Sexual Dichromatism Drives Diversification Within a Major

Copyedited by: AV MANUSCRIPT CATEGORY: Systematic Biology

Syst. Biol. 68(6):859–875, 2019 © The Author(s) 2019. Published by Oxford University Press, on behalf of the Society of Systematic Biologists. All rights reserved. For permissions, please email: [email protected] DOI:10.1093/sysbio/syz023 Advance Access publication April 23, 2019 Sexual Dichromatism Drives Diversiﬁcation within a Major Radiation of African Amphibians

, ∗ , , , DANIEL M. PORTIK1 2 ,RAYNA C. BELL1 3,DAV I D C. BLACKBURN4,AARON M. BAUER5,CHRISTOPHER D. BARRATT6 7 8, , , , , WILLIAM R. BRANCH9 10†,MARIUS BURGER11 12,ALAN CHANNING13,TIMOTHY J. COLSTON14 15,WERNER CONRADIE9 16 , , , J. MAXIMILIAN DEHLING17 ,ROBERT C. DREWES18,RAFFAEL ERNST19 20,ELI GREENBAUM21,VÁCLAV GVOŽD´IK22 23,JAMES , HARVEY24,ANNIKA HILLERS25 26,MAREIKE HIRSCHFELD25,GREGORY F. M. JONGSMA4,JOS KIELGAST27,MARCEL T. Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 , KOUETE4,LUCINDA P. L AWSON28 29,ADAM D. LEACHÉ30,SIMON P. L OADER31,STEFAN LÖTTERS32,ARIE VAN DER MEIJDEN33,MICHELE MENEGON34,SUSANNE MÜLLER32,ZOLTÁN T. N AGY35,CALEB OFORI-BOATENG36,ANNEMARIE OHLER37,THEODORE J. PAPENFUSS1,DANIELA RÖßLER32,ULRICH SINSCH17 ,MARK-OLIVER RÖDEL25,MICHAEL VEITH32, JENS VINDUM18,ANGE-GHISLAIN ZASSI-BOULOU38, AND JIMMY A. MCGUIRE1 1Museum of Vertebrate Zoology, University of California, Berkeley, CA 94720, USA; 2Department of Ecology and Evolutionary Biology, University of Arizona, Tucson, AZ 85721, USA; 3Department of Vertebrate Zoology, National Museum of Natural History, Smithsonian Institution, Washington, DC 20560-0162, USA; 4Florida Museum of Natural History, University of Florida, Gainesville, FL 32611, USA; 5Department of Biology, Villanova University, 800 Lancaster Avenue, Villanova, PA 19085, USA; 6Department of Environmental Sciences, University of Basel, Basel 4056, Switzerland; 7German Centre for Integrative Biodiversity Research (iDiv) Halle-Jena-Leipzig, Leipzig 0413, Germany; 8Max Planck Institute for Evolutionary Anthropology, Leipzig 0413, Germany; 9Port Elizabeth Museum, P.O. Box 11347, Humewood 6013, South Africa; 10 Department of Zoology, Nelson Mandela Metropolitan University, P.O. Box 77000, Port Elizabeth 6031, South Africa; 11 African Amphibian Conservation Research Group, Unit for Environmental Sciences and Management, North-West University, Potchefstroom 2520, South Africa; 12Flora Fauna & Man, Ecological Services Ltd. Tortola, British Virgin, Island; 13Unit for Environmental Sciences and Management, North-West University, Potchefstroom 2520, South Africa; 14 Department of Biological Sciences, Florida State University, Tallahassee, FL 32306, USA; 15Zoological Natural History Museum, Addis Ababa University, Arat Kilo, Addis Ababa, Ethiopia; 16 School of Natural Resource Management, Nelson Mandela University, George Campus, George 6530, South Africa; 17 Department of Biology, Institute of Sciences, University of Koblenz-Landau, Universitätsstr. 1, D-56070 Koblenz, Germany; 18 California Academy of Sciences, San Francisco, CA 94118, USA; 19 Museum of Zoology, Senckenberg Natural History Collections Dresden, Königsbrücker Landstr. 159, Dresden 01109, Germany; 20Department of Ecology, Technische Universität Berlin, Rothenburgstr. 12, Berlin 12165, Germany; 21Department of Biological Sciences, University of Texas at El Paso, El Paso, TX 79968, USA; 22The Czech Academy of Sciences, Institute of Vertebrate Biology, Brno, Czech Republic; 23Department of Zoology, National Museum, Prague, Czech Republic; 24Pietermaritzburg, KwaZulu-Natal, South Africa; 25Museum für Naturkunde, Leibniz Institute for Evolution and Biodiversity Science, Biodiversity Dynamics, Invalidenstr. 43, Berlin 10115, Germany; 26AcrosstheRiver–ATransboundary Peace Park for Sierra Leone and Liberia, The Royal Society for the Protection of Birds, 164 Dama Road, Kenema, Sierra Leone; 27 Natural History Museum of Denmark, University of Copenhagen, Universitetsparken 15, Copenhagen 2100, Denmark; 28Department of Biological Sciences, University of Cincinnati, 614 Rieveschl Hall, Cincinnati, OH 45220, USA; 29Life Sciences, Field Museum of Natural History, 1400 S. Lake Shore Dr., Chicago, IL 60605, USA; 30Department of Biology, Burke Museum of Natural History and Culture, University of Washington, Seattle, WA, USA; 31 Life Sciences Department, Natural History Museum, London SW7 5BD, UK; 32Biogeography Department, Trier University, Universitätsring 15, Trier 54296, Germany; 33CIBIO Research Centre in Biodiversity and Genetic Resources, InBIO, Universidade do Porto, Campus Agrario de Vairão, Rua Padre Armando Quintas, No. 7, 4485-661 Vairão, Vila do Conde, Portugal; 34Tropical Biodiversity Section, Science Museum of Trento, Corso del lavoro e della Scienza 3, Trento 38122, Italy; 35Royal Belgian Institute of Natural Sciences, OD Taxonomy and Phylogeny, Rue Vautier 29, B-1000 Brussels, Belgium; 36Forestry Research Institute of Ghana, P.O. Box 63, Fumesua, Kumasi, Ghana; 37 Département Origines et Evolution, Muséum National d’Histoire Naturelle, UMR 7205 ISYEB, 25 rue Cuvier, Paris 75005, France; and 38Institut National de Recherche en Sciences Exactes et Naturelles, Brazzaville BP 2400, République du Congo †Deceased. ∗ Correspondence to be sent to: Department of Ecology and Evolutionary Biology, University of Arizona, Tucson, AZ 85721, USA; E-mail: [email protected]

Received 27 July 2018; reviews returned 15 February 2019, 18 March 2019; accepted 9 April 2019 Associate Editor: Michael Alfaro

Abstract.—Theory predicts that sexually dimorphic traits under strong sexual selection, particularly those involved with intersexual signaling, can accelerate speciation and produce bursts of diversification. Sexual dichromatism (sexual dimorphism in color) is widely used as a proxy for sexual selection and is associated with rapid diversification in several animal groups, yet studies using phylogenetic comparative methods to explicitly test for an association between sexual dichromatism and diversification have produced conflicting results. Sexual dichromatism is rare in frogs, but it is both striking and prevalent in African reed frogs, a major component of the diverse frog radiation termed Afrobatrachia. In contrast to most other vertebrates, reed frogs display female-biased dichromatism in which females undergo color transformation, often resulting in more ornate coloration in females than in males. We produce a robust phylogeny of Afrobatrachia to investigate the evolutionary origins of sexual dichromatism in this radiation and examine whether the presence of dichromatism is associated with increased rates of net diversification. We find that sexual dichromatism evolved once within hyperoliids and was followed by numerous independent reversals to monochromatism. We detect significant diversification rate heterogeneity in Afrobatrachia and find that sexually dichromatic lineages have double the average net diversification rate of monochromatic lineages. By conducting trait simulations on our empirical phylogeny, we demonstrate that our inference of trait-dependent diversification is robust. Although sexual dichromatism in hyperoliid frogs is linked to their rapid diversification and supports macroevolutionary predictions of speciation by sexual selection, the function of dichromatism in reed frogs remains unclear. We propose that reed frogs are a compelling system for studying the roles of natural and sexual selection on the evolution of sexual dichromatism across micro- and macroevolutionary timescales. [Afrobatrachia; Anura; color evolution; diversification; macroevolution; sexual selection.] 859

