantity of the antimicrobial peptide Brevinin-1 Da in pooled samples of R. dalmatina tadpoles (A) and froglets (B). To obtain detectable quantities of AMP, we had to pool groups of three samples during sample preparation preceding chemical analysis. Letters in lower case indicate homogeneous subsets according to Tukey HSD post-hoc tests. Two overlapping data points are depicted next to each other in panel B in the control treatment. Note that in froglets the low Bd dose and the high Bd dose treatment contained only 5 replicates as opposed to 6 replicates in the other groups

Body mass of tadpoles was positively affected by development stage, but treatment had no significant effect on it, either alone, or in interaction with development stage. In case of froglets, treatment had a marginally non-significant effect on body mass, where individuals exposed to the high Bd dose treatment tended to be heavier than others. Length of larval development had no effect on body mass, either alone or in interaction with treatment (For details see Table 2). Treatment had no significant effect on development, either measured as tadpoles’ development stage at sampling or as the length of larval development in case of froglets (Table 2).

Table 2 The effect of treatments on development and body mass of individuals at two life stages in the two studied species. Results are based on General Linear Models. Significant differences are highlighted in bold

Bufo bufo

None of the control individuals were infected, but we obtained relatively high infection prevalence and intensities in toadlets exposed to the low Bd dose treatment and in both life stages exposed to the high Bd dose treatment (Table 1; Fig. 1B and C).

We detected 22 different bufadienolide compounds in B. bufo extracts, three of which we identified with the help of the standards as arenobufagin, telocinobufagin and bufotalin (Table 3). The presence of individual compounds showed varied age-dependent patterns: some bufadienolides were present at both life stages in all (e.g., compound 7) or nearly all individuals (e.g., compound 13), while others occurred in a high proportion of individuals only after metamorphosis (e.g., compounds 10–12; Table 3). After metamorphosis, toadlets produced more bufadienolide compounds and also experienced a two to three fold increase in TBQ compared to tadpoles (Fig. 3).

Table 3 Percentages of Bufo bufo individuals that contained various bufadienolide compounds in the two development stages. We could unambiguously identify three compounds based on the standards and detected another 19 compounds as unknown bufadienolides based on their characteristic UV spectra. Analytical properties of the detected compounds are provided. (N: sample size (number of individuals), m/z: mass/charge)
Fig. 3

Toxin content of B. bufo larvae and toadlets in the control and Bd-exposure treatments: Number of bufadienolide compounds (NBC; mean ± SE) in tadpoles (A) and toadlets (B), and total bufadienolide quantity (TBQ; mean ± SE) in tadpoles (C) and toadlets (D) after exposure to zero (control), low or high Bd zoospore concentrations. Note that the scale is not continuous in case of NBC. Letters in lower case indicate homogeneous subsets according to Tukey HSD post-hoc tests

The number of bufadienolide compounds (NBC) did not differ among treatments in either life stage (GLM; tadpoles: F2,49 = 0.49, P = 0.62; toadlets: N = 38, F2,35 = 1.6, P = 0.22; Fig. 3 A and B). NBC was positively related to dry mass in tadpoles (B = 0.16, SE = 0.08, F1,51 = 4.56, P = 0.038), but not in toadlets (F1,36 = 1.78, P = 0.19). Infection intensity and development had no significant effect on NBC either in tadpoles (infection intensity: F1,50 = 1.82, P = 0.18; development stage: F1,50 = 0.002, P = 0.97) or in toadlets (infection intensity: F1,36 = 2.31, P = 0.13; length of larval development: F1,36 = 0.02, P = 0.88).

Total bufadienolide quantity (TBQ) was not affected by treatment in B. bufo tadpoles (GLM; F2,50 = 0.31, P = 0.74; Fig. 3C), but differed significantly among treatments in case of toadlets (F2,34 = 4.08, P = 0.026; Fig. 3D). TBQ was not related to dry mass (F1,51 = 10.68, P = 0.20), but this relationship was positive in case of toadlets (B = 2.21, SE = 0.69, F1,34 = 10.39, P = 0.003). According to Tukey HSD post hoc tests on the residuals of the regression of TBQ on toadlet dry mass (R = 0.47, F1,37 = 10.06, P = 0.003), relative TBQ was lower by 23% in the high Bd dose treatment than in the low Bd dose treatment (Mean difference = − 33.31, SE = 11.93, P = 0.022), but these two treatment groups did not differ from the control (high Bd dose: Mean difference = − 23.99, SE = 11.49, P = 0.11; (low Bd dose: Mean difference = 9.32, SE = 11.49, P = 0.7; Fig. 3D). Infection intensity and development stage did not have an effect on NBC either in tadpoles (infection intensity: F1,51 = 0.69, P = 0.41; development stage: F1,51 = 0.04, P = 0.85) or in toadlets (infection intensity: F1,33 = 0.038, P = 0.85; length of larval development: F1,33 = 1.46, P = 0.24).

Body mass was affected by development stage in case of tadpoles: individuals that were less developed also had a lower body mass. Treatment and its interaction with development stage had no significant effect on body mass (Table 2). Within the high Bd dose treatment, where Bd prevalence was sufficiently high to be analyzed, tadpole body mass was again positively related to development stage (F1,15 = 29.6, P < 0.001), while infection intensity had no effect on mass (GLM: F1,14 = 0.003, P = 0.96). Fourteen days after completion of metamorphosis, body mass of toadlets was in a negative relationship with the length of larval development (Table 2). Treatment and its interaction with the length of larval development had no significant effect on toadlet body mass (Table 2). Within the low Bd dose treatment group, neither infection intensity, nor the length of larval development had an effect on body mass of toadlets (GLM: infection intensity: F1,10 = 2.01, P = 0.19; length of larval development: F1,10 = 0.38, P = 0.55). However, in the high Bd dose treatment, infection intensity had a significant negative effect (GLM: B = − 4.38, SE = 1.73, F1,10 = 6.39, P = 0.03), and the length of larval development a marginally significant positive effect (B = − 5.01, SE = 2.28, F1,9 = 5.01, P = 0.052) on toadlet body mass. Development was not affected by treatment at either life stage (Table 2), and it was not related to infection intensity in either one of the assessed treatment groups (Spearman correlation in case of tadpoles in the high Bd dose treatment: R = 0.025, N = 17, P = 0.93; Pearson correlations in case of toadlets: low Bd dose: R = − 0.301, N = 12, P = 0.34; high Bd dose: R = 0.157, N = 12, P = 0.63).


In the present study experimental exposure to Bd did not result in significantly increased production of the antimicrobial peptide Brevinin-1 Da in R. dalmatina tadpoles, nor did it influence bufadienolide toxin synthesis in B. bufo tadpoles. However, Bd-exposure during the larval stage negatively affected chemical defences in metamorphosed individuals of both species. These results suggest that neither larvae nor freshly metamorphosed individuals respond to Bd-exposure with enhanced synthesis of antimicrobial chemicals, and that infection during the larval stage may rather carries costs that manifest in decreased quantities of chemical defences in metamorphs. Our results further indicate that larvae of the agile frog (R. dalmatina) were resistant to infection with a highly virulent Bd isolate, as indicated by low prevalence and infection intensities. At the same time, tadpoles of the common toad (B. bufo) were not resistant, but tolerant, as suggested by high Bd prevalence and high infection intensities, but no malign effects on life history traits: exposure to Bd did not influence body mass or development rate in larvae or metamorphs in either species.

