A review of chemical defense in harlequin toads (Bufonidae: Atelopus) (2024)

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  We review the quantity and diversity of toxins in Atelopus toads.

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  Sampling and methodological biases likely affect known toxin quantity and diversity.

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  Atelopus extinctions threaten the loss of undescribed toxins.

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  Few data exist on the ecology and evolution of Atelopus chemical defenses.

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  Atelopus is a promising study system for toxin sequestration and synthesis.

2.1. Literature review

We examined peer-reviewed literature published prior to November 2021 describing the composition and toxicity of Atelopus chemical defenses as well as auxiliary literature that describes the pharmacology of relevant toxins, toxin detection and quantification methods, and Atelopus ecology, morphology, taxonomy, and evolution. Articles were found using keyword searches through the UC Berkeley Library, Google Scholar, and Google Search with phrases such as “Atelopus toxic,” “Atelopus peptides,” “atelopidtoxin,” “chiriquitoxin,” “zetekitoxin,” “Atelopus bufadienolides,” etc. An exhaustive search was performed specifically for literature detailing the detection, quantification, and identification of Atelopus small-molecule toxins; in total, seventeen peer-reviewed papers were identified that met one or more of these criteria (see Supplementary Table S1 for a complete list). Becker et al. (2011) claims to have detected zetekitoxins in Atelopus zeteki via HPLC, a method which would require a zetekitoxin AB standard for positive identification (Table 1). Given the extraordinary rarity of purified zetekitoxin AB (Yotsu-Yamash*ta et al., 2004), it seems highly unlikely Becker et al. (2011) had access to sufficient quantities for use as an HPLC standard. Thus, we exclude this paper from our analyses. Additionally, we reviewed a PhD thesis (Brown, 1972) describing toxicity assessments of several Atelopus species, as well as the isolation and detection of guanidinium alkaloids in A. zeteki. Some of the data presented therein appears to have been published elsewhere (Brown et al., 1977; Fuhrman et al., 1969), but we include the unpublished data from Brown (1972) in our analyses. Two additional papers were identified that detailed the presence or absence of skin peptides produced by Atelopus that may or may not be used in defense (Ellison et al., 2014; Woodhams et al., 2006). Owing to a lack of information on Atelopus skin peptide diversity and function, we focus our review on guanidinium alkaloids and cardiac glycosides.

2.2. Geographic and phylogenetic mapping of Atelopus toxin profiles

Sixteen of the eighteen Atelopus toxin assessment papers (including Brown, 1972) compiled during literature review described sampling location. In a few papers, only country-level sampling locations were provided or the species identification was dubious, so we excluded some of these samples from our combined assessments (see Table S2 for details). When GPS coordinates were not provided, we obtained coordinates using the geocoding service provided by Google Maps (https://developers-dot-devsite-v2-prod.appspot.com/maps/documentation/utils/geocoder). If the specific location name provided in a paper was not available, coordinates were determined by inputting larger geographic regions known to contain the locations of interest. See Supplementary Table S2 for a complete inventory of sampling locations, location names, and coordinates. Maps were generated through the ArcGIS Online application, Map Viewer Classic (Esri, Redlands, CA, USA), and edited using Adobe Illustrator (Adobe Inc., 2021).

To visualize the phylogenetic distribution of Atelopus toxins (Fig. 3a), we obtained a chronogram of Atelopus species from Ramírez et al. (2020) and pruned it to include a single tip per species in R v3.6.1 (R Core Team, 2019) using packages phytools v0.7.70 (Revell, 2012) and ape v5.5 (Paradis and Schliep, 2019).

4.1. Guanidinium alkaloids

Guanidinium alkaloids are low molecular weight neurotoxins that target voltage-gated sodium channels (VGSCs). The eponymous positively charged guanidinium moiety (Fig. 1) interacts with the extracellular facing end of the sodium ion channel, while the rest of the molecule effectively seals off the pore (Narahashi, 2008). With the flow of sodium ions occluded, nerves lose the ability to produce action potentials and thus can no longer send signals (Narahashi et al., 1964). Guanidinium alkaloid poisoning is characterized by tingling, ataxia, paralysis, and death by respiratory failure or bradycardia (Durán-Riveroll and Cembella, 2017; How et al., 2003).

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Fig. 1

Guanidinium alkaloids detected in Atelopus. Purified quantities of Zetekitoxin C have been insufficient to estimate chemical structure (Brown et al., 1977).