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In The Descent of Man and Selection in Relation to Sex, Eclectus results from intrasexual competition to attract Darwin (1871) observed that many closely related taxa mates and intersexual differences in exposure to visual differed primarily in secondary sexual characters and predators (Heinsohn et al. 2005). Likewise, the dynamic suggested that sexual selection plays a role in the sexual dichromatism in frogs that form large breeding diversiﬁcation of species. The concept of speciation aggregations may serve to identify other competing through sexual selection was later developed into males rather than to attract mates (Sztatecsny et al. a theory that links the coevolution of secondary 2012; Kindermann and Hero 2016; Bell et al. 2017b). sexual traits and mating preferences to premating Consequently, investigating the evolution of secondary

reproductive isolation (Lande 1981, 1982; Kirkpatrick sexual characters like dichromatism across ecologically Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 1982; West-Eberhard 1983). This conceptual framework diverse taxonomic groups is essential if we aim to predicts that both the strength of sexual selection generalize about the function of sexually dimorphic and the prevalence of sexually selected traits have traits and better understand the roles of natural and a positive association with speciation rate (Lande sexual selection in biological diversification. 1981, 1982; West-Eberhard 1983; Barraclough et al. Secondary sexual traits are diverse and prevalent 1995). If divergence in secondary sexual characters and among anurans (frogs and toads) and include sexual size sexual selection are indeed major drivers of speciation, dimorphism, structures for acoustic signaling, nuptial then clades exhibiting elaborate sexually dimorphic pads, spines, elongated fingers, glands, and sexual traits and/or elevated sexual selection should display dichromatism (Duellman and Trueb 1986). Sexual size higher species richness and increased diversification dimorphism is common and occurs in >90% of species rates at macroevolutionary scales. Empirically, these (Shine 1979; Han and Fu 2013), and body size evolution of predictions have mixed support across a range of the sexes is often attributed to fecundity for females and sexually dimorphic traits that serve as proxies for intrasexual competition, energetic constraints, agility, sexual selection (reviewed in Kraaijeveld et al. 2011), and predation for males (Salthe and Duellman 1973; indicating that macroevolutionary trends for traits Wells 1977; Shine 1979; Woolbright 1983; De Lisle and involved with intersexual signaling and mate-choice Rowe 2013; Han and Fu 2013; Nali et al. 2014). Male may differ from those more strongly influenced by frogs of most species produce acoustic or vibrational intrasexual or natural selection. For example, traits signals to attract females, and these signals are essential under ecological selection such as body size dimorphism for mate recognition and relaying social information have no consistent relationship with diversification (Ryan 1980; Gerhardt 1994; Gerhardt and Huber 2002). (Kraaijeveld et al. 2011), whereas sexual dichromatism, Structures like spines and tusks, which are present in a form of sexual dimorphism in which the sexes differ males of many species, are used in male–male combat in color, often functions as a mate recognition signal and (e.g. McDiarmid 1975) whereas the diverse assortment is associated with diversification in several taxonomic of glands and nuptial pads, which are widespread groups (Misof 2002; Stuart-Fox and Owens 2003; Alfaro in male frogs, are likely involved in courtship and et al. 2009; Kazancıoglu et al. 2009; Wagner et al. 2012). amplexus (Duellman and Trueb 1986). In contrast to Accordingly, sexual dichromatism has become a widely these widespread secondary sexual characters, anuran used proxy for studying the effects of sexual selection sexual dichromatism is rare, occurring in only ∼2% of on speciation rate. However, phylogenetic comparative frog species (Bell and Zamudio 2012; Bell et al. 2017b). analyses explicitly testing for an association between Behavioral studies in a handful of frog species indicate sexual dichromatism and diversification have produced that these sexual color differences may be subject conflicting results, even in well-studied groups such to natural selection as well as inter- and intrasexual as birds (Barraclough et al. 1995; Owens et al. 1999; selection (Maan and Cummings 2009; Sztatecsny et al. Morrow et al. 2003; Phillimore et al. 2006; Seddon 2012). Sexual selection on coloration has historically et al. 2013; Huang and Rabosky 2014) where sexual been dismissed in frogs with the assumption that dichromatism plays an important role in signaling and communication is predominately acoustic (reviewed mate-choice (Price 2008). Differences in methodologies in Starnberger et al. 2014); however, several studies and the spatial, temporal, and taxonomic scales document the importance of color signals in courtship among studies may partially explain these contrasting behavior and mate-choice, even in nocturnal species results. In particular, recent studies have highlighted (Gomez et al. 2009, 2010; Jacobs et al. 2016; Yovanovich concerns about the ability to distinguish between et al. 2017; Akopyan et al. 2018). If sexual dichromatism trait-dependent and trait-independent diversification in frogs contributes to premating reproductive isolation, scenarios using phylogenetic comparative methods then sexually dichromatic clades may fit the conceptual (Rabosky and Goldberg 2015; Beaulieu and O’Meara framework of speciation by sexual selection and display 2016). In a broader sense, however, this disparity among higher species richness and increased diversification studies may reflect more nuanced or novel mechanisms rates at macroevolutionary scales. underlying the evolution of sexual dichromatism such Afrobatrachia is a frog radiation (Arthroleptidae, that it does not consistently fit the conceptual framework Brevicipitidae, Hemisotidae, Hyperoliidae) that of speciation by sexual selection. For instance, the includes over 400 species distributed across sub- striking sexual dichromatism in parrots of the genus Saharan Africa that reflects much of the morphological,

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Juvenile (Phase J)

♂ ♀ Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 Male Phase F Male Phase J Female Phase F

FIGURE 1. Illustration of ontogenetic color change occurring in hyperoliid frogs (Hyperolius dintelmanni shown) that underlies sexual dichromatism. In dichromatic species, females undergo a color change from Phase J to Phase F in response to steroid hormones at the onset of sexual maturity. Males retain the juvenile coloration (Phase J) or undergo a parallel change in color (to Phase F), however the proportion of the male color phases in populations varies across species. Secondary monochromatism can evolve from dichromatism through the loss of Phase J males (both sexes undergo an ontogenetic color change to Phase F) or through the loss of ontogenetic color change in both sexes (both sexes retain Phase J coloration at sexual maturity).