The knowledge available regarding the occurrence of AMP synthesis in larval anurans is limited and controversial. Skin-associated granular glands and their ducts are mostly immature before metamorphosis in most species [41, 42], suggesting no, or limited AMP production (but see [43]). However, at the same time, gland products may also be secreted by a merocrine process, where the secretum reaches the skin surface via exocytosis directly or through the epidermal interstitium [41, 44, 45]. Furthermore, the adaptive immune system is suppressed during metamorphosis to prevent immune responses against newly emerging tissue types [46], suggesting a greater reliance on innate immune defences against invading pathogens during this susceptible period. Schadich et al. [47] found no evidence of AMP synthesis in tadpoles of Litoria ewingii despite intense AMP production after metamorphosis, while Wabnitz et al. [48] demonstrated efficient AMP synthesis in the skin of Litoria splendida at both larval and adult life stages. While Woodhams et al. [49] did not find AMPs in an early larval stage (development stage 25 according to Gosner, [50]) in two closely related species, Rana arvalis and R. temporaria, we detected that R. dalmatina tadpoles produce de novo the same AMP as do adults [39] at least during late larval development (development stage 37 according to Gosner). Whether this discrepancy among studies indicates species-specific differences in AMP synthesis, or if AMP production starts in Ranids some time later during larval development remains to be determined.

Our expectation that AMP synthesis is boosted upon exposure to Bd in R. dalmatina was not met. Phenotypic plasticity in chemical defences is not well understood [51], but some studies suggest that environmental stressors such as competition [52] and pathogen presence [53, 54] can induce an increase in AMP synthesis in metamorphosed anuran amphibians. Furthermore, adult European water frogs (Pelophylax lessonae and P. esculentus) are capable of elevating AMP synthesis in response to Bd, if the skin microbiota is suppressed [55]. Although during the tadpole stage we also observed a slight increase in the relative amount of Brevinin-1 Da upon Bd exposure, this was not significant, and we detected a sharp reduction in AMP quantity in both Bd-exposed groups in froglets. Wild caught, infected adults of the frog Litoria serrata exhibited similarly reduced quantities of AMPs compared to uninfected conspecifics [56], but whether this was a cause or consequence of infection remained unknown. Our results suggest that reduced AMP expression can be a consequence of exposure to Bd. A reduced synthesis of AMPs may result from immunosuppression by Bd, as demonstrated in case of the adaptive immune system [57,58,59]. Lowered AMP production may also be a direct cost of infection, or may be indirectly caused by elevated corticosterone levels resulting from Bd-infection [60, 61] because elevated corticosterone levels can cause reduced AMP synthesis [62, 63]. Whether the reduced quantity of Brevinin-1 Da is still large enough to be effective against Bd in metamorphs, or other agents of the immune system can take over the role of Brevinin-1 Da in preventing severe infection requires further investigation.

Interestingly, AMP synthesis was reduced in metamorphs arising from the low Bd dose treatment without detectable amounts of Bd on all but one froglet. There are, however, precedents for significant effects of Bd exposure on various traits in the absence of confirmed infection. For example, Garner et al. [64] experienced significant mortality in B. bufo tadpoles and freshly metamorphosed individuals due to Bd exposure without detectable infection loads. Also, Bd exposure reduced growth in European treefrogs (Hyla arborea) without detectable levels of infection [65]. Individuals are probably able to prevent initial infections or naturally clear Bd infections acquired but may suffer the costs of the mounted immune response [66]. Alternatively, Bd presence in the surrounding aquatic environment may be sufficient to induce pathology or responses (e.g., via waterborne chemicals) even if infection does not occur, as shown in case of crayfishes (Procambarus spp. and Orconectes virilis) which are alternative hosts of Bd [67].

Exposure to Bd had no detectable effect either on the number of bufadienolide compounds or on total bufadienolide quantity in case of B. bufo tadpoles. Similarly, NBC of toadlets was also not affected by Bd presence. However, TBQ in the high Bd dose treatment was significantly lower compared to the low Bd dose treatment in toadlets. These results clearly indicate that Bd-prevalence and higher infection loads did not induce enhanced toxin synthesis. Besides their role in the chemical defence system, bufadienolides also contribute to the osmotic homeostasis of toads [68, 69]. The decreased TBQ in the high Bd dose may have resulted from a compensatory response to the altered electrolyte balance (reduced sodium and potassium concentrations) due to Bd infection [16], but this speculation needs experimental confirmation. Alternatively, Bd infection can lead to structural damage in the skin and its glands [10], which may have contributed to the lowered toxin production in the high Bd dose treatment. Whatever the cause is, heavily infected toadlets unable to produce the increase in toxin content after metamorphosis, may suffer from detrimental consequences because skin secreted bufadienolides can act as repellents against vertebrate predators [70, 71], they play a role in immune defence [37, 38, 72] and are important for osmotic homeostasis [68, 69]. Lowered TBQ in the high Bd dose treatment, thus, indicates a possible indirect negative effect of Bd infection, similarly to what we found in regard to the chemical defence of R. dalmatina.

Exposure to Bd did not affect the measured life history traits in either species at the tadpole stage. In larval anurans only the mouthparts are keratinized structures [17], thus, mortalities due to chytridiomycosis are rare and susceptibility varies among species [10, 18, 73, 74]. However, sublethal negative effects of Bd infection can also occur due to mouthpart damage [75, 76], lethargy and poor swimming performance, resulting in lowered body mass and growth [18, 74, 76, 77]. In the present study only a very few tadpoles of R. dalmatina became infected and in these individuals infection intensity was very low in both Bd treatments. In case of B. bufo tadpoles, prevalence of infection and infection intensities were high, especially in the high Bd dose treatment, so that the lack of fitness consequences was somewhat surprising. Two out of the three identified bufadienolide compounds (arenobufagin and telocinobufagin) were previously documented to moderately inhibit the growth of Bd [38]. These and some of the other unidentified bufadienolide compounds may have contributed to the high Bd-tolerance of B. bufo individuals, which lack AMPs. These results suggest that the studied populations exhibit low susceptibility to Bd infection during larval development: tadpoles of R. dalmatina appear to be highly resistant, while B. bufo larvae may be highly tolerant.

During metamorphic climax, Bd starts to colonise newly keratinized skin surfaces and spreads out on the entire animal [17, 78]. In the present study, both the prevalence of Bd (GPL, IA042) and infection intensities were high in case of B. bufo, which is consistent with previous studies that used the same isolate and resulted in significant negative effects on body mass and mortality of B. bufo originated from the United Kingdom [64, 79]. Mortalities due to chytridiomycosis in the closely related B. spinosus (formerly a subspecies of B. bufo) were also observed in Spain [64, 80]. Contrary to this, the length of larval development was not affected by Bd exposure in our experiment, and we did not observe significant negative effects on body mass of toadlets. Only in the high Bd dose treatment, infection intensity was negatively related to body mass: lighter individuals had higher infection intensities, than heavier ones. Whether this pattern was a cause or consequence of infection remains unclear. All in all, these results suggest that B. bufo individuals of the studied population in Central Europe may be more tolerant to Bd than those in Western European populations. Alternatively to this spatial hypothesis, the pattern may also arise due to temporal differences; toads may have adapted to the presence of Bd since the earlier studies [64, 79, 80]. Furthermore, populations may respond differently to Bd infection from year to year due to phenotypic plasticity or epigenetic changes. These processes could also have contributed to the observed differences between the present and former studies in toad susceptibility. Regarding R. dalmatina, we detected low prevalence of infection in the low Bd dose treatment and moderate prevalence with low infection intensities in the high Bd dose treatment, and no adverse effects of Bd-exposure on life history traits of froglets. These results are in line with those of previous field studies suggesting that R. dalmatina is resistant to chytridiomycosis [21, 81, 82]. The less keratinized skin as well as the presence of Brevinin-1 Da may make the skin of R. dalmatina less suitable for Bd growth, hence the lower probability and intensity of infection as compared to B. bufo, which speculations need further investigations in the future.