Although guanidinium alkaloids have been detected in many marine animals (Chau et al., 2011), their occurrence in terrestrial taxa is limited to five amphibian families: Salamandridae, Dendrobatidae (Colostethus), Brachycephalidae, Rhacophoridae (Polypedates), and Bufonidae (Atelopus; Daly et al., 1994; Kim et al., 1975; Lüddecke et al., 2018; Pires et al., 2005; Tanu et al., 2001). Tetrodotoxin has also been reported in a single species of the salamander family Ambystomatidae (Yotsu et al., 1990a), however that finding has since been called into question (Hanifin, 2010). Lastly, tetrodotoxin has been suggested to cooccur with bufadienolides in Clinotarsus cultripes (Gosavi et al., 2014), a ranid, and chiriquitoxin has been suggested to occur in Hypsiboas crepitans, a hylid (Lamadrid-Feris et al., 2015); however, these findings are based on preliminary data that have not been verified by more sensitive techniques. Five guanidinium alkaloids have been detected and structurally identified in Atelopus: tetrodotoxin, 4,9-anhydrotetrodotoxin, 4-epitetrodotoxin, chiriquitoxin, and zetekitoxin AB (Fig. 1). While not structurally identified, zetekitoxin C has been detected in Atelopus zeteki and is likely also a guanidinium alkaloid (Brown et al., 1977).

4.2. Bufadienolides

Bufadienolides are cardiac glycosides (CGs), steroidal toxins that bind to and inhibit Na⁺/K⁺-ATPases (Fig. 2; Botelho et al., 2019). Na⁺/K⁺-ATPase inhibition ultimately causes a buildup of Ca²⁺ ions within nerve and muscle cells, which increases the contractility of muscle tissues (Blaustein et al., 2009). CG poisoning manifests as hypertension, gastrointestinal distress, abnormal heart rate, and – in high enough doses – death (Roberts et al., 2016). CG inhibition of Na⁺/K⁺-ATPase also alters some signaling pathways and is the topic of intense research for potential anticancer therapies (Reddy et al., 2020). Whereas other CGs have a five membered lactone ring attached to the central steroid structure, bufadienolides are characterized by a six membered lactone ring (Fig. 2; Rodríguez et al., 2017).

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Fig. 2

Bufadienolides detected in Atelopus.

Source. All four Atelopus species that have been tested for bufadienolides were found to possess this class of toxins (Daly et al., 1997; Flier et al., 1980). Bufadienolides are endogenously synthesized by toads, likely from cholesterol (Chiadao and Osuch, 1969; Garraffo and Gros, 1986; Porto and Gros, 1971). Interestingly, bufadienolides and other CGs are present at low levels in mammal and amphibian tissues, and likely have a highly conserved role as endogenous hormones (Dmitrieva et al., 2000; Flier et al., 1980; Lenaerts et al., 2018; Schoner and Scheiner-Bobis, 2005). Bufadienolides may also be used for sodium and water regulation in toads. For example, exposure to saline solutions altered the concentration of digitalis-like compounds (likely bufadienolides, see Dufresnes et al., 2019; Rodríguez et al., 2017) in the skin and brain of Bufotes viridis (Lichtstein et al., 1991). Thus, a possible evolutionary pathway for bufadienolide defense in toads is via natural selection on the regulation of endogenous CGs (Flier et al., 1980) coupled with the development of Na⁺/K⁺-ATPase target site insensitivity, whereby amino acid substitutions result in a weaker affinity of Na⁺/K⁺-ATPase for CGs. Target site insensitivity to CGs has been demonstrated in the α3 Na⁺/K⁺-ATPase subunit of bufonid toads – including Atelopus spumarius – and toad-feeding reptiles (Moore et al., 2009; Ujvari et al., 2015) and in a tandem duplicate of the α1 Na⁺/K⁺-ATPase in toad-feeding frogs (Leptodactylus;Mohammadi et al., 2021). More than one hundred different bufadienolides have been detected in the skins, eggs, or granular gland secretions of bufonid toads (Rodríguez et al., 2017). The mechanisms underlying the diversity of bufadienolides in toads has been largely uninvestigated, though microbial biotransformation may play a role (Hayes et al., 2009b; Kamalakkannan et al., 2017).

4.3. Unidentified toxins

Toxin diversity in Atelopus is incompletely characterized, and toxins whose identities are unknown have been detected in multiple species. For various reasons, including small quantities and methodological limitations, investigation into these chemicals has been insufficient to clarify their structures, pharmacology, and/or chemical characteristics (see Section 5.2 for a discussion of the methods used to identify Atelopus toxins).

Several unidentified toxins that mirror guanidinium alkaloids in effect or chemistry have been detected in Atelopus. The only toxin found in A. certus is water soluble and likely positively charged, both of which are features of guanidinium alkaloids. While this unknown chemical was determined to not be TTX, too little was purified for further analysis (Yotsu-Yamash*ta and Tateki, 2010). In competitive binding assays, A. spurrelli skin extracts inhibit saxitoxin binding, a characteristic of guanidinium alkaloids. Given that TTX is a minor component of A. spurrelli skin extracts, one or more unidentified TTX-like toxins are believed responsible for A. spurrelli toxicity (Daly et al., 1994). Similarly, TTX is a trace component in A. “ignescens,” and the tetrodotoxin-like chemicals which account for the remaining toxicity of A. “ignescens” skin extracts to mice are uncharacterized (Daly et al., 1994). Finally, aqueous A. senex skin extracts injected into mice caused the same symptoms as known guanidinium alkaloids (Brown, 1972). While A. senex skin extracts likely contain guanidinium alkaloids, the individual identities of these toxins have not been determined.