ecological, and reproductive mode diversity present in distinct female color patterns in reed frogs may play anurans (Portik and Blackburn 2016). Afrobatrachian a role in courtship and mate recognition at breeding frogs display many unusual secondary sexual characters sites where upwards of 8 hyperoliid species congregate including the hair-like skin structures of male Hairy in a single night (Drewes and Vindum 1994; Kouamé Frogs (Trichobatrachus), an elongate third digit in et al. 2013; Portik et al. 2018). The link between sexual males (up to 40% of body length in Arthroleptis dichromatism and rapid speciation across disparate and Cardioglossa), extreme body size dimorphism animal clades (Misof 2002; Stuart-Fox and Owens 2003; (Leptopelis, Chrysobatrachus, Breviceps), and prominent Alfaro et al. 2009; Kazancıoglu et al. 2009; Wagner et al. pectoral glands (Leptopelis) or gular glands on the male 2012) demonstrates that when sexual dichromatism vocal sac (Hyperoliidae). Afrobatrachian frogs also functions primarily as an intersexual signal under have the highest incidence of sexual dichromatism strong sexual selection, there are predictable outcomes among anurans, which is striking and prevalent in on diversification rate. Therefore, as a first step toward many hyperoliid reed frogs (Hyperolius, Heterixalus). understanding the potential function of dichromatism In sexually dichromatic hyperoliids, the difference in hyperoliids, including the plausibility of intersexual in coloration is female-biased: both sexes exhibit a signaling, we aim to assess whether this trait fits the consistent juvenile coloration upon metamorphosis predictions of speciation by sexual selection on a (termed Phase J), but at the onset of maturity sex macroevolutionary scale. steroids trigger a color and/or color pattern change in In this study, we reconstruct the evolutionary females (Phase F), whereas males typically retain the history of sexual dichromatism within Afrobatrachia juvenile color pattern (Fig. 1)(Schiøtz 1967; Richards and investigate whether diversification rate shifts 1982; Hayes 1997; Hayes and Menendez 1999). In many in this continental radiation are associated with dichromatic reed frog species adult males can also dichromatism. We produce a well-resolved species display the Phase F coloration, but this generally occurs tree of Hyperoliidae from genomic data (>1000 loci) in lower frequency than the Phase J morph (Fig. 1) and greatly increased taxonomic sampling relative to (Schiøtz 1967, 1999; Amiet 2012; Kouamé et al. 2015; previous studies (Wieczorek et al. 2000; Veith et al. Portik et al. 2016a). The function of the ontogenetic color 2009; Portik and Blackburn 2016). To explore the shift in female reed frogs is poorly understood (Bell and evolution of sexual dichromatism within the broader Zamudio 2012), but it may be similar to female-biased phylogenetic and biogeographic context of African sexual dichromatism in other vertebrates, which can frogs, we incorporate all published sequence data result from a reversal in mating system in which females of Afrobatrachia to produce a robust, time-calibrated compete for males (Andersson 1994) or sexual niche phylogeny. Using methods that improve the accuracy partitioning in which males and females use different of character state reconstructions by accounting for trait habitats (Shine 1989; Heinsohn et al. 2005). Alternatively, and diversification rate heterogeneity (King and Lee

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2015; Beaulieu and O’Meara 2016), we test the hypothesis KIAA2013) were also incorporated based on published that sexual dichromatism evolved repeatedly within sequence data (Portik and Blackburn 2016). The final Afrobatrachia (Veith et al. 2009). Finally, we estimate net marker set for probe design included 1265 genes from diversification rates from our time-calibrated phylogeny 4 species and 5060 individual sequences, with a total using the hidden state speciation and extinction (HiSSE) of 995,700 bp of target sequence. These sequences framework (Beaulieu and O’Meara 2016). Specifically, were used to design a MYbaits-3 custom bait library we examine if diversification rate shifts occur within (MYcroarray, now Arbor Biosciences) consisting of 120 Afrobatrachia and if so, whether they are associated with mer baits and a 2× tiling scheme (every 60 bp), which

the occurrence of sexual dichromatism. Given recent resulted in 60,179 unique baits. Transcriptomes, target Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 concerns raised about the ability to distinguish between sequences, and probe designs are available on Dryad trait-dependent and trait-independent diversification (Portik et al. 2016c). scenarios (Rabosky and Goldberg 2015; Beaulieu Genomic DNA was extracted using a high-salt and O’Meara 2016), we assess the performance of extraction method (Aljanabi and Martinez 1997) and available state-dependent diversification methods with individual genomic libraries were prepared following a trait simulation study conducted using our empirical Meyer and Kircher (2010) with modifications described phylogeny. in Portik et al. (2016b). Samples were pooled for capture reactions based on phylogenetic relatedness, and the combined postcapture libraries were sequenced on 3 lanes of an Illumina HiSeq2500 with 100 bp MATERIALS AND METHODS paired-end reads. Raw sequence data were cleaned Species Tree Estimation of Family Hyperoliidae following Singhal (2013) and Bi et al. (2012), and the cleaned reads of each sample were de novo assembled, Taxonomic sampling.—We included 254 hyperoliid filtered, and mapped as described in Portik et al. samples in our sequence capture experiment with (2016b). The final filtered assemblies were aligned using multiple representatives per species when possible. MAFFT (Katoh et al. 2002, 2005; Katoh and Standley Although there are 230 currently recognized hyperoliid 2013) and trimmed using trimAl (Capella-Gutierrez species (AmphibiaWeb 2019), this family is in a state et al. 2009). We enforced additional postprocessing of taxonomic flux with recent studies recommending filters for alignments, including a minimum length both the synonymy of species names and the splitting of 90 bp and a maximum of 30% total missing of species complexes (Rödel et al. 2002, 2003, 2009; data across an alignment, resulting in 1047 exon Wollenberg et al. 2007; Schick et al. 2010; Conradie alignments totaling 561,180 bp. Raw sequencing reads et al. 2012, 2013, 2018; Dehling 2012; Greenbaum et al. are deposited in the NCBI Sequence Read Archive 2013; Liedtke et al. 2014; Loader et al. 2015; Portik (BioProject: PRJNA521610), newly generated sequences et al. 2016a; Barratt et al. 2017; Bell et al. 2017a). We for the 5 captured nuclear loci are deposited in estimate that our sampling represents approximately 143 GenBank (Accession numbers: MK497946–MK499204), distinct hyperoliid lineages including 12 of 17 described and all sequence capture alignments are available at hyperoliid genera. The 5 unsampled genera are https://osf.io/ykthm/. either monotypic (Arlequinus, Callixalus, Chrysobatrachus, Kassinula) or species poor (Alexteroon, 3 spp.). Our sampling of the remaining genera is proportional to their Species tree estimation.—We used the sequence capture recognized diversity and includes several known species data set to estimate a species tree for Hyperoliidae complexes with lineages not reflected in the current using ASTRAL-III (Mirarab et al. 2014; Mirarab and taxonomy. We also sampled outgroup species from the Warnow 2015; Zhang et al. 2017), which uses unrooted following families: Arthroleptidae (7 spp), Brevicipitidae gene trees to estimate the species tree. This method (1 sp), Hemisotidae (1 sp), and Microhylidae (1 sp). employs a quartet-based approach that is consistent Museum and locality information for all specimens is under the multispecies coalescent process, and therefore provided in Supplementary Table S1 available on Dryad appropriate for resolving gene tree discordance resulting at http://dx.doi.org/10.5061/dryad.1740n0h. from incomplete lineage sorting (Mirarab et al. 2014; Mirarab and Warnow 2015). This approach also allows Sequence capture data and alignments.—The full details for missing taxa in alignments, which were present of transcriptome sequencing, probe design, library in our sequence capture data and are problematic for preparation, sequence captures, and bioinformatics other coalescent-based summary methods such as MP- pipelines are described in Portik et al. (2016b), but here, EST (Liu et al. 2010). We kept samples with the most we outline major steps of the transcriptome-based exon complete sequence data to collapse the alignments captures. We sequenced, assembled, and filtered the to a single representative per lineage and generated transcriptomes of 4 divergent hyperoliid species and unrooted maximum likelihood (ML) gene trees with selected 1260 orthologous transcripts for probe design. 200 bootstrap replicates for each locus using RAxML v8 We chose transcripts 500–850 bp in length that ranged (Stamatakis 2014) under the GTRCAT model. The set of from 5% to 15% average pairwise divergence. Five 1047 gene trees was used to infer a species tree with additional nuclear loci (POMC, RAG-1, TYR, FICD, and ASTRAL-III, and node support was assessed with (i)