Our results provide evidence that exposure to Bd can have negative effects on two different types of chemical defences of phylogenetically distant species in a later life-stage, even in the absence of obvious immediate effects on life-history traits, or, indeed, actual Bd infection. Because the investigated chemical defences are widespread among amphibians, the results of this study are likely to be applicable to many species with similar chemical defences, and this hidden effect of Bd presence in aquatic habitats should be considered in the future. Whether weakened chemical defences lead to lowered fitness in affected individuals remains an open question that will need further investigation.


Experimental procedures

In March 2016, we collected 40 eggs from each of nine freshly laid egg clutches of R. dalmatina from a pond in the Pilis-Visegrádi-Hills, Hungary (47.767058 N, 18.981325 E). We transported them to the Experimental Station Júliannamajor of the Plant Protection Institute, Centre for Agricultural Research. The Közép-Duna-Völgyi KTVF issued the permission to conduct the study (PE/KTF:3596-6-8/2016) and the Ethical Commission of the ATK NÖVI approved the investigation in accordance with Good Scientific Practice guidelines and national legislation. We placed eggs from each clutch separately into plastic boxes (24 × 16 × 13 cm) holding 1 L of reconstituted soft water (RSW; [83]) at a constant temperature of 19 °C and a 12:12 h light:dark cycle. Eggs were disinfected by bathing them for 3 days in 10 mg/L chloramphenicol in order to prevent accidental Bd infection [84] because Bd is reportedly present in the study area [21], even if with low prevalence [82]. Although chloramphenicol is an antibiotic agent, it is also effective against Bd [84, 85] and can be safely used for the disinfection of amphibian eggs [86].

Five days after hatching, when larvae were at development stage 25 (Gosner), we started the experiment with 12 healthy-looking tadpoles from each family. We maintained tadpoles individually in plastic boxes (15 × 12 × 12 cm) filled with 1 L of RSW, and fed tadpoles with slightly boiled and smashed spinach complemented with Spirulina powder (1 m/m%) ad libitum. We changed water twice a week using different dip nets for each treatment to prevent contamination across treatments. Temperature was 19.4 ± 0.7 °C (mean ± SD) during the experiment. The light:dark cycle was adjusted weekly to outdoor conditions, starting with 12:12 h light:dark in late March which we gradually changed to 14:10 h by the end of April. We exposed tadpoles during the entire larval development to sterile culture broth (control), or to a low or high zoospore concentration in liquid Bd culture (low Bd dose and high Bd dose treatments hereafter; for details, see below). We assigned tadpoles to the treatments using stratified randomization, and arranged rearing boxes into randomized spatial blocks, each containing one replicate from each treatment. We exposed individuals of nine families to the three treatments in four replicates, which resulted in a total of 108 experimental units. Hatchlings that were not used in the experiment were released at the site of origin.

Thirty days after start of the experiment, we randomly selected half of the tadpoles, sampled their skin secretions (for details see below) then gently blotted them dry and weighed them to the nearest mg using an OHAUS-PA213 analytical balance. Thereafter, we euthanized and preserved tadpoles in 70% ethanol and stored samples at 4 °C until further analysis.

We monitored development of remaining tadpoles daily. When an individual reached development stage 42 (emergence of forelimbs; according to Gosner) we poured the water off, placed back the tadpole into the same box covered with a transparent and perforated lid, added 100 ml RSW and lifted one side of the container by ca. 2 cm to provide metamorphs with both a body of water and a dry surface. Once metamorphs reached stage 46 (complete tail resorption; according to Gosner) we placed individuals into new, covered boxes of the same size as before, equipped with wet paper towels and a piece of cardboard egg-holder as a shelter. We fed the froglets with small crickets (Acheta domestica, instar stage 1–2) ad libitum. Dates of metamorphosis and of completion of tail resorption were registered daily. Fourteen days after completion of tail resorption we weighed animals and humanely euthanized them using the “cooling then freezing” method [87], preserved individuals in 70% ethanol and kept samples at 4 °C until further processing.

We conducted the same experiment using individuals of B. bufo. However, we observed high mortality independently from treatments (15 out of 36 tadpoles died both in the control and in the low Bd-dose treatment, and 10 out of 36 tadpoles died in the high Bd-dose treatment), presumably due to a bacterial bloom caused by the simultaneous presence of Spirulina and culture broth (an effect we did not observe in case of R. dalmatina tadpoles in this experiment, or when tadpoles were fed solely with spinach in previous experiments). Consequently, 17 days after start of the first experiment, we re-started this part of the study and conducted the same experiment as described above with 108 randomly chosen B. bufo tadpoles from the same location with the following additional differences: (1) Eggs originating from 14 different clutches were kept in mixed groups in outdoor mesocosms containing 130 L of aged tap water, 40 g beech leaves and 0.5 L pond water with no prior chloramphenicol treatment. (2) Once brought into the laboratory after hatching, we fed tadpoles with smashed and slightly boiled spinach only (no Spirulina). (3) Light exposure adjustment was different because of the delayed start (4) Because of their smaller size, we fed toadlets with springtails (Folsomia sp.) after metamorphosis.

Maintenance of Bd culture and experimental exposure

We experimentally infected tadpoles with the global pandemic lineage (GPL) of Bd. This isolate originated from a dead Alytes obstetricans (IA042) collected in 2004 from a mass mortality event in Spanish Pyrenees. Cultures were maintained in mTGhL broth (8 g tryptone, 2 g gelatine-hydrolysate and 4 g lactose in 1000 ml distilled water) in 25 cm2 cell culture flasks at 4 °C and passed every three months into sterile mTGhL. One week before use, we inoculated 100 ml mTGhL broth with 1–2 ml of these cultures in 175 cm2 cell culture flasks and incubated them for seven days at 22 °C. We assessed the concentration of intact zoospores using a Bürker chamber at × 400 magnification. During inoculation of tadpoles’ rearing boxes, the mean initial concentrations were ~ 1.8 × 106 (used to infect R. dalmatina) and ~ 2 × 106 (used to infect B. bufo) zoospores (zsp)/ml in the flasks. These cultures were used for the high Bd dose treatment and we prepared a 100-fold dilution with sterile mTGhL broth for the low Bd dose treatment. After each water change, we inoculated 1 ml of these cultures into the tadpoles’ rearing boxes holding 1 L RSW, resulting in 18–20 zsp/ml in the low Bd dose treatment and 1800–2000 zsp/ml in the high Bd dose treatment. Similar zoospore concentrations have been used widely and successfully in studies involving experimental infection [18, 75, 88]. We inoculated controls with the same quantity of sterile mTGhL broth. Contaminated water and equipment were disinfected overnight with VirkonS before disposal [89].

Assessment of infection intensity using qPCR

We assessed infection intensity from dissected mouthparts in case of preserved tadpoles and from toe clips in case of metamorphs. We homogenized tissue samples, extracted DNA using PrepMan Ultra Sample Preparation Reagent (Thermo Fisher Scientific, Waltham, Massachusetts, USA) according to previous recommendations [90], and stored extracted DNA at − 20 °C until further analyses. We assessed infection intensity using real-time quantitative real-time polymerase chain reaction (qPCR) following a standard amplification methodology targeting the ITS-1/5.8S rDNA region [90] on a BioRad CFX96 Touch Real-Time PCR System. To avoid PCR inhibition by ingredients of PrepMan, samples were diluted ten-fold with double-distilled water. We ran samples in duplicate. In case the result was equivocal, we repeated reactions in duplicate. If it again returned an equivocal result, we considered that sample to be Bd positive [91]. Genomic equivalent (GE) values were estimated from standard curves based on four dilutions of a standard (100, 10, 1 and 0.1 zoospore genomic equivalents; provided by J. Bosch; Museo Nacional de Ciencias Naturales, Madrid, Spain).