There are unidentified Atelopus toxins which either differ substantially from guanidinium alkaloids or whose properties are almost completely unknown. Aqueous skin extracts of A. planispina injected in mice cause symptoms that differ from those of guanidinium alkaloid poisoning, specifically cessation of respiration before cardiac arrest (Fuhrman et al., 1969). The unidentified toxin is unlikely a bufadienolide because bufadienolides are weakly soluble in water (Flier et al., 1980) and do not cause the symptoms observed with A. planispina toxins (Roberts et al., 2016). Thus, A. planispina represents a likely source of novel Atelopus toxins, which warrants further research. Secondly, an unidentified major toxin has been detected in a single specimen of A. zeteki and has received no further investigation (Yotsu-Yamash*ta and Tateki, 2010). However, the method used on that specimen was incapable of detecting ZTX AB, the most common major toxin found in A. zeteki (Table S2; Yotsu-Yamash*ta and Tateki, 2010), so it is plausible that the chemical was ZTX AB.

5.1. Extraction and purification

In most studies, extractions are performed on isolated skin or eggs, though whole-body extractions are also possible (Mebs et al., 2018a; Yotsu-Yamash*ta et al., 1992). Usually, tissues are broken into small pieces and suspended in a solvent with properties most amenable to the toxin type of interest. If the tissues are hom*ogenized, subsequent dialysis is performed to separate soluble chemicals from the slurry (Fuhrman et al., 1969; Pavelka et al., 1977; Shindelman et al., 1969). A variety of extract cleaning methods can be used, many of which involve some form of filtration via chromatography (Daly et al., 1994; Mebs and Schmidt, 1989; Shindelman et al., 1969). Final toxin separation and purification may be performed through chromatography or free-flowing electrophoresis (Brown et al., 1977; Yotsu-Yamash*ta et al., 2004).

A study published in 1977 found that higher levels of guanidinium alkaloids were extracted from A. chiriquiensis eggs and skin when 3% acetic acid was used as opposed to water, with the effect most pronounced in egg extractions (Pavelka et al., 1977). Following acid extraction, the toxins exhibited enhanced solubility in water. The authors suggest guanidinium alkaloids in Atelopus may exist to some extent in an insoluble bound form, from which the toxins are released following hydrolysis with acid (Pavelka et al., 1977). Several species of TTX-possessing pufferfish (Matsui et al., 2000; Matsumoto et al., 2010; Yotsu-Yamash*ta et al., 2001) and gastropods (Hwang et al., 2007) are known to possess TTX-binding proteins, thus another possibility is the acidic denaturation of a guanidinium alkaloid-binding protein. Previous studies (before 1977) used distilled water for the initial extraction, and thus may have reported lower toxin levels than were present in the toads tested. With a few exceptions (Daly et al., 1994, 1997; Mebs et al., 1995), subsequent studies on Atelopus toxins followed Pavelka et al. (1977) and performed acidic extractions.

While bufadienolides have a variety of structures and physical properties (Rodríguez et al., 2017), they tend to be poorly soluble in water (Flier et al., 1980; Li et al., 2009; Zhang et al., 2008). Furthermore, bufadienolides degrade over 24-h timescales in highly acidic or basic solutions (Li et al., 2015). Thus, aqueous and/or acidic extractions of Atelopus tissues may largely exclude bufadienolides and mouse bioassays of such extractions likely do not account of the contribution of bufadienolides to Atelopus toxicity. Bufadienolide-specific toad extractions commonly employ methanol as a solvent (Barnhart et al., 2017; Daly et al., 1997; Flier et al., 1980; Inoue et al., 2020; Petroselli et al., 2018).

Considering the severity of Atelopus declines (La Marca et al., 2005), nonfatal extraction methods may be critical for future research on toxins in wild Atelopus populations. One method involves collecting small skin punches from animals in the field, and has been utilized to measure TTX levels in salamanders but has not been benchmarked yet for accuracy against whole-body extractions (Bucciarelli et al., 2014; Hanifin et al., 2002). Completely noninvasive methods involve the collection of granular gland secretion via manual or electrical stimulation of amphibian skin (Conceição et al., 2007; Rozek et al., 1998). Although these methods have not been thoroughly tested on amphibians that possess guanidinium alkaloids, we suspect that they would be fruitful. The sampling of museum specimens for toxins represents another avenue for Atelopus research and could enable the assessment of species that have gone extinct. In a couple of studies, analyses were performed on the storage alcohol of Atelopus museum specimens (Mebs et al., 1995, 2018a). However, toxins in museum specimens may degrade over time and specimens stored in formalin are not suitable for toxin analyses (Mebs et al., 1995).