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quartet support values, or local posterior probabilities divergence dating analyses in BEAST v1.8.1 (Drummond computed from gene tree quartet frequencies, which also et al. 2012) involved transforming the ML afrobatrachian allows the calculation of branch lengths in coalescent frog tree to an ultrametric tree, and for this we used units (Sayyari and Mirarab 2016), and (ii) 200 replicates of penalized likelihood with the “chronopl” function of multilocus bootstrapping, where each of the 200 RAxML APE (Sanderson 2002; Paradis et al. 2004), setting age bootstrap trees per locus are used to infer a species tree bounds to allow divergence times to be compatible with and a greedy consensus tree is created from the 200 our BEAST calibration priors. We performed BEAST species trees to calculate percent support across nodes analyses using a ﬁxed ultrametric starting tree topology

(Seo 2008). by removing relevant operators acting on the tree model. Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 We used 4 secondary calibration points with normal distributions to constrain the most recent common Evolutionary Relationships of Afrobatrachian Frogs ancestors (MRCAs) of Afrobatrachia to 80 Ma ± 5 SD, (Hemisotidae + Brevicipitidae) to 50 Ma ± 5 SD, DNA barcoding.—We obtained sequence data from the Arthroleptidae to 40 Ma ± 5 SD, and Hyperoliidae to mitochondrial marker 16S ribosomal RNA (16S) for all 40 Ma ± 5 SD. These secondary calibration points are samples included in the sequence capture experiment based on a consensus of age estimates for afrobatrachian and for additional species that we were unable to include frogs resulting from multiple studies incorporating fossil in our sequence capture experiment due to insufficient calibrations and/or secondary calibrations (Roelants DNA yield. Polymerase chain reactions (PCRs) were et al. 2007; Kurabayashi and Masayuki 2013; Loader carried out in 12.5 L volumes consisting of: 1.25 L et al. 2014; Portik and Blackburn 2016). We used the Roche 10× (500 mM Tris/HCl, 100 mM KCl, 50 mM Yule model of speciation as the tree prior, applied (NH ) SO , 20 mM MgCl , pH = 8.3), 0.75 L25mM 4 2 4 2 an uncorrelated relaxed lognormal clock, and ran 2 MgCl , 0.75 L 2 mM dNTPs, 0.25 L 10.0 Mforward 2 analyses for 30,000,000 generations sampling every 3000 primer, 0.25 L 10.0 M reverse primer, 8.40 LHO, 2 generations. Runs were assessed using TRACER v1.5.0 0.10 L Taq, and 0.75 L DNA. Amplification involved ◦ (Rambaut et al. 2013) to examine convergence, and a initial denaturation at 94 C for 4 min, followed by ◦ ◦ ◦ maximum clade credibility tree with median heights was 35 cycles of 95 C for 60 s, 51 C for 60 s, 72 Cfor ◦ created from 7500 trees after discarding a burn-in of 2500 90 s, and a final extension at 72 C for 7 min, using the trees. primer pairs 16SA and 16SB (Palumbi et al. 1991). The PCR amplifications were visualized on an agarose gel, cleaned using ExoSAP-IT, and sequenced using BigDye v3.1 on an ABI3730 sequencer (Applied Biosystems). Evolution of Sexual Dichromatism and State-Dependent Newly generated 16S sequence data are deposited in Diversification GenBank (Accession numbers: MK509481–MK509743). We scored the presence or absence of sexual dichromatism for hyperoliid species in our data set GenBank data.—To expand our taxonomic sampling, from multiple sources, including publications (Schiøtz we included all available published sequence data of 1967, 1999; Channing 2001; Channing and Howell 2006; afrobatrachian frogs. We built a molecular data matrix Wollenberg et al. 2007; Rödel et al. 2009; Veith et al. 2009; using the 5 nuclear loci included in our captures Amiet 2012; Bell and Zamudio 2012; Channing et al. 2013; (FICD, KIAA2013, POMC, TYR, and RAG-1) and the Conradie et al. 2013; Portik et al. 2016a), examination of mtDNA marker 16S. This resulted in the inclusion of 30 museum specimens (Portik 2015), and the collective field additional hyperoliids and 130 arthroleptid, brevicipitid, observations from all authors. Sexual dichromatism in and hemisotid species, though many are represented hyperoliids is quite pronounced and typically involves solely by 16S mtDNA data. Nuclear loci were aligned differences in both color and pattern. A species was using MUSCLE (Edgar 2004), and 16S sequences were considered sexually dichromatic if adult females and aligned using MAFFT with the E-INS-i algorithm (Katoh adult males exhibit distinct color patterns (Phase F and et al. 2002, 2005). The final concatenated alignment of the Phase J, respectively). We note that although many expanded taxonomic data set consisted of 283 taxa and sexually dichromatic species also exhibit variation in 3991 bp, with 36% total missing data. male color phase (e.g., adult males in the population display Phase J and Phase F), the females of these Phylogenetic methods and divergence dating analyses.— dichromatic species consistently display one color phase We reconstructed the phylogenetic relationships of (Phase F). A summary of the sexual dichromatism afrobatrachian frogs from the 5 nuclear genes and data is provided in Supplementary Table S2 available mtDNA data set using a ML approach in RAxML v8 on Dryad. (Stamatakis 2014). To preserve the relationships inferred We reconstructed the evolution of sexual with our species tree analyses of sequence capture loci, dichromatism on the time-calibrated phylogeny the sequence capture hyperoliid species tree containing of Afrobatrachia using Bayesian ancestral state 153 taxa (143 ingroup and 10 outgroup taxa) was used reconstruction in BEAST (Drummond et al. 2012) as a partial constraint tree for the ML analysis of the and with HiSSE analyses using the R package HiSSE 283-taxon alignment of 6 loci. A preliminary step for our (Beaulieu and O’Meara 2016) (described below). We

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performed Bayesian ancestral state reconstructions trait occur in the phylogeny (Rabosky and Goldberg using several combinations of clock and character 2015; Beaulieu and O’Meara 2016). The improved models in BEAST v1.8.1 (Drummond et al. 2012) character-independent diversification models, referred following King and Lee (2015). The topology and branch to as CID-2 and CID-4, contain the same number of lengths were fixed by removing all tree operators, and diversification rate parameters as the BiSSE and HiSSE sexual dichromatism was treated as a binary alignment. models, respectively. We fit 26 different models to our Because all outgroup families are monochromatic, sexual dichromatism data set (Table 1): 6 represent we fixed the root state by adding a placeholder BiSSE-like models, 4 are variations of the CID-2 model,