Skin secretion sampling in R. dalmatina and analysis of Brevinin-1 Da

We analysed skin secretions in two ontogenetic stages: 30 days after start of the experiment, when larvae were in development stage 36.9 ± 1.4 (mean ± SD; according to Gosner) and 14 days after completion of metamorphosis. Following the procedure described in [92] we collected skin secretions non-invasively by bathing tadpoles or froglets individually in 5 ml polypropylene tubes containing 5 ml collection buffer with 0.1 mM norepinephrine (NE) bitartrate for 15 min. After removing animals, we acidified the NE solution by adding 50 µL of 99% trifluoracetic acid (TFA), to reach a final concentration of 1 V/V%. Samples were stored at − 20 °C until further analyses. As indicated by preliminary assessments of Brevinin-1 Da concentrations, we had to pool groups of three samples within treatments to obtain detectable quantities of the targeted AMP. As a pre-step of purification, we activated each reverse-phase Sep-Pak cartridges (200 mg, LiChrolut RP-18, Merck-Millipore) with 2 ml acetonitrile and subsequently rinsed with 2 ml solvent A (HPLC-grade water with 0.12 V/V% TFA; according to [93]). Next, we loaded the skin extracts onto the cartridges, saved the solution, rinsed cartridges with 2 ml solvent A, and loaded the saved solution onto the cartridges again. Finally, we rinsed cartridges with 4 ml solvent A. We eluted the purified skin secretion with 2 ml solvent B (70:30 acetonitrile:HPLC grade water, acidified with TFA to a final concentration of 0.1 V/V%) into 2 ml polypropylene tubes and dried samples using a vacuum centrifuge (Savant, Integrated Speed Vac System, ISS 100).

We accomplished peptide identification and quantification using nano-UHPLC-MS/MS liquid chromatography-mass spectrometry, with a Maxis II ETD QqTOF (Bruker Daltonics, Bremen, Germany) coupled to an Ultimate 3000 nanoRSLC system (Dionex, Sunnyvale, CA, USA) under the control of Hystar v.3.2 (Bruker Daltonics, Bremen, Germany). We dissolved samples in 2% acetonitrile and 0.1% formic acid in water, out of which 5 µl were injected onto an Acclaim PepMap100 C-18 trap column (100 µm × 20 mm, Thermo Scientific, Sunnyvale, CA, USA). We performed sample desalting and preconcentration with 0.1% TFA for 8 min with a flow rate of 5 µl/min. Peptides were separated on an ACQUITY UPLC M-Class Peptide BEH C18 column (130 Å, 1.7 µm, 75 µm × 250 mm, Waters, Milford, MA, USA) at 48 °C, using a flow rate of 300 nl/min. We used the following HPLC solvents: solvent A containing 0.1% formic acid in water and solvent B containing 0.1% formic acid in acetonitrile, with the gradient: 4% B from 0 to 11 min, followed by a 120 min gradient to 50% B, then elevated the concentration of solvent B to 90% in 1 min and kept it there for 10 min. After each sample a blank was run to avoid carry-over. Sample ionization was achieved in the positive electrospray ionization mode via a CaptiveSpray nanoBooster ion source with capillary voltage set to 1300 V, at 0.2 Bar nanoBooster pressure. The drying gas was heated to 150 °C with 3 L/min flow rate. For external mass calibration, we used the low concentration tuning mix from Agilent technologies via direct infusion. Internal mass calibration was performed via lock mass for each run using sodium formate with the following ion transfer parameters: prepulse storage 10 µs, collision transfer 10 µs, quadrupole ion energy 5 eV, Funnel 1 RF 400 Vpp, Multipole RF 400 Vpp. The collision RF was set to 1200 Vpp with 120 µs ion transfer time. We identified the chromatographic peaks as Brevinin-1 Da by comparing retention time and mass spectrum of the 453.35514+ fragment to a commercially purchased molecular standard (Biocenter Kft., Szeged, Hungary). We analysed chromatograms using the software Compass Data Analysis (version 4.0, Bruker daltonics Inc., Billerica, USA) to obtain chromatogram area values as quantity estimates for statistical analysis.

Skin toxin sampling in B. bufo and analysis of bufadienolides

We collected bufadienolide samples from tadpoles preserved at development stage 36.3 ± 0.9 (mean ± SD; according to Gosner) and from 14 day old toadlets (that finished metamorphosis 14 days earlier). We homogenized whole bodies with a homogenizer (VWR VDI 12) equipped with a dispersing tool (IKA S12N-7S), dried samples under vacuum at 45 °C using a rotary evaporator (Büchi Rotavapor R-134, Flawil, Switzerland), weighed dry mass (dry body mass henceforth) and re-dissolved samples in 1 ml absolute HPLC-grade methanol, aided by brief exposure to ultrasound in a bath sonicator (Tesla UC005AJ1). As the last step of sample preparation, we filtered samples through FilterBio nylon syringe filters (pore size = 0.22 μm) and stored them at − 20 °C until further analyses.

We analysed bufadienolide compounds by means of high-performance liquid chromatography coupled with diode-array detector and electrospray ionization mass spectrometry (HPLC–DAD-ESI–MS). We identified the chromatographic peaks as bufadienolides based on the UV spectrum [34] and by comparing their retention time and mass spectrum to those of the following commercially available standards: bufalin, bufotalin, resibufogenin, gamabufotalin, areno- and telocinobufagin (Biopurify Phytochemicals, Chengdu, China), cinobufagin (Chembest, Shanghai, China), cinobufotalin (Quality Phytochemicals, New Jersey, USA) and digitoxigenin (Santa Cruz Biotechnology, Dallas, TX, USA) or to compounds established by analysing a large sample obtained from an adult male common toad by gently massaging the parotoid glands.

We performed HPLC–MS measurements on a Shimadzu LC–MS 2020 instrument (Shimadzu, Kyoto, Japan) that consisted of a binary gradient solvent pump, a vacuum degasser, a thermostated autosampler, a column oven, a diode array detector and a single-quadrupole mass analyser with electrospray ionization (ESI–MS). Chromatographic separations were carried out at 35 °C on a Kinetex C18 2.6 µm column (100 mm × 3 mm i.d., Phenomenex) in series with a C18 guard column (4 mm × 3 mm i.d.) using 10 µL injections. Eluent A was 5% aqueous acetonitrile with 0.05% formic acid and eluent B was acetonitrile with 0.05% formic acid. The flow rate was 0.8 mL/min and the gradient was as follows: 0–2 min: 10.5–21.1% B; 2–15 min: 21.1–26.3% B; 15–24 min: 26.3–47.4% B; 24–25 min: 47.4–100% B; 25–30 min: 100% B; 30–31 min: 100–10.5% B; 31–35 min: 10.5% B. ESI conditions were as follows: desolvation line (DL) temperature: 250 °C; heat block temperature: 400 °C; drying N2 gas flow: 15 L/min; nebulizer N2 gas flow: 1.5 L/min; positive ionization mode. Data were acquired and processed using the LabSolutions 5.42v software (Shimadzu).

Statistical analyses

We analysed the data on the two species and life stages separately. We excluded two individuals due to extremely slow development (development stage was lower than the mean by more than 3 SD), and accidentally lost three individuals. This resulted in the following sample sizes used in the analyses: R. dalmatina; control: 35, low Bd dose: 36, high Bd dose: 35; B. bufo: control: 34, low Bd dose: 36, high Bd dose: 35.

We averaged GE values obtained from qPCR runs for each sample and subsequently rank-transformed means, because GE values that fall outside the standard interpolation curve are not estimated reliably.