5.2. Toxin identification

After extraction and purification, a variety of methods can be applied to determine guanidinium alkaloid identity (Table 1). Thin layer chromatography (TLC) and nuclear magnetic resonance (NMR) were among the earliest and most used of these in Atelopus studies. TLC separates chemicals and allows for assessment of purity. Spots on a TLC plate can be sprayed with the Weber reagent, an aqueous solution of sodium nitroprusside, potassium ferricyanide, and sodium hydroxide (Weber, 1928) that turns red in the presence of fifty or more mouse units of guanidinium alkaloids (approximately equivalent to 11μg of TTX; Brown et al., 1977). Alternatively, the spotted TLC plate can be sprayed with an alkaline solution and heated (Mebs and Schmidt, 1989). This converts guanidinium alkaloids into 2-aminoquinazoline derivatives that fluoresce under UV light (Nakamura and Yasumoto, 1985). 1H NMR and C-13 NMR spectra give information on the electronic environments, neighboring atoms, and quantities of carbon and hydrogen atoms in a molecule, respectively (Klein, 2017). In addition to serving as a method of detection, NMR has been critical in determining the chemical structures of Atelopus guanidinium alkaloids (Yotsu-Yamash*ta et al., 2004; Yotsu et al., 1990b).

The next technology that became widely used in Atelopus toxin studies was developed by Yotsu et al. (1989). This liquid chromatography-fluorescence detection system (LC-FLD) takes advantage of the fluorescence of 2-aminoquinazoline derivatives (generated by heating guanidinium alkaloids in alkaline solutions) to identify guanidinium alkaloids that have been separated by liquid chromatography, and was the first method used to detect 4-epiTTX and 4,9-anhydroTTX in Atelopus extracts (Yotsu-Yamash*ta et al., 1992; Yotsu et al., 1989). LC-FLD utilizes reverse phase chromatography, which is incapable of separating all TTX analogs (Bane et al., 2014). Additionally, because TTX analogs exhibit a wide range of fluorescent intensities, analogs may not all be detectable under the same set of conditions (Shoji et al., 2001). For instance, detection of 11-deoxytetrodotoxin, a common analog in newts and brachycephalid toads (Hanifin, 2010), requires higher post column reaction temperatures than does the detection of TTX (Yotsu et al., 1989). Studies of Atelopus toxins which utilized LC-FLD either employed post-column reaction temperatures below those needed for 11-deoxytetrodotoxin detection (Yotsu-Yamash*ta et al., 1992) or neglect to specify the post-column reaction temperature (Daly et al., 1994; Mebs et al., 1995; Yotsu-Yamash*ta and Tateki, 2010). All Atelopus LC-FLD studies utilized a C18 chromatography column but varied in mobile phase composition (heptafluorobutyric acid or trifluoroacetic acid) and concentration of NaOH in the post-column reaction (ranging from 2.8N to 4N; Daly et al., 1994; Mebs et al., 1995; Yotsu-Yamash*ta et al., 1992, Yotsu-Yamash*ta and Tateki, 2010). LC-FLD cannot detect low levels of CHTX, and it is incapable of detecting ZTX AB (Yotsu-Yamash*ta et al., 1992; Yotsu-Yamash*ta and Tateki, 2010). Four Atelopus species have had their guanidinium alkaloids studied exclusively through LC-FLD: A. “ignescens,” A. spumarius sensu lato, A. spurrelli, and A. subornatus (Daly et al., 1994; Mebs et al., 1995). Thus, these species could have undetected CHTX, ZTX AB, or 11-deoxytetrodotoxin.

Most recently, methods that incorporate electrospray ionization-mass spectrometry (ESI-MS) have emerged as promising guanidinium alkaloid assays in Atelopus studies (Mebs et al., 1995, 2018a; Yotsu-Yamash*ta and Tateki, 2010). Whereas quantification by LC-FLD requires calibration with standards of each TTX analog being measured, quantification of TTX analogs can be performed by ESI-MS with the exclusive use of a TTX standard (Chen et al., 2011, Nakagawa et al., 2006, Shoji et al., 2001). The simplest use of ESI-MS is to directly screen toad extracts for ions indicative of toxin presence, such as the MH+ion of TTX (320m/z; Mebs et al., 1995). However, the presence of other chemicals in the extract/matrix can reduce sensitivity (Bane et al., 2014), thus chromatography can be used prior to ESI-MS for chemical separation (Yotsu-Yamash*ta and Tateki, 2010). Mebs et al. (2018a) is the only study of Atelopus toxins to employ high resolution hydrophilic interaction liquid chromatography (HR-HILIC), a highly sensitive method which achieves better separation of analogs than reverse phase chromatography (Knutsen et al., 2017). Subsequent analysis by ESI-MS detected only TTX and 4-epiTTX in an A. hoogmoedi extract (Mebs et al., 2018a). As HR-HILIC-LC/MS can detect nearly every major analog of TTX found in nature (Yotsu-Yamash*ta et al., 2013), the exclusive detection of TTX and 4-epiTTX suggests that A. hoogmoedi lacks other TTX analogs including 6-epitetrodotoxin and 11-deoxytetrodotoxin, which are frequently detected in newts (Hanifin, 2010). This is consistent with the apparent absence of 6-epitetrodotoxin and tetrodonic acid in A. carbonerensis (Mebs et al., 1995; Yotsu-Yamash*ta et al., 1992). Application of HR-HILIC-LC/MS to other Atelopus species could determine whether 6-epitetrodotoxin, 11-deoxytetrodotoxin, and tetrodonic acid are universally absent in the genus.