monochromatic taxon to the root with a zero branch 5 are variations of the CID-4 model, 9 are various Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 length, creating a hard prior distribution on the root HiSSE models with 2 hidden states, and 2 are HiSSE where P(monochromatic root) =1, and P(dichromatic models with a single hidden state. One variation of CID- root) =0. A stochastic Mk model of character evolution 4 includes the 9-rate model developed by Harrington (Lewis 2001) was used with symmetrical (Mk1) or and Reeder (2017). Within each of these classes, the asymmetrical (Mk2) transition rates between states, models vary mainly in the number of distinct transition and each character model was analyzed using a rates (q), extinction fraction rates (ε), and net turnover strict clock (SC) model (enforcing a homogenous trait rates (), and the most complex HiSSE model includes rate) and a random local clock model (allowing for 4 net turnover rates, 4 extinction fraction rates, and 8 heterotachy), resulting in 4 analysis combinations. distinct transition rates. We enforced a monochromatic The analyses involving the random local clock allowed root state for all models and evaluated the fit of the estimation of the number and magnitude of rate changes 26 models using AIC scores, AIC scores, and Akaike ω using MCMC. All analyses were run for 200 million weights ( i)(Burnham and Anderson 2002). From the generations with sampling every 20,000 generations, best-fit model, we estimated confidence intervals for resulting in 10,000 retained samples. We used the relevant parameters and transformed ε and to obtain marginal likelihood estimator with stepping stone speciation (), extinction (), and net diversification sampling, with a chain length of 1 million and 24 path rates using the “SupportRegion” function in HiSSE steps, to estimate the log-marginal likelihood of each (Beaulieu and O’Meara 2016). We performed ancestral run (Baele et al. 2012, 2013). We performed 5 replicates state estimations for each of the 26 models using the per analysis to ensure consistency in the estimated marginal reconstruction algorithm implemented in the log-marginal likelihood, and subsequently compared “MarginRecon” function of HiSSE, again enforcing a the 4 different analyses using log-Bayes factors, monochromatic root state. Our final estimation and calculated as the difference in log-marginal likelihoods, visualization of diversification rates and node character to select the best fit clock model and character model states on the afrobatrachian phylogeny took model combination. We summarized transitions between uncertainty into account by using the model averaging character states and created consensus trees to estimate approach described by Beaulieu and O’Meara (2016), the posterior probabilities of character states across such that model contributions to rates and states were nodes. proportional to their likelihoods. We performed HiSSE analyses using the R package HiSSE (Beaulieu and O’Meara 2016) to identify if sexual dichromatism in afrobatrachian frogs is associated Simulated traits and state-dependent diversification.—The with increased diversification rates, and to reconstruct identified bias toward the detection of trait-dependent ancestral states while accounting for transition rate diversification in the BiSSE framework (Rabosky and and diversification rate heterogeneity. The HiSSE model Goldberg 2015; Beaulieu and O’Meara 2016) prompted builds upon the popular binary-state speciation and us to investigate if a similar outcome would be detected extinction (BiSSE) model (Maddison et al. 2007)by in our data set, and whether the HiSSE framework could incorporating “hidden states” representing unmeasured improve our ability to distinguish whether the observed traits that could impact the diversification rates states are correlated with diversification rates. One estimated for states of the observed trait. The HiSSE concern raised by Beaulieu and O’Meara (2016)isthat model is therefore able to account for diversification neutrally evolving traits simulated on trees generated rate heterogeneity that is not linked to the observed from a complex heterogenous rate branching process can trait, while still identifying trait-dependent processes. lead to false signals of trait-dependent diversification, The HiSSE framework also includes a set of null signifying the HiSSE framework may be sensitive to models that explicitly assume the diversification process particular types of tree shapes. is independent from the observed trait, without To evaluate the performance of these methods given constraining diversification rates to be homogenous our empirical tree topology, we simulated neutrally across the tree. The inclusion of these character- evolving traits on the afrobatrachian frog phylogeny independent models circumvents a significant problem and tested for trait-dependent diversification using identified in the BiSSE framework, in which the simple both the BiSSE and HiSSE frameworks. We conducted “null” model of constant diversification rates is typically independent simulations of a binary trait with the rejected in favor of trait-dependent diversification “sim.history” function in R package PHYTOOLS (Revell when diversification rate shifts unrelated to the 2012) using the unequal rates q-matrices obtained from

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TABLE 1. Summary of models and ﬁts using HiSSE analyses, with sexual dichromatism as the observed state Net Extinction Distinct Hidden State- turnover fraction transition Description Model Loglik AIC AIC wtAIC States dependent rates rates rates HiSSE 19 −1033.5 2082.9 0.0 0.445 Yes Yes 4 1 3 HiSSE 15 −1026.0 2084.01.00.266 Yes Yes 4 4 8 HiSSE 18 −1029.7 2085.42.50.130 Yes Yes 4 1 8 CID-4 14 −1035.3 2086.53.60.075 Yes No 4 1 3 CID-2 8 −1031.7 2087.34.40.049 Yes No 2 2 8

HiSSE 16 −1033.5 2088.96.00.022 Yes Yes 4 4 3 Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 HiSSE 25 −1036.1 2092.29.30.004 only for state 0 Yes 3 3 4 CID-4 12 −1035.3 2092.59.60.004 Yes No 4 4 3 CID-4 26 −1029.7 2093.410.40.002 Yes No 4 4 9 CID-2 10 −1036.2 2094.411.5 <0.001 Yes No 2 1 8 HiSSE 24 −1041.12102.319.3 <0.001 only for state 1 Yes 3 3 4 BiSSE-like 2 −1046.32102.619.7 <0.001 No Yes 2 1 2 BiSSE-like 1 −1046.32104.621.7 <0.001 No Yes 2 2 2 BiSSE-like 3 −1053.52115.132.1 <0.001 No No 1 1 2 HiSSE 22 −1055.72121.538.5 <0.001 Yes Yes 4 1 3 HiSSE 21 −1053.62127.244.3 <0.001 Yes Yes 4 1 8 CID-4 13 −1060.2 2132.449.4 <0.001 Yes No 4 1 1 HiSSE 20 −1061.9 2135.752.8 <0.001 Yes Yes 4 1 1 HiSSE 17 −1060.4 2138.755.8 <0.001 Yes Yes 4 4 1 CID-4 11 −1060.4 2138.855.8 <0.001 Yes No 4 4 1 CID-2 9 −1068.12144.261.3 <0.001 Yes No 2 1 1 CID-2 7 −1068.12146.263.3 <0.001 Yes No 2 2 1 BiSSE-like 4 −1071.42152.869.8 <0.001 No Yes 2 2 1 HiSSE 23 −1073.92153.770.8 <0.001 Yes Yes 4 1 1 BiSSE-like 5 −1075.22158.375.4 <0.001 No Yes 2 1 1 BiSSE-like 6 −1077.62161.278.3 <0.001 No No 1 1 1 Note: The top ranked model is highlighted in bold.

our empirical data and enforcing a root state of trait RESULTS absence. We required a minimum of 10% of taxa to exhibit the derived state and conducted simulations until Phylogenetic Relationships we obtained 1500 replicates meeting this criterion. We The sequence capture data set consisting of 1047 used ML to fit a BiSSE model and the typical “null” loci and 561,180 bp produced a well-resolved species model with equal speciation and extinction rates to tree with a normalized quartet score of 0.877 and each simulated trait using the R package DIVERSITREE only 8 of 150 nodes (5%) with quartet scores below (Maddison et al. 2007; FitzJohn et al. 2009; FitzJohn 2012). 0.9 (Supplementary Fig. S1 available on Dryad). The We performed likelihood ratio tests and calculated AIC multilocus bootstrapping analysis produced similar scores to determine if the constraint model could be results with low support for only 7 nodes, 6 of which rejected with confidence (AIC > 2orP-value < 0.05), also received low support in the species tree analyses and summarized the number of instances each of the 2 (Supplementary Fig. S1 available on Dryad). The higher- models was favored across the simulations. We analyzed level relationships recovered in the species tree are the simulated data in the HiSSE framework as with largely congruent with those recovered by Portik and our empirical data, but with a reduced set of 5 models Blackburn (2016), though we found strong support representing each major category of model. This reduced for a different placement of the genus Acanthixalus as model set included 2 BiSSE-like models that differed sister to the clade containing Semnodactylus, Paracassina, only in the constraint of , a CID-2 and CID-4 model, Phlyctimantis, and Kassina, which together are sister to and a HiSSE model with 2 hidden states, 3 transition all other hyperoliids. Our improved sampling provides rates, equal ε, and distinct . We set a probability of one the first comprehensive assessment of evolutionary for trait absence at the root state to match the manner in relationships within the speciose genera Afrixalus and which traits were simulated and evaluated the fit of the Hyperolius. The monophyly of Hyperolius is supported, ω 5 models using AIC scores and Akaike weights ( i)for however Afrixalus is paraphyletic, and we found each simulation, using a threshold of AIC greater than a sister relationship between the Ethiopian-endemic 2 to favor a model. Specifically, we were interested in Afrixalus enseticola and the Malagasy-Seychelles species whether the unconstrained BiSSE-like or HiSSE models of Heterixalus and Tachycnemis, which are in turn were favored, resulting in the detection of a false pattern sister to all remaining Afrixalus (Supplementary Fig. of trait-dependent diversification, or if the CID-2 or CID- S1 available on Dryad). We found strong support for 4 models were selected, capturing the expected pattern the southern African species Hyperolius semidiscus as where the diversification process was independent from sister to all other lineages in the genus, and furthermore trait evolution. recovered Hyperolius parkeri, Hyperolius lupiroensis, and