In R. dalmatina we compared Brevinin-1 Da quantity among treatments using general linear models (GLM) entering log-transformed values of the chromatogram areas as the dependent variable. We used log-transformation to enhance normality of model residuals and homogeneity of variances. In case of 14 days old froglets, we only had 5–5 replicates in Bd-exposed treatments because of mortality. For each triplet, we calculated mean values of body mass, development stage and infection intensity. The initial model included treatment as a fixed factor, and body mass, development stage and infection intensity as covariates.

To calculate the number of bufadienolide compounds (NBC) present in each B. bufo individual, we assumed a compound to be present if the signal-to-noise ratio (S/N) of its peak was at least three. We estimated the quantity of each compound from the area values of chromatogram peaks based on the calibration curve of the bufotalin standard, and summed up these values to obtain an estimate of total bufadienolide quantity (TBQ) for each individual (for a similar approach see [34, 94]). We analysed effects of treatment on both toxin variables separated by life stages using GLMs. In case of TBQ, we entered as the dependent variable log-transformed values in case of tadpoles and square-root transformed values in case of metamorphs to enhance normality of model residuals and homogeneity of variances. Initial models included treatment as a fixed factor and dry mass, development stage and infection intensity as covariates. In case of a significant treatment effect, we used Tukey HSD post-hoc tests to reveal significant differences between treatment groups. When a covariate also had a significant effect, we ran post-hoc tests on residuals extracted from the regression of the dependent variable on the covariate.

We analysed body mass data using GLMs. Initial models included treatment as a fixed factor and an estimate of the speed of development (Gosner stage in case of tadpoles and length of larval development in case of metamorphs) as a covariate, as well as their interaction. To enhance normality of model residuals and homogeneity of variances, we entered log-transformed values of body mass in case of R. dalmatina tadpoles and B. bufo metamorphs. In case of the high Bd dose treatment in B. bufo tadpoles, and both Bd treatments in metamorphs, where Bd prevalence was high, we also investigated the effect of infection intensity on body mass in the Bd-exposed treatment groups with GLMs, including infection intensity and development stage as covariates. We analysed variation in development stage using GLMs with treatment as a fixed factor, except for R. dalmatina metamorphs where we used Kruskal–Wallis tests due to the non-normal distribution of model residuals and inhomogeneity of variances. In case of B. bufo tadpoles in the high Bd dose treatment we analysed the relationship between infection intensity and development stage using Spearman rank correlation and Pearson correlations in both Bd treatments of metamorphs.

We verified normal distribution of model residuals using Shapiro–Wilk tests and by inspecting diagnostic plots, and homogeneity of variances using Levene’s tests. We applied a backward stepwise model simplification procedure [95] to avoid potential problems due to the inclusion of non-significant terms [96]. We obtained statistics for removed variables by re-entering them one by one to the final model. All tests were two-tailed. Statistics were calculated using SPSS Statistics 20.0 for Windows.

Data availability

All data used in the analyses will be available from Figshare Repository. (Ujszegi et al., 2021).



Antimicrobial peptide


Batrachochytrium dendrobatidis


Batrachochytrium salamandrivorans


Genomic equivalent


General linear model


Global pandemic lineage


High performance liquid chromatography


Mass spectrometry


Number of bufadienolide compounds




Quantitative polymerase chain reaction


Reconstituted soft water


Standard deviation


Total bufadienolide quantity


Trifluoracetic acid


  1. 1.

    Collins JP, Storfer A. Global amphibian declines: sorting the hypotheses. Divers Distrib. 2003;9(2):89–98.

    Article  Google Scholar 

  2. 2.

    Stuart SN, Chanson JS, Cox NA, Young BE, Rodrigues ASL, Fischman DL, et al. Status and trends of amphibian declines and extinctions worldwide. Science. 2004;306(October):1783–6.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  3. 3.

    Campbell Grant EH, Miller DAW, Schmidt BR, Adams MJ, Amburgey SM, Chambert T, et al. Quantitative evidence for the effects of multiple drivers on continental-scale amphibian declines. Sci Rep. 2016;6:25625.

    Article  CAS  Google Scholar 

  4. 4.

    Blumer C, Zimmermann DR, Weilenmann R, Vaughan L, Pospischil A. Chlamydiae in free-ranging and captive frogs in Switzerland. Vet Pathol. 2007;44(2):144–50.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  5. 5.

    Skerratt LF, Berger L, Speare R, Cashins S, McDonald KR, Phillott AD, et al. Spread of chytridiomycosis has caused the rapid global decline and extinction of frogs. EcoHealth. 2007;4(2):125–34.

    Article  Google Scholar 

  6. 6.

    Price SJ, Garner TWJ, Nichols RA, Balloux F, Ayres C, Mora-Cabello De Alba A, et al. Collapse of amphibian communities due to an introduced Ranavirus. Curr Biol. 2014;24(21):2586–91.

  7. 7.

    Daszak P, Berger L, Cunningham AA, Hyatt AD, Green DE, Speare R. Emerging infectious diseases and amphibian population declines. Emerg Infect Dis. 1999;5(6):735–48.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  8. 8.

    Brunner JL, Storfer A, Gray MJ, Hoverman JT. Ranavirus ecology and evolution: from epidemiology to extinction. In: Gray MJ, Chinchar G V, editors. Ranaviruses: Lethal pathogens of ectothermic vertebrates. New York: Springer; 2015.

  9. 9.

    Garner TWJ, Schmidt BR, Martel A, Pasmans F, Muths E, Cunningham AA, et al. Mitigating amphibian chytridiomycoses in nature. Philos Trans R Soc B. 2016;371(1709):20160207.

    Article  Google Scholar 

  10. 10.

    Van Rooij P, Martel A, Haesebrouck F, Pasmans F. Amphibian chytridiomycosis: a review with focus on fungus-host interactions. Vet Res. 2015;46(1):1–22.

    Article  CAS  Google Scholar 

  11. 11.

    Martel A, Spitzen-van der Sluijs A, Blooi M, Bert W, Ducatelle R, Fisher MC, et al. Batrachochytrium salamandrivorans sp. nov. causes lethal chytridiomycosis in amphibians. Proc Natl Acad Sci USA. 2013;110(38):15325–9

  12. 12.

    Thomas V, Wang Y, Van Rooij P, Verbrugghe E, Baláž V, Bosch J, et al. Mitigating Batrachochytrium salamandrivorans in Europe. Amphib Reptil. 2019;40(3):265–90.

    Article  Google Scholar 

  13. 13.

    Scheele BC, Pasmans F, Skerratt LF, Berger L, Martel A, Beukema W, et al. Amphibian fungal panzootic causes catastrophic and ongoing loss of biodiversity. Science. 2019;363:1459–63.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  14. 14.

    Lips KR. Overview of chytrid emergence and impacts on amphibians. Philos Trans R Soc B. 2016;371(1709):20150465.

    Article  Google Scholar 

  15. 15.

    Berger L, Speare R, Daszak P, Green DE, Cunningham AA, Goggin LC, et al. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc Natl Acad Sci USA. 1998;95(15):9031–6.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  16. 16.

    Voyles J, Young S, Berger L, Campbell C, Voyles WF, Dinudom A, et al. Pathogenesis of chytridiomycosis, a cause of catastrophic amphibian declines. Science. 2009;326:582–5.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  17. 17.

    Marantelli G, Berger L, Speare R, Keegan L. Distribution of the amphibian chytrid Batrachochytrium dendrobatidis and keratin during tadpole development. Pac Conserv Biol. 2004;10:173–9.

    Article  Google Scholar 

  18. 18.

    Blaustein AR, Romansic JM, Scheessele EA, Han BA, Pessier AP, Longcore JE. Interspecific variation in susceptibility of frog tadpoles to the pathogenic fungus Batrachochytrium dendrobatidis. Conserv Biol. 2005;19(5):1460–8.