HR-HILIC-LC/MS forms the basis of a mass spectrometry-guided screening method developed by Kudo et al. (2014, 2020): it enables the detection of unknown compounds for subsequent structural analysis. The pattern of unique TTX analogs and other chemicals detected in newts via this method has led to the determination of a putative TTX biosynthetic pathway (Kudo et al., 2014, Kudo et al., 2016, Kudo et al., 2020, Kudo et al., 2021, Kudo and Yotsu-Yamash*ta, 2019). Use of this screening method on Atelopus could help determine the extent of convergence in the biosynthetic pathway(s) of amphibian associated TTX. Furthermore, the search for genes in amphibians and bacteria which encode for inferred biosynthetic enzymes could give powerful insight into the ultimate source of TTX (Kudo et al., 2014).

The identification of individual Atelopus bufadienolides has only been attempted once (Flier et al., 1980). After verification of bufadienolide presence, Flier et al. (1980) performed HPLC on A. “ignescens” skin extractions. By comparing the elution order with those of bufadienolide standards, the preliminary identities of six Atelopus bufadienolides were determined. A variety of methods are now available which make it possible to identify many bufadienolides precisely and sensitively (Zhan et al., 2020).

5.3. Quantification of toxicity and pharmacological activity

Most of the methods described in Section 5.2 can quantify individual chemicals, subject to their detection and identification limitations (Table 1). In contrast, the methods described below can be used to determine the combined toxicity or pharmacological activity of multiple toxins.

The mouse bioassay (MBA) is the most frequently used method of toxicity quantification in Atelopus toxin studies. MBAs may also be used throughout the toxin extraction and purification process to identify toxic fractions (Shindelman et al., 1969; Yotsu-Yamash*ta and Tateki, 2010; Yotsu et al., 1990b). MBAs involve injecting varying doses of toxin intraperitoneally into mice and measuring survival times (Brown et al., 1977). The symptoms observed during MBAs may suggest the presence or absence of different toxins (Fuhrman et al., 1969). Toxin quantities determined by MBAs are reported in mouse units (MUs). The standard definition of one MU is that it is enough toxin to kill a 20g male mouse in 30min by intraperitoneal injection and is equivalent to 0.22 μg of tetrodotoxin (Kawabata, 1978; Yasumoto, 1991). However, Atelopus toxin studies published prior to 1988 use conflicting definitions of MUs based on post-injection survival times of 20min (Brown et al., 1977; Pavelka et al., 1977), 30min (Mebs and Schmidt, 1989) and 1h (Kim et al., 1975; Shindelman et al., 1969). Other studies do not specify the survival time used in their MU definition (Fuhrman et al., 1969, 1976). Lastly, female mice and mice of different strains were sometimes used for MBAs (Pavelka et al., 1977). Post 1989, all Atelopus studies that use MBAs for toxin quantification apply the standard MU definition or use the standard MU to TTX equivalent conversion outlined in Yasumoto (1991). The variability in MU definition between Atelopus studies may complicate the comparability of the toxicities they report.

Binding assays can be used to determine the quantity of chemicals with specific pharmacological activities. While other variants of toxin binding assays exist (Stokes et al., 2012), those applied in studies of Atelopus toxins thus far measure the binding inhibition of radioactively labeled reference chemicals in hom*ogenized brain tissue or red blood cells by toad skin extracts. Binding inhibition assays for guanidinium alkaloids and bufadienolides use [³H]Saxitoxin and [³H]Ouabain as reference chemicals, respectively (Daly et al., 1994, 1997; Flier et al., 1980). Modern binding assays can be an order of magnitude more sensitive than the mouse bioassay in detecting guanidinium alkaloids (Kawabata, 1978; Stokes et al., 2012), and have fewer ethical considerations (Stern et al., 2018; Taylor et al., 2011; Wilder-Kofie et al., 2011).

6.1. Guanidinium alkaloids

Sequestered toxin profiles are constrained by the availability of toxins (an environmental factor; Mebs et al., 2018b; Yoshida et al., 2020) and the capacity of the organism to sequester those chemicals (a genetic factor; Davison et al., 2021). Atelopus may sequester guanidinium alkaloids from bacteria, and, if so, it is unclear whether such bacteria are horizontally or vertically transmitted. In the case of horizontal transmission, Atelopus toxin profiles would be constrained by the geographic distributions of guanidinium-alkaloid synthesizing bacteria, whereas vertical transmission would ensure the availability of particular guanidinium alkaloids, regardless of any biogeographic patterns in microbial diversity. In both scenarios, Atelopus guanidinium alkaloid profiles would also be shaped by the genetic capacity to sequester specific toxins, establish symbioses with toxin-producing bacteria, or, in the possible case of CHTX, to modify sequestered chemicals (see Section 4.1.3). If, instead, Atelopus guanidinium alkaloids are endogenously synthesized, toxin profiles should reflect genetically controlled synthetic abilities and, possibly, plasticity in defensive responses to environmental cues (e.g., predation attempts, reproductive status, etc). Thus it is likely that both genetic and environmental factors shape Atelopus guanidinium toxin profiles.