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Dichromatic Hemisotidae Brevicipitidae Monochromatic Arthroleptidae Hyperoliidae

SemnodactylusAcant hixalus Paracassina

Phlyctimantis Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019

Kassina

Opisthothylax Kassininae A. enseticola Tachycnemis

Hyperoliidae Heterixalus

Hyperoliinae

Afrixalus

H. nasutus Clade 2 com plex

Clade 1

tothylax rerella CrypMo Net Diversification

lius

ypero H

0.003 0.16

FIGURE 2. Ancestral state reconstruction of sexual dichromatism in afrobatrachian frogs from HiSSE using model averaging to account for uncertainty in both models and reconstructions. Circles at tips and nodes are colored by state with node pie charts indicating probability of a state assignment. Branches are colored using a gradient from the model-averaged net diversiﬁcation rate, with black representing the slowest rate. The arrow highlights the node identiﬁed as displaying an unambiguous transition from monochromatism to sexual dichromatism.

the Hyperolius nasutus complex as sister to 2 larger assignments for genera not sampled in our molecular subclades of Hyperolius (Clades 1 and 2; Fig. 2, study (Hyperoliinae: Alexteroon, Arlequinus, Callixalus, Supplementary Fig. S1 available on Dryad). In an effort Chrysobatrachus, Kassinula), which should be included in to distinguish the main division within Hyperoliidae, future phylogenetic studies to conﬁrm these placements. we recognize the subfamilies Kassininae Laurent, 1972 The phylogenetic analyses of the Afrobatrachia and Hyperoliinae Laurent, 1943 and deﬁne the content supermatrix produced family-level relationships within each on the basis of our species tree analysis as consistent with previous analyses (Pyron and Wiens follows: (i) Kassininae: Acanthixalus, Kassina, Paracassina, 2011; Portik and Blackburn 2016; Feng et al. 2017), Phlyctimantis, and Semnodactylus; (ii) Hyperoliinae: including a sister relationship between Hyperoliidae Afrixalus, Cryptothylax, Heterixalus, Hyperolius, Morerella, and Arthroleptidae, and between Hemisotidae and and Opisthothylax (Fig. 2). We retain previous subfamily Brevicipitidae (Supplementary Fig. S2 available on

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TABLE 2. Summary of trait rate analyses for the evolution of sexual dichromatism Log- marginal Transitions Transitions Rate Rate Rate Trait rate analysis likelihood 0 > 11> 0 shifts minimum maximum Strict clock Mk1 −92.12 15 7 – – – Strict clock Mk2 −81.83 2 27 – – – Relaxed local clock Mk1 −85.10 16 8 3.30.001 0.019 Relaxed local clock Mk2 −81.73 2 27 0.80.013 0.016 Note: For transition summaries, monochromatism is coded as “0” and dichromatism as “1.” The average number of rate shifts is provided, rather Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 than median.

Dryad). This expanded taxonomic data set also included character reconstructions revealed that this estimate improved sampling for the Malagasy hyperoliid genus is due to low posterior probability estimates for 2 Heterixalus and the H. nasutus complex, for which we critical nodes within Hyperoliinae (Supplementary Fig. recovered results consistent with Wollenberg et al. (2007) S3 available on Dryad). The presence of dichromatism and Channing et al. (2013), respectively. We recovered at one or both of these nodes results in a single an Eocene age for the time to most recent common origin of sexual dichromatism. In contrast, the SC Mk2 ancestor (TMRCA) of the families Hyperoliidae, model demonstrated approximately 27 independent Arthroleptidae, and Brevicipitidae, of approximately losses of sexual dichromatism within hyperoliids, 42.4 Ma (95% highest posterior density region [HPD]: including losses in entire groups (Afrixalus, H. nasutus 39.1–51.6 Ma), 45.4 Ma (95% HPD: 35.5–48.7 Ma), and complex) and numerous reversals within Heterixalus and 41.8 Ma (95% HPD: 33.9–49.8 Ma) (Supplementary Fig. Hyperolius (notably in Clade 2) (Supplementary Fig. S4 S2 available on Dryad). These dates are younger than available on Dryad). previous estimates (Roelants et al. 2007; Loader et al. Our HiSSE analyses revealed that a HiSSE model 2014) but are consistent with more recent multilocus with 4 net turnover rates, equal extinction fraction phylogenetic analyses (Portik and Blackburn 2016; Feng rates, and 3 distinct transition rates was the best- et al. 2017). We found diversification events began fit model (Model 19, Table 1). The second and third within Hyperoliinae approximately 35.8 Ma (95% HPD: ranked models (AIC of 1.0, 2.5) were also HiSSE 30.1–41.6 Ma) and within Kassininae approximately 38.3 models that varied in the number of extinction fraction Ma (95% HPD: 30.9–45.6 Ma). The majority of speciation rates or the number of transition rates. Together these events within Hyperolius occurred from the late Miocene 3 HiSSE models accounted for 84.1% of the model to the Plio-Pleistocene (Supplementary Fig. S2 available weight (Table 1) and support a signal of state-dependent on Dryad). diversification in which sexual dichromatism and a hidden state are associated with diversification rates. The net diversification rates inferred using the best-fit Evolution of Sexual Dichromatism and State-Dependent model were nearly twice as high in sexually dichromatic Diversification lineages (0.157) as compared with monochromatic lineages (0.091) in the absence of the hidden state We determined sexual dichromatism occurs in 60 of (Fig. 3), and the combination of the hidden state the 173 (34%) hyperoliid frog species included in our and dichromatism or monochromatism resulted in analyses. The greatest number of dichromatic species much lower net diversification rate estimates (0.02 and Hyperolius occurs in Clade 1 (35 of 39 species, 89%), <0.001, respectively). The trait reconstructions for all Hyperolius followed by Clade 2 (18 of 50 species, 36%) and 4 state combinations indicated that the dichromatism Heterixalus the genus (4 of 11 species, 36%) (Figs. 2 and plus hidden state combination occurs in the MRCA 5). The Bayesian ancestral state reconstructions using of 2 species-poor or monotypic genera (Cryptothylax, 4 model combinations produced different patterns of Morerella)(Supplementary Fig. S5 available on Dryad), character reconstructions and rates of trait evolution and is associated with a markedly lower diversification (Supplementary Fig. S3 available on Dryad). Overall, we rate (Fig. 3). The model-averaged ancestral state found models with asymmetric character transition rates reconstructions demonstrated strong support for a single (Mk2) outperformed those with symmetric rates (Mk1), origin of sexual dichromatism and 25 independent regardless of the clock model used (log-transformed reversals to monochromatism within Hyperoliinae, with Bayes factor range: 3.40–10.39; Table 2). The analysis reversal patterns similar to the Mk2 analyses (Fig. 2 and with the highest log-marginal likelihood incorporated Supplementary Fig. S5 available on Dryad). the relaxed local clock Mk2 model, though it was not a significantly better fit than the simpler SC Mk2 model (log-transformed Bayes factor of 0.10) indicating there Simulated traits and state-dependent diversification.— is low variation in lineage-specific evolutionary rates Analyzed in the BiSSE framework, we found that for this trait. The ancestral character reconstructions of many of the trait simulations on the phylogeny of the SC Mk2 model indicated a median of 2 transitions Afrobatrachia resulted in the rejection of the “null” from monochromatism to dichromatism. However, the model of equal diversification rates across character