    Article  Google Scholar 

  19. 19.

    Kilpatrick AM, Briggs CJ, Daszak P. The ecology and impact of chytridiomycosis: an emerging disease of amphibians. Trends Ecol Evol. 2010;25(2):109–18.

    PubMed  Article  PubMed Central  Google Scholar 

  20. 20.

    Walker SF, Bosch J, Gomez V, Garner TW, Cunningham AA, Schmeller DS, et al. Factors driving pathogenicity versus prevalence of the amphibian pathogen Batrachochytrium dendrobatidis and chytridiomycosis in Iberia. Ecol Lett. 2010;13:372–82.

    PubMed  Article  PubMed Central  Google Scholar 

  21. 21.

    Baláž V, Vörös J, Civiš P, Vojar J, Hettyey A, Sós E, et al. Assessing Risk and guidance on monitoring of Batrachochytrium dendrobatidis in Europe through identification of taxonomic selectivity of infection. Conserv Biol. 2014;28(1):213–23.

    PubMed  Article  PubMed Central  Google Scholar 

  22. 22.

    Conlon JM. The contribution of skin antimicrobial peptides to the system of innate immunity in anurans. Cell Tissue Res. 2011;343(1):201–12.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  23. 23.

    Daly JW. The chemistry of poisons in amphibian skin. Proc Natl Acad Sci USA. 1995;92(1):9–13.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  24. 24.

    Macfoy C, Danosus D, Sandit R, Jones TH, Garraffo MH, Spande TF, et al. Alkaloids of anuran skin: antimicrobial function? Zeitschrift für Naturforsch. 2005;60:932–7.

    CAS  Article  Google Scholar 

  25. 25.

    Gomes A, Giri B, Saha A, Mishra R, Dasgupta SC, Debnath A, et al. Bioactive molecules from amphibian skin: their biological activities with reference to therapeutic potentials for possible drug development. Indian J Exp Biol. 2007;45(7):579–93.

    CAS  PubMed  PubMed Central  Google Scholar 

  26. 26.

    Tempone AG, de Souza Carvalho Melhem M, Oliveira Prado F, Motoie G, Mitsuyoshi Hiramoto R, Maria Antoniazzi M, et al. Amphibian secretions for drug discovery studies: a search for new antiparasitic and antifungal compounds. Lett Drug Des Discov. 2007;4(1):67–73.

  27. 27.

    König E, Bininda-Emonds ORP, Shaw C. The diversity and evolution of anuran skin peptides. Peptides. 2015;63:96–117.

    CAS  Article  PubMed  PubMed Central  Google Scholar 

  28. 28.

    Chinchar GV, Wang J, Murti G, Carey C, Rollins-Smith LA. Inactivation of frog virus 3 and channel catfish virus by esculentin-2P and ranatuerin-2P, two antimicrobial peptides isolated from frog skin. Virology. 2001;288(2):351–7.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  29. 29.

    Zasloff M. Antimicrobial peptides of multicellular organisms. Nature. 2002;415:389–95.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  30. 30.

    Rollins-Smith LA, Conlon JM. Antimicrobial peptide defenses against chytridiomycosis, an emerging infectious disease of amphibian populations. Dev Comp Immunol. 2005;29(7):589–98.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  31. 31.

    Conlon JM, Mechkarska M, Lukic ML, Flatt PR. Potential therapeutic applications of multifunctional host-defense peptides from frog skin as anti-cancer, anti-viral, immunomodulatory, and anti-diabetic agents. Peptides. 2014;57:67–77.

    CAS  Article  PubMed  PubMed Central  Google Scholar 

  32. 32.

    Woodhams DC, Rollins-Smith LA, Alford RA, Simon MA, Harris RN. Innate immune defenses of amphibian skin: antimicrobial peptides and more. Anim Conserv. 2007;10(4):425–8.

    Article  Google Scholar 

  33. 33.

    Tennessen JA, Woodhams DC, Chaurand P, Reinert LK, Billheimer D, Shyr Y, et al. Variations in the expressed antimicrobial peptide repertoire of northern leopard frog (Rana pipiens) populations suggest intraspecies differences in resistance to pathogens. Dev Comp Immunol. 2009;33(12):1247–57.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  34. 34.

    Hayes RA, Crossland MR, Hagman M, Capon RJ, Shine R. Ontogenetic variation in the chemical defenses of cane toads (Bufo marinus): toxin profiles and effects on predators. J Chem Ecol. 2009;35(4):391–9.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  35. 35.

    Ujszegi J, Móricz ÁM, Krüzselyi D, Hettyey A. Skin toxin production of toads changes during early ontogeny but is not adjusted to the microbiota of the aquatic environment. Evol Ecol. 2017;31(6):925–36.

    Article  Google Scholar 

  36. 36.

    Üveges B, Fera G, Móricz ÁM, Krüzselyi D, Bókony V, Hettyey A. Age- and environment-dependent changes in chemical defences of larval and post-metamorphic toads. BMC Evol Biol. 2017;17:137.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  37. 37.

    Cunha Filho GA, Schwartz CA, Resck IS, Murta MM, Lemos SS, Castro MS, et al. Antimicrobial activity of the bufadienolides marinobufagin and telocinobufagin isolated as major components from skin secretion of the toad Bufo rubescens. Toxicon. 2005;45(6):777–82.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  38. 38.

    Barnhart K, Forman ME, Umile TP, Kueneman J, McKenzie V, Salinas I, et al. Identification of bufadienolides from the boreal toad, Anaxyrus boreas, active against a fungal pathogen. Microb Ecol. 2017;74:990–1000.

    CAS  Article  PubMed  PubMed Central  Google Scholar 

  39. 39.

    Conlon JM, Seidel B, Nielsen PF. An atypical member of the brevinin-1 family of antimicrobial peptides isolated from the skin of the European frog Rana dalmatina. Comp Biochem Physiol C. 2004;137(2):191–6.

    Google Scholar 

  40. 40.

    Mebs D, Wagner MG, Pogoda W, Maneyro R, Kwet A, Kauert G. Lack of bufadienolides in the skin secretion of red bellied toads, Melanophryniscus spp. (Anura, Bufonidae), from Uruguay. Comp Biochem Physiol C. 2007;144(4):398–402.

  41. 41.

    Delfino G, Brizzi R, Alvarez BB. Serous cutaneous glands in Phyllomedusa hypochondrialis (Anura, Hylidae): secretory patterns during ontogenesis. Tissue Cell. 1998;30(1):30–40.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  42. 42.

    Viertel B, Richter S. Anatomy: viscera and endocrines. In: McDiarmid RW, Altig R, editors. Tadpoles: the biology of anuran larvae. Chicago: University of Chicago Press; 1999. p. 92–148.

    Google Scholar 

  43. 43.

    Lacombe C, Cifuentes-Diaz C, Dunia I, Auber-Thomay M, Nicolas P, Amiche M. Peptide secretion in the cutaneous glands of South American tree frog Phyllomedusa bicolor: an ultrastructural study. Eur J Cell Biol. 2000;79(9):631–41.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  44. 44.

    Terreni A, Nosi D, Greven H, Delfino G. Development of serous cutaneous glands in Scinax nasica (Anura, Hylidae): patterns of poison biosynthesis and maturation in comparison with larval glands in specimens of other families. Tissue Cell. 2003;35:274–87.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  45. 45.

    Quagliata S, Malentacchi C, Delfino C, Brunasso AMG, Delfino G. Adaptive evolution of secretory cell lines in vertebrate skin. Caryologia. 2006;59(2):187–206.

    Article  Google Scholar 

  46. 46.