TTX. Tetrodotoxin is the most widespread Atelopus toxin, having been detected in ten of the sixteen chemically assessed species (Fig. 3a). It is usually a major toxin component and has been found in Atelopus toads from across the entire geographic range of the genus (Fig. 3b) and within both major clades (Fig. 3a). Atelopus is the only taxon in Bufonidae known to possess guanidinium alkaloids (Rodríguez et al., 2017); toxin assessments of species within the sister taxon of Atelopus (Oreophrynella;Kok et al., 2018) and other “atelopodid” bufonids (Melanophryniscus and Dendrophryniscus; Graybeal, 1997) have failed to detect guanidinium alkaloids (Daly et al., 1994; Mebs et al., 1995). Thus, the phylogenetic distribution of TTX within Bufonidae suggests a single origin of TTX defense in the common ancestor of Atelopus with possible secondary losses in some species, i.e., A. glyphus, A. limosus, and A. cruciger (Fig. 3a). If TTX is sequestered from bacteria in Atelopus, the absence of TTX could be reflective of differences in microbiome composition rather than the loss of sequestration ability. However, better sampling of the Amazonian-Guianan Atelopus clade and of Bufonidae more generally is needed before a definitive model of the origin and evolution of TTX defense in Atelopus can be proposed.

4,9-anhydroTTX and 4-epiTTX frequently cooccur with TTX, which is expected given the aqueous equilibrium between the three chemicals. Interestingly, 4-epiTTX has once been detected in the absence of 4,9-anhydroTTX in a single male specimen of A. hoogmoedi from Suriname (Fig. 3b; Mebs et al., 2018a).

CHTX. Chiriquitoxin was discovered in the now-extinct Atelopus chiriquiensis in 1975 and was thought to be unique to that species until its detection in A. limosus and A. glyphus more than three decades later (Kim et al., 1975; Yotsu-Yamash*ta and Tateki, 2010). CHTX is a major component in all three species (Table S1), and appears to be restricted to Central America, having only been found in Costa Rican and Panamanian Atelopus (Fig. 3b). The three species with CHTX form a polyphyletic group. A. certus and A. senex possess guanidinium alkaloid-like toxins (Brown, 1972; Fuhrman et al., 1969), and their phylogenetic placement (Fig. 3a) and Central American ranges (Kahn et al., 2005; Veselý and Batista, 2021) suggest they would be promising targets for CHTX testing. However, A. senex is extinct (IUCN, 2021). TTX was also a major component in A. chiriquiensis, consistently detected alongside CHTX (Table S1). Although TTX is the likely metabolic precursor of CHTX (Yotsu et al., 1990b), TTX is not known to be present in A. glyphus or A. limosus (Yotsu-Yamash*ta and Tateki, 2010), indicating that TTX, if present in these species, could be completely converted to CHTX.

ZTX AB. Zetekitoxin AB (ZTX AB) has only been found in two of the seven assessed Central American Atelopus: A. varius and A. zeteki (Fig. 3a). These sister species are closely related (Fig. 3a; Lötters et al., 2011; Ramírez et al., 2020) and a recent whole-genome analysis does not support the species boundary between them (Byrne et al., 2020), suggesting that A. varius and A. zeteki are the same species. A. varius and A. zeteki exhibit intraspecific variation in the presence and absence of TTX and ZTX AB. The exclusive presence of ZTX AB in A. varius and A. zeteki suggests that ZTX AB sequestration or synthesis is under some degree of genetic control. This is corroborated by the occurrence of Colostethus panamensis, a dendrobatid poison frog, in El Valle de Antón. C. panamensis occupies the same habitat as A. zeteki but possesses only TTX, not ZTX AB (Daly et al., 1994). Nevertheless, the paucity of data makes it difficult to draw any conclusion on the broader phylogenetic or geographic distribution of this toxin.

6.2. Bufadienolides

Bufadienolides have been detected in all Atelopus which have been tested with methods that are sensitive to these substances: A. “ignescens,” A. spurrelli, A. varius, and A. zeteki (Fig. 3a; Daly et al., 1997; Flier et al., 1980). Thus, while cardiac glycosides appear geographically restricted to Andean and Central American harlequin toads (Fig. 3b), this is likely an artifact of incomplete sampling. Identification of Atelopus bufadienolides has been attempted only once, revealing the likely presence of major components telocinobufa*gin and bufotalin, and minor components including marinobufa*gin, cinobufa*gin, bufalin, and arenobufa*gin, as well as two unidentified bufadienolides (Fig. 2; Flier et al., 1980). In other bufonids, bufadienolide profiles are highly variable between populations, species, and life history stages. Factors implicated in this variation include population structuring, environmental factors like climate and habitat quality, and microbial toxin biotransformation (Bókony et al., 2016, 2019; Cao et al., 2019; Hayes et al., 2009a, 2009b; Inoue et al., 2020; Kamalakkannan et al., 2017). Consequently, there is likely undiscovered bufadienolide diversity and variation within Atelopus.