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0.20 a) 350 BiSSE model comparisons ‘Null’ 300 BiSSE 250 0.15 200

Frequency 150

100 Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 0.10 50

0 0102051525 et Diversification Rate et Diversification

N 0.05 b) 350 HiSSE model comparisons CID-4 300 HiSSE 250

0 200

Mono/- Dichro/- Mono/+ Dichro/+ 150 Frequency 100 FIGURE 3. The net diversification rate confidence intervals estimated from the best-fit HiSSE model for various combinations of 50 monochromatism, dichromatism, and the presence (+) or absence (−) 0 of the hidden state. 02468 10 12 14 Delta AIC score states. Based on the significance of likelihood ratio tests, we rejected the “null” model in favor of trait- FIGURE 4. Histograms of the delta AIC comparisons resulting from dependent diversification for 565 (37.6%) of our 1500 (a) BiSSE analyses and (b) HiSSE analyses of the 1500 simulations of neutrally evolving traits on the Afrobatrachia phylogeny. The vertical comparisons. We recovered similar results using a delta dotted line represents a delta AIC of two. In the BiSSE analyses (a), AIC cutoff value of 2, for which we found support delta AIC scores of less than two are considered a failure to reject the for trait-dependent diversification in 546 (36.4%) of the “null” model of constant diversification rates, whereas scores above simulations (Fig. 4). In many of these cases, we found two for the BiSSE model are interpreted as favoring a correlation between diversification rates and the simulated neutral trait. The unexpectedly strong support for the BiSSE model, and HiSSE analyses (b) included 5 models, but only two were consistently 160 (10.6%) of the comparisons resulted in delta AIC selected as the top ranking, including the HiSSE and CID-4 (character- values ranging from 10 to 40. independent diversification) models. For these analyses delta AIC > 2 In contrast to the BiSSE analyses, the addition of was considered as support for a particular model, whereas delta AIC < the character independent models (CID-2 and CID-4) 2 was considered as equivocal support for the model. in HiSSE dramatically reduced the detection of a false association between simulated traits and diversification problematic for the investigation of trait-dependent rates by providing appropriate null models. In addition diversification using these available methods, adding to the 2 CID models, our set of 5 models also included support to our empirical analyses in which we detected 2 BiSSE-like models and a typical HiSSE model. Based an association between diversification rates and sexual on a delta AIC cutoff value of 2, out of the 1500 dichromatism. analyses performed the CID-4 model was selected 1132 times (75.4%), the HiSSE model was chosen 125 times DISCUSSION (8.3%), and the remaining 243 analyses (16.2%) showed equivocal support (AIC = 0–2) for either the CID-4 or Hyperoliid Relationships and the Origin of Sexual HiSSE model (Fig. 4). In the cases of equivocal support, Dichromatism the CID-4 and HiSSE models were always the top 2 Afrobatrachian frogs account for more than half of models, which should be interpreted as a lack of support all African amphibians and this continental radiation for trait-dependent diversification. Our error rate with exhibits incredible variation in ecomorphology, the HiSSE model being favored in only 8.3% of our reproductive mode, and other life history traits (Portik simulations was substantially lower than the Beaulieu and Blackburn 2016). Within Afrobatrachia, the family and O’Meara (2016) “difficult tree” scenario in which Hyperoliidae is the most species-rich (∼230 species) the HiSSE model was favored in 29% of the data sets. with surprisingly little diversity in ecomorphology These simulation results strongly suggest the branching and reproductive mode (Schiøtz 1967, 1999; Portik pattern of our empirical phylogeny is not inherently and Blackburn 2016), but with incredible variation in

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coloration and sexual dichromatism. These phenotypic monochromatism result directly from differences characteristics have generated considerable taxonomic in hormone sensitivities among species. Reversals to confusion within hyperoliids (e.g., Ahl 1931), hindering monochromatism are especially prominent in Hyperolius a clear understanding of species diversity and the Clade 2 (12 independent events; Fig. 2, Supplementary evolutionary history of this radiation. Here, we have Fig. S4 available on Dryad), highlighting the potential for produced the most comprehensive hyperoliid species future research in this group to identify the physiological tree to date and found support for an Eocene origin basis and molecular underpinnings of these transitions. of 2 subfamilies representing a major division within For example, 3 monochromatic species of the Hyperolius