    Flajnik MF, Hsu E, Kaufman JF, Du Pasquier L. Changes in the immune system during metamorphosis of Xenopus. Immunol Today. 1987;8(2):58–64.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  47. 47.

    Schadich E, Cole ALJ, Squire M, Mason D. Skin peptides of different life stages of Ewing’s tree frog. J Exp Zool A. 2010;313A(8):532–7.

  48. 48.

    Wabnitz PA, Walters H, Tyler MJ, Wallace JC, Bowie JH. First record of host defence peptides in tadpoles. The magnificent tree frog Litoria splendida. J Pept Res. 1998;52(6):477–81.

  49. 49.

    Woodhams DC, Bell SC, Bigler L, Caprioli RM, Chaurand P, Lam BA, et al. Life history linked to immune investment in developing amphibians. Conserv Physiol. 2016;4(1):cow025.

  50. 50.

    Gosner KL. A simplified table for staging anuran embryos larvae with notes on identification. Herpetologica. 1960;16(3):183–90.

    Google Scholar 

  51. 51.

    Hettyey A, Tóth Z, Van Buskirk J. Inducible chemical defences in animals. Oikos. 2014;23:1025–8.

    Article  Google Scholar 

  52. 52.

    Groner ML, Rollins-Smith LA, Reinert LK, Hempel J, Bier ME, Relyea RA. Interactive effects of competition and predator cues on immune responses of leopard frogs at metamorphosis. J Exp Biol. 2014;217(October):351–8.

    PubMed  PubMed Central  Google Scholar 

  53. 53.

    Simmaco M, Mangoni ML, Boman A, Barra D, Boman HG. Experimental infections of Rana esculenta with Aeromonas hydrophila: a molecular mechanism for the control of the normal flora. Scand J Immunol. 1998;48(4):357–63.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  54. 54.

    Mangoni ML, Miele R, Renda TG, Barra D, Simmaco M. The synthesis of antimicrobial peptides in the skin of Rana esculenta is stimulated by microorganisms. FASEB J. 2001;15:1431–2.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  55. 55.

    Woodhams DC, Bigler L, Marschang R. Tolerance of fungal infection in European water frogs exposed to Batrachochytrium dendrobatidis after experimental reduction of innate immune defenses. BMC Vet Res. 2012;8:197–209.

    PubMed  PubMed Central  Article  Google Scholar 

  56. 56.

    Woodhams DC, Geiger CC, Reinert LK, Rollins-Smith LA, Lam B, Harris RN, et al. Treatment of amphibians infected with chytrid fungus: learning from failed trials with itraconazole, antimicrobial peptides, bacteria, and heat therapy. Dis Aquat Organ. 2012;98(1):11–25.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  57. 57.

    Fites JS, Ramsey JP, Holden WM, Collier SP, Sutherland DM, Reinert LK, et al. The invasive chytrid fungus of amphibians paralyzes lymphocyte responses. Science. 2013;342(6156):366–9.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  58. 58.

    Rollins-Smith LA, Fites JS, Reinert LK, Shiakolas AR, Umile TP, Minbiole KP. Immunomodulatory metabolites released by the frog-killing fungus Batrachochytrium dendrobatidis. Infect Immun. 2015;83(12):4565–70.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  59. 59.

    Grogan LF, Robert J, Berger L, Skerratt LF, Scheele BC, Castley JG, et al. Review of the amphibian immune response to chytridiomycosis, and future directions. Front Immunol. 2018;9:2536.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  60. 60.

    Gabor CR, Fisher MC, Bosch J. A non-invasive stress assay shows that tadpole populations infected with Batrachochytrium dendrobatidis have elevated corticosterone levels. PLoS ONE. 2013;8(2):e56054.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  61. 61.

    Gabor CR, Fisher MC, Bosch J. Elevated corticosterone levels and changes in amphibian behavior are associated with Batrachochytrium dendrobatidis (Bd) infection and Bd lineage. PLoS ONE. 2015;10(4):1–13.

    Article  CAS  Google Scholar 

  62. 62.

    Simmaco M, Boman A, Mangoni ML, Mignogna G, Miele R, Barra D, et al. Effect of glucocorticoids on the synthesis of antimicrobial peptides in amphibian skin. FEBS Lett. 1997;416(3):273–5.

    CAS  Article  PubMed  PubMed Central  Google Scholar 

  63. 63.

    Rollins-Smith LA, Ramsey JP, Pask JD, Reinert LK, Woodhams DC. Amphibian immune defenses against chytridiomycosis: impacts of changing environments. Integr Comp Biol. 2011;51(4):552–62.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  64. 64.

    Garner TWJ, Walker S, Bosch J, Leech S, Rowcliffe JM, Cunningham AA, et al. Life history tradeoffs influence mortality associated with the amphibian pathogen Batrachochytrium dendrobatidis. Oikos. 2009;118(5):783–91.

    Article  Google Scholar 

  65. 65.

    Luquet E, Garner TW, Léna JP, Bruel C, Joly P, Lengagne T, et al. Genetic erosion in wild populations makes resistance to a pathogen more costly. Evolution (N Y). 2012;66(6):1942–52.

    Google Scholar 

  66. 66.

    Sheldon BC, Verhulst S. Ecological immunology: costly parasite defences and trade-offs in evolutionary ecology. Trends Ecol Evol. 1996;11(8):317–21.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  67. 67.

    McMahon TA, Brannelly LA, Chatfield MWH, Johnson PTJ, Joseph MB, McKenzie VJ, et al. Chytrid fungus Batrachochytrium dendrobatidis has nonamphibian hosts and releases chemicals that cause pathology in the absence of infection. Proc Natl Acad Sci USA. 2013;110(1):210–5.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  68. 68.

    Lichtstein D, Gati I, Babila T, Haver E, Katz U. Effect of salt acclimation on digitalis-like compounds in the toad. Biochim Biophys Acta. 1991;1073:65–8.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  69. 69.

    Lichtstein D, Gati I, Haver E, Katz U. Digitalis-like compounds in the toad Bufo viridis: tissue and plasma levels and significance in osmotic stress. Life Sci. 1992;51:119–28.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  70. 70.

    Toledo RC, Jared C. Cutaneous granular glands and amphibian venoms. Comp Biochem Physiol A. 1995;111(1):1–29.

    Article  Google Scholar 

  71. 71.

    Hayes RA, Piggott AM, Dalle K, Capon RJ. Microbial biotransformation as a source of chemical diversity in cane toad steroid toxins. Bioorg Med Chem Lett. 2009;19(6):1790–2.

    CAS  Article  PubMed  PubMed Central  Google Scholar 

  72. 72.

    Tempone AG, Pimenta DC, Lebrun I, Sartorelli P, Taniwaki NN, de Andrade HF, et al. Antileishmanial and antitrypanosomal activity of bufadienolides isolated from the toad Rhinella jimi parotoid macrogland secretion. Toxicon. 2008;52(1):13–21.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  73. 73.

    Kleinhenz P, Boone MD, Fellers G. Effects of the amphibian Chytrid fungus and four insecticides on Pacific Treefrogs (Pseudacris regilla). J Herpetol. 2012;46(4):625–31.

    Article  Google Scholar 

  74. 74.

    Hanlon SM, Lynch KJ, Kerby J, Parris MJ. Batrachochytrium dendrobatidis exposure effects on foraging efficiencies and body size in anuran tadpoles. Dis Aquat Organ. 2015;112(3):237–42.

    PubMed  Article  PubMed Central  Google Scholar 

  75. 75.

    Venesky MD, Parris MJ, Storfer A. Impacts of Batrachochytrium dendrobatidis infection on tadpole foraging performance. EcoHealth. 2009;6(4):565–75.