7.1. Atelopus toxin profiles

While many individual Atelopus toxins have been detected and characterized both chemically and pharmacologically, the adaptive importance of the specific compositions of Atelopus toxin co*cktails remains uninvestigated. From an antipredator perspective, for instance, it is unclear whether possessing ZTX AB or CHTX rather than TTX as a major toxin component would affect harlequin toad fitness. It is possible that the unique binding patterns of different guanidinium alkaloids (see Section 4.1) result in functional differences relevant to warding off specific predators. Alternatively, considering the similar toxicities of TTX, CHTX, and ZTX to mice, the identity of the major alkaloid component in each Atelopus species may be of no adaptive significance. Future studies that involve exposing potential Atelopus predators to different guanidinium alkaloids could determine whether major toxin component identity influences the effectiveness of Atelopus chemical defenses.

The implications of simultaneously maintaining guanidinium alkaloids and cardiac glycosides, two chemically and pharmacologically distinct toxin classes, are worth consideration. Having diverse toxin types can enable organisms to be defended against multiple natural enemies, as demonstrated in chemically defended plants (Lindroth and Hwang, 1996). Furthermore, toxins can synergize to magnify each other's effects (Nelson and Kursar, 1999; Raaymakers et al., 2017). The respective targets of bufadienolides and guanidinium alkaloids, VGSC and Na⁺/K⁺-ATPase proteins, both influence sodium ion concentrations and may thus interact physiologically. In astrocytes, a type of glial cell, inhibition of VGSCs results in lower Na⁺/K⁺-ATPase activity. VGSCs may maintain Na⁺ ion concentrations at levels necessary for Na⁺/K⁺-ATPase function (Sontheimer et al., 1994). The interaction between these membrane proteins could have consequences for the function of Atelopus toxin co*cktails. Interestingly, tetrodotoxin reduces the toxicity of cardiac glycosides (CGs) when injected directly into the brain of cats but potentiates CG toxicity when given intravenously (Peres-Gomes and Ribeiro, 1979). The difference in effect between TTX administered to the brain and TTX administered intravenously is probably a consequence of the impermeability of the blood-brain barrier to TTX (Zimmer, 2010). Tissue-specific expression of uncharacterized receptors for toxin-binding proteins could also result in delivery-dependent differences in poisoning symptoms. In dogs, intravenous TTX increases survival times and reduces cardiac arrhythmia following cardiac glycoside poisoning (Bernstein, 1969). The prevalence of system- and delivery-dependent results highlight the need to investigate the interactions of bufadienolide and guanidinium alkaloids in predator systems that are biologically relevant to Atelopus.

If Atelopus sequester guanidinium alkaloids, bufadienolide biosynthesis could provide some level of chemical defense when said sequestration is disrupted. Captive-raised Atelopus that lack guanidinium alkaloids retain bufadienolides (Daly et al., 1997). Melanophryniscus is another genus of bufonid toads where autogenous toxin production may compensate for variability in toxin sequestration (Cei et al., 1968; Hantak et al., 2013). Although Melanophryniscus that are fed an alkaloid-free diet gradually lose lipophilic alkaloids from their skins (Mebs et al., 2018b) Melanophryniscus may be able to upregulate the synthesis of indole alkaloids when lipophilic alkaloids are low (Jeckel et al., 2015). Similarly, the myobatrachid, Pseudophryne semimarmorata, synthesizes more indole alkaloids when raised in captivity without access to dietary lipophilic alkaloids (Smith et al., 2002). In contrast to Melanophryniscus and Pseudophryne, bufadienolide quantities are similar in A. varius with and without guanidinium alkaloids, suggesting that bufadienolide production is not upregulated in response to low TTX levels (Daly et al., 1997). More investigation is needed to clarify the functional role and regulation of autogenous toxins in Atelopus.

7.2. Variation in toxicity between and within species

A common characteristic of chemically defended organisms is variation in toxicity, from the individual to the species level and in response to temporal, environmental, and physiological changes (Speed et al., 2012). Atelopus is no different. Individual harlequin toads range from completely nontoxic to toxic enough to kill thousands of mice (Fig. 4 inset). The causes of this variation are unknown and presumably depend on the source of a toxin and the ability of the toad to bioaccumulate the toxin or host its producers; however, some patterns do emerge which parallel those observed in better-studied systems. It is important to note that the toxicity values reported for Atelopus are primarily reflective of the guanidinium alkaloids present in their skin because the acidic aqueous extraction methods commonly used prior to toxin quantification likely exclude some or all bufadienolides from the resulting toxic fractions (see Section 5.1). When methods sensitive to bufadienolides were used, bufadienolide quantities in Atelopus were found to be large enough to contribute to the overall toxicity of the toads (Daly et al., 1997; Flier et al., 1980). Studies published prior to 1989 employed alternative and conflicting mouse unit definitions, which impede the comparability of reported Atelopus toxicities (see Section 5.3 for an expanded discussion).