Hyperoliidae: Kassininae (26 species) and Hyperoliinae cinnamomeoventris complex that are endemic to the Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 (∼200 species). Within Hyperoliinae, we clarified islands of São Tomé and Príncipe (Hyperolius drewesi, relationships within the hyperdiverse genus Hyperolius Hyperolius molleri, Hyperolius thomensis;bothsexes (∼150 species), which consists of 2 major clades (Fig. 2, with Phase F coloration) are derived from a mainland Supplementary Figs. S1 and S2 available on Dryad). clade containing a mix of sexually dichromatic species These clades represent parallel radiations with species (Hyperolius cinnamomeoventris, Hyperolius olivaceus; distributed across a variety of habitats and altitudes females Phase F, males Phase J) and monochromatic throughout sub-Saharan Africa, which showcases species (Hyperolius veithi; both sexes Phase J) (Fig. 5). Hyperoliidae as a rich comparative framework for Variation among these closely related species is well future biogeographic research. suited for investigating both the evolutionary and Although both Kassininae and Hyperoliinae include ecological contexts underlying transitions from sexual colorful species, sexual dichromatism only occurs dichromatism to monochromatism. within Hyperoliinae where it is present in the genera Tachycnemis, Cryptothylax, and Morerella, several species Sexual Dichromatism Is Linked to Increased Diversification of Heterixalus, and more than half of the Hyperolius species we sampled (Fig. 2, Supplementary Table S2 Rates available on Dryad). A previous study hypothesized Sexual dichromatism is an important predictor of that sexual dichromatism evolved multiple times diversification in several taxonomic groups including within Hyperoliidae (Veith et al. 2009); however, we cichlids (Wagner et al. 2012), labrid fishes (Alfaro et al. found overwhelming support for a single origin of 2009; Kazancıoglu et al. 2009), agamid lizards (Stuart- sexual dichromatism (Fig. 2, Supplementary Fig. S3 Fox and Owens 2003), and dragonflies (Misof 2002). available on Dryad) and over 20 independent reversals There is no such association in bees (Blaimer et al. 2018), to monochromatism that range from early transitions and in birds the relationship between dichromatism that characterize entire genera (e.g., Afrixalus) to recent and species richness/speciation rate varies among reversals within Heterixalus and Hyperolius (Fig. 2, studies that differ in methodology and taxonomic scale Supplementary Fig. S4 available on Dryad). This (Barraclough et al. 1995; Owens et al. 1999; Morrow transition bias from dichromatism to monochromatism et al. 2003; Phillimore et al. 2006; Seddon et al. also occurs in birds (Price and Birch 1996; Omland 2013; Huang and Rabosky 2014). Some state-dependent 1997; Burns 1998; Kimball et al. 2001; Hofmann et al. speciation and extinction model sets, such as the BiSSE 2008; Dunn et al. 2015; Shultz and Burns 2017), in which method, are no longer considered adequate for robustly secondary monochromatism can evolve as the result detecting trait-dependent diversification (Rabosky and of a color change in either sex (Kimball and Ligon Goldberg 2015; Beaulieu and O’Meara 2016). In contrast 1999; Johnson et al. 2013; Price and Eaton 2014; Dunn to BiSSE, the HiSSE method contains an expanded model et al. 2015). Similar transitional pathways to secondary set that can link diversification rate heterogeneity to monochromatism occur in hyperoliids, in which observed traits, hidden traits, or character independent females may lose the ability for color transformation processes. In particular, the inclusion of appropriate at sexual maturity (both sexes retain the Phase J null models (character-independent models) reduces juvenile coloration) or males undergo obligatory the inference of trait-dependent diversification when a ontogenetic color change at maturity (both sexes phylogeny contains diversification rate shifts unrelated develop Phase F coloration) (Figs. 1, 5). Characterizing to the focal trait (Beaulieu and O’Meara 2016). Our juvenile coloration across species (which remains trait simulation study recapitulated this result (Fig. 4) unknown in many hyperoliids) and documenting while demonstrating that the branching pattern of ontogenetic color change (including the prevalence our Afrobatrachia phylogeny is unlikely to drive of male color phases, e.g., Portik et al. 2016a) will be false inferences of trait-dependent diversification using essential steps toward differentiating between these 2 HiSSE (e.g., the “worst-case” scenario of Beaulieu and forms of monochromatism. In experimental settings, O’Meara 2016). We found that sexually dichromatic the hormone estradiol induces color transformation hyperoliid lineages have nearly double the average in both sexes in the dichromatic species Hyperolius diversification rate of monochromatic lineages, and argus and Hyperolius viridiflavus. Conversely, the that these diversification rates are not inflated by a effects of testosterone appear to differ across species hidden trait. The shift to the sexual dichromatism (Richards 1982; Hayes and Menendez 1999), suggesting plus hidden state character combination occurred only that evolutionary transitions from dichromatism to once in the common ancestor of 2 genera (Cryptothylax,

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FIGURE 5. Illustration of (a) several Hyperolius species in the predominately sexually dichromatic Clade 1 and (b) several Hyperolius species in Clade 2 that exhibit multiple transitions to secondary monochromatism. Males are positioned in the top rows, with females below, and the phylogenetic relationships among species are depicted (though not all taxa have been included) Photo credits: Daniel Portik, Jos Kielgast, Bryan Stuart, Andrew Stanbridge.

Morerella; Supplementary Fig. S5 available on Dryad) closely related species with high ecological similarity and is actually associated with lower diversification but differing almost exclusively in signaling traits (West- rates (Fig. 3). Together, our results demonstrate Eberhard 1983; Dominey 1984; Panhuis et al. 2001; that diversification rate heterogeneity occurs within Turelli et al. 2001; Andersson 1994; Mendelson and Afrobatrachia, and that the origin and persistence of Shaw 2012; Ritchie 2007; Safran et al. 2013). Among sexual dichromatism in hyperoliid frogs is linked to their frog species with similar morphologies (e.g., treefrogs), rapid diversification across sub-Saharan Africa. ecological divergence primarily results from changes in body size. Frogs are generally opportunistic, gape- limited predators (Duellman and Trueb 1986), and in How Does Sexual Dichromatism Influence Diversification hyperoliids food partitioning is strongly dictated by Rate? body size (Luiselli et al. 2004). At one well-characterized Sexual dichromatism is a common proxy for sexual community site, the females of 7 sexually dichromatic selection in macroevolutionary studies (reviewed in Hyperolius species display minimal differences in body Kraaijeveld et al. 2011), especially for testing the size (Portik et al. 2018) but exceptional divergence in prediction that clades with variation in secondary sexual color (Portik et al. 2016a). Five of these species occur in characters under strong sexual selection exhibit higher Hyperolius Clade 1, and within this clade there tend to diversification rates and species richness (Lande 1981, be striking interspecific color differences in females, but 1982; West-Eberhard 1983; Barraclough et al. 1995). not males, among closely species (Fig. 5). This pattern Although our trait-dependent diversification analyses of interspecific variation in secondary sexual characters strongly support a faster rate of net diversification in the absence of ecological divergence is consistent associated with sexual dichromatism, we currently lack with the predictions of speciation by sexual selection. behavioral and natural history data in hyperoliids to By inference, this would imply that dichromatism in establish whether sexual dichromatism is actually under hyperoliids—specifically female color—is an essential strong sexual selection. At more recent timescales, mate recognition signal (sensu Mendelson and Shaw speciation by sexual selection can result in a group of 2012). In frogs, male calls are well-established mate

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recognition signals that can be under strong sexual microhabitats, and as a consequence are subject to selection (Ryan 1980; Gerhardt 1994), and these acoustic different selective pressures and predation regimes signals are demonstrably important for hyperoliid (Heinsohn et al. 2005; Bell and Zamudio 2012). During frogs. Males form dense breeding aggregations and the breeding season, male hyperoliids are exposed choruses (Bishop et al. 1995), calls differ between on calling sites and Hayes (1997) proposed that more closely related species (Schiøtz 1967, 1999; Gilbert cryptic male coloration may reduce predation pressure. and Bell 2018) and females prefer conspeciﬁc calls This hypothesis is consistent with a greater number over calls of syntopic heterospeciﬁcs (Telford and of observations of predation events on females of

Passmore 1981). However, visual displays can also dichromatic species (Grafe 1997; Portik et al. 2018). Downloaded from https://academic.oup.com/sysbio/article-abstract/68/6/859/5477408 by Ustav biologie obratlovcu AV CR, v.v.i. user on 23 October 2019 serve as important courtship signals in frogs, typically Quantifying differences in predation rates between in conjunction with acoustic signaling (Gomez et al. the sexes, between monochromatic and dichromatic 2009, 2010; Starnberger et al. 2014; Jacobs et al. 2016; species, and between Phase F and Phase J males within Yovanovich et al. 2017; Akopyan et al. 2018). Although dichromatic species would address whether this aspect most studies have documented female preference for of natural selection is also shaping the evolution of sexual male coloration, the recent discovery of female displays dichromatism. Although these mechanisms and other during nocturnal phyllomedusine treefrog courtship differences in natural selection pressures may influence highlights the possibility of male mate-choice and the evolution of sexual dichromatism in hyperoliids, intersexual selection of female coloration (Akopyan et al. they are generally not expected to elevate rates of 2018). The notable lack of heterospecific matings among reproductive isolation or drive diversification rate shifts dichromatic species at high-diversity breeding sites comparable to the effects of sexual selection. Therefore, (Portik et al. 2018) strongly suggests behavioral isolation we propose that hyperoliids are a compelling system for may be linked to divergent mate recognition signals, disentangling the roles of sexual selection and natural including male call (Telford and Passmore 1981), gular selection in the evolution of sexual dichromatism, and gland compounds (Starnberger et al. 2013), or female how these mechanisms have promoted diversification coloration. Our knowledge of reproductive behavior in at both microevolutionary and macroevolutionary hyperoliids is largely based on a single dichromatic timescales. species (H. marmoratus; Dyson