    PubMed  Article  PubMed Central  Google Scholar 

  76. 76.

    Venesky MD, Hanlon SM, Lynch K, Parris MJ, Rohr JR. Optimal digestion theory does not predict the effect of pathogens on intestinal plasticity. Biol Lett. 2013;9(2):20130038.

    PubMed  PubMed Central  Article  Google Scholar 

  77. 77.

    Parris MJ, Cornelius TO. Fungal pathogen causes competitive and developmental stress in larval amphibian communities. Ecology. 2004;85(12):3385–95.

    Article  Google Scholar 

  78. 78.

    McMahon TA, Rohr JR. Transition of Chytrid fungus infection from mouthparts to hind limbs during Amphibian metamorphosis. EcoHealth. 2015;12(1):188–93.

    PubMed  Article  PubMed Central  Google Scholar 

  79. 79.

    Fisher MC, Bosch J, Yin Z, Stead DA, Walker J, Selway L, et al. Proteomic and phenotypic profiling of the amphibian pathogen Batrachochytrium dendrobatidis shows that genotype is linked to virulence. Mol Ecol. 2009;18(3):415–29.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  80. 80.

    Bosch J, Martínez-Solano I. Chytrid fungus infection related to unusual mortalities of Salamandra salamandra and Bufo bufo in the Peñalara Natural Park, Spain. Oryx. 2006;40(1):84–9.

    Article  Google Scholar 

  81. 81.

    Ficetola GF, Valentini A, Miaud C, Noferini A, Mazzotti S, Dejean T. Batrachochytrium dendrobatidis in amphibians from the Po River Delta, Northern Italy. Acta Herpetol. 2011;6(2):297–302.

    Google Scholar 

  82. 82.

    Vörös J, Herczeg D, Fülöp A, Gál J, Dán Á, Harmos K, et al. Batrachochytrium dendrobatidis in Hungary: an overview of recent and historical occurrence. Acta Herpetol. 2018;13:125–40.

    Google Scholar 

  83. 83.

    APHA. Standard methods for the examination of water and wastewater. 16. Washington, D.C: American Public Health Association; 1985. 1268 p.

  84. 84.

    Bishop PJ, Speare R, Poulter R, Butler M, Speare BJ, Hyatt A, et al. Elimination of the amphibian chytrid fungus Batrachochytrium dendrobatidis by Archey’s frog Leiopelma archeyi. Dis Aquat Organ. 2009;84(1):9–15.

    PubMed  Article  PubMed Central  Google Scholar 

  85. 85.

    Young S, Speare R, Berger L, Skerratt LF. Chloramphenicol with fluid and electrolyte therapy cures terminally ill green tree frogs (Litoria caerulea) with chytridiomycosis. J Zoo Wildl Med. 2012;43(2):330–7.

    PubMed  Article  PubMed Central  Google Scholar 

  86. 86.

    Ujszegi J, Molnár K, Hettyey A. How to disinfect anuran eggs? Sensitivity of anuran embryos to chemicals widely used for the disinfection of larval and post-metamorphic amphibians. J Appl Toxicol. 2021;41:387–98.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  87. 87.

    Shine R, Amiel J, Munn AJ, Stewart M, Vyssotski AL, Lesku JA. Is “cooling then freezing” a humane way to kill amphibians and reptiles? Biol Open. 2015;4(7):760–3.

    PubMed  PubMed Central  Article  Google Scholar 

  88. 88.

    Garner TW, Rowcliffe JM, Fisher MC. Climate change, chytridiomycosis or condition: an experimental test of amphibian survival. Glob Change Biol. 2011;17(2):667–75.

    Article  Google Scholar 

  89. 89.

    Johnson ML, Berger L, Philips L, Speare R. Fungicidal effects of chemical disinfectants, UV light, desiccation and heat on the amphibian chytrid Batrachochytrium dendrobatidis. Dis Aquat Organ. 2003;57(3):255–60.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  90. 90.

    Boyle DG, Boyle DB, Olsen V, Morgan JAT, Hyatt AD. Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Dis Aquat Organ. 2004;60(2):141–8.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  91. 91.

    Kriger KM, Hero J-M, Ashton KJ. Cost efficiency in the detection of chytridiomycosis using PCR assay. Dis Aquat Organ. 2006;71:149–54.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  92. 92.

    Woodhams DC, Rollins-smith LA, Carey C, Reinert L, Tyler MJ, Alford RA. Population trends associated with skin peptide defenses against Chytridiomycosis in Australian frogs. Oecologia. 2006;146(4):531–40.

    PubMed  Article  PubMed Central  Google Scholar 

  93. 93.

    Conlon JM. Purification of naturally occurring peptides by reversed-phase HPLC. Nat Protoc. 2007;2(1):191–7.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  94. 94.

    Hagman M, Hayes RA, Capon RJ, Shine R. Alarm cues experienced by cane toad tadpoles affect post-metamorphic morphology and chemical defences. Funct Ecol. 2009;23:126–32.

    Article  Google Scholar 

  95. 95.

    Grafen A, Hails R. Modern statistics for the life sciences. Oxford, New York: Oxford University Press; 2002. p. 351.

    Google Scholar 

  96. 96.

    Engqvist L. The mistreatment of covariate interaction terms in linear model analyses of behavioural and evolutionary ecology studies. Anim Behav. 2005;70(4):967–71.

    Article  Google Scholar 

Download references


We thank T. Sendula, Zs. Mikó, M. Szederkényi for assistance during the experiment, B. Dalmadi Kiss for help in analytical work, the Pilisi Parkerdő Zrt. for allowing us to use their roads, and V. Verebélyi for the graphics depicting tadpoles and metamorphs. Zoospore genomic equivalent standards were kindly offered by J. Bosch.


Research was supported by the Lendület Programme of the Hungarian Academy of Sciences (MTA, LP2012-24/2012), an FP7 Marie Curie Career Integration Grant (PCIG13-GA-2013-631722), a János Bolyai Scholarship of the MTA, a Bolyai + Scholarship of the New National Excellence Programme of the Ministry of Human Capacities, Hungary (ÚNKP, EMMI), the Otto Kinne Foundation and the National Research, Development and Innovation Office (NKFIH) of Hungary (Grant No. K-124375). These funding bodies provided the financial support necessary for designing, preparation and performance of the study and also supported authors during molecular, analytical and statistical analyses and during the writing of the manuscript. TWJG was funded by NERC (Standard Grants NE/S000992/1, NE/N009967/1 and NE/K012509/1) which provided support during designing the study and writing the manuscript.

Author information




JU, JV, TWJG and AH conceived the ideas and designed methodology; JU, TD and AH accomplished the experiment; KL, ÁMM, DK and LD performed and evaluated chemical analytical measurements. JU, MZN and JV accomplished qPCR analyses. JU and AH analysed the data and led the writing of the manuscript. All authors contributed critically to the drafts and gave final approval for publication. All authors read and approved the final manuscript.

Corresponding author

Correspondence to János Ujszegi.

Ethics declarations

Ethics approval and consent to participate

The Közép-Duna-Völgyi KTVF issued the permission to conduct the study (PE/KTF:3596-6-8/2016) and the Ethical Commission of the ATK NÖVI approved the investigation in accordance with Good Scientific Practice guidelines and national legislation.

Consent for publication

Not applicable.

Competing interest

The authors have no conflict of interest to declare.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Ujszegi, J., Ludányi, K., Móricz, Á.M. et al. Exposure to Batrachochytrium dendrobatidis affects chemical defences in two anuran amphibians, Rana dalmatina and Bufo bufo. BMC Ecol Evo 21, 135 (2021).

Download citation


  • Antimicrobial peptide
  • Bufadienolide
  • Indirect effect
  • Infectious diseases
  • Innate immunity