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Fig. 4

Geographic (N=892 individuals) and species-level (inset) variation in adult Atelopus toxicity. Toxicity values primarily reflect quantity of guanidinium alkaloids (see section 5.1). Numbers left of species names detail the total number of toxicity assessments and number of specimens assessed (in parentheses) for each species. One mouse unit (MU) is sufficient to kill a single average-weight mouse in 30min upon injection (Yasumoto, 1991). When toxicity values were given in TTX equivalents or when TTX quantity alone was given, conversion to MUs used the conversion factor 1 MU=0.22 μg TTX (Yasumoto, 1991). See Supplementary Table S2 for coordinates, toxicity values, species names, sources, and details on unreported sample size data. Toxicity data summarized and graphed using ggplot2 v3.3.3 (Wickham, 2016), dplyr v1.0.6 (Wickham et al., 2021), and cowplot v1.1.1 (Wilke, 2020).

The most toxic harlequin toads (presently known) are found in Central America (e.g. up to 1200 MU/toad for A. zeteki and 948 MU/toad for A. chiriquiensis;Kim et al., 1975; Pavelka et al., 1977) and the montane cloud forests of the state of Mérida, Venezuela (A. carbonerensis, up to 1000 MU/toad; Dole and Durant, 1974; Yotsu-Yamash*ta et al., 1992). Atelopus from the Guiana Shield and the Andes south of Venezuela generally have low toxin levels (Fig. 4). However, while a minimum of 889 Central American harlequin toads have been assessed for toxicity, at least 39 from all other geographic regions have been tested (Fig. 4 inset, see Table S2 for details on reported sample sizes). The lack of standardization in older Atelopus toxicity studies also prevents confirmation of these patterns. Future research on Atelopus toxicity could prioritize the sampling of toads from the Andes and Guiana Shield.

Atelopus toxicity may be influenced by selection pressure from predators. In Taricha, a genus of newts that possesses as many as 60,000 MU of TTX per individual, high toxin levels are thought to be driven by a coevolutionary relationship with TTX-resistant predatory garter snakes (Thamnophis) (Hague et al., 2020, Hanifin et al., 2002, Williams et al., 2003). The intensity of reciprocal selection between these species varies geographically, resulting in populations with drastically different toxicities and toxin resistances (Hague et al., 2020). A similar situation is possible between Atelopus and one or more predator species (such as Erythrolamprus epinephalus). Future studies could investigate covariance in Atelopus predator toxin resistance and Atelopus toxicity across the sympatric ranges of both taxa to see if a coevolutionary arms race is taking place.

Some of the observed intrapopulation variation in Atelopus toxicity (Daly et al., 1994; Kim et al., 1975; Mebs et al., 1995; Pavelka et al., 1977) might be attributable to the experiences of sampled individuals. While also found in the skin epithelium and the liver, Atelopus toxins are primarily localized in the granular glands, which are distributed across the body and can eject their contents when a toad feels threatened (Mebs et al., 2018a; Toledo and Jared, 1995). A toad that was recently attacked may have temporarily diminished its skin-associated stores of alkaloids and steroids. Over longer time periods, encounters with predators could lead to higher toxicity in Atelopus individuals. Predator cue exposure and simulated predator attacks induce increased toxicity in some amphibian species that possess guanidinium alkaloids or bufadienolides (Benard and Fordyce, 2003; Bucciarelli et al., 2017), but not in others (Brossman et al., 2014; Üveges et al., 2017). The plasticity of Atelopus chemical defenses needs investigation and could provide insight into the ecological significance of their toxins.

Reproductive cycles and development may also play a role in toxicity variation. Gravid Atelopus chiriquiensis females have lower skin-associated toxin levels than males but are comparably poisonous when the toxicities of their eggs are accounted for (Pavelka et al., 1977). Thus, Atelopus may provision their eggs with toxins, possibly as a defensive measure, and this process likely involves the diversion of skin toxins into the reproductive system (Pavelka et al., 1977; Yotsu-Yamash*ta and Tateki, 2010). More generally, toxicity may vary with Atelopus age and/or metamorphic stage, as seen in other bufadienolide-defended toads (Hayes et al., 2009a; Üveges et al., 2017) and tetrodotoxin-defended newts (Gall et al., 2011; Tsuruda et al., 2002) and octopi (Williams, 2008; Williams et al., 2011). In other toxin-sequestering frog species, body size has been positively correlated with granular gland capacity (Saporito et al., 2010) and overall toxin quantity (Jeckel et al., 2015). Atelopus toxin provisioning and ontogenetic changes in toxicity could be investigated for mechanisms involved in toxin transport and accumulation.

Most speculatively, unidentified environmental factors may influence the success of microbe-toad symbioses, which are possible sources of guanidinium alkaloids in Atelopus. Some Atelopus species have extremely high site fidelity and individuals may thus be exposed to relatively constant microenvironments throughout their lives (Crump, 1986; Tarvin et al., 2014). Atelopus occupy a variety of habitats, and the possibility of covariance between toxicity and abiotic factors such as temperature and precipitation remains an area of interest.

^{Appendix A}Supplementary data to this article can be found online at https://doi.org/10.1016/j.toxcx.2022.100092.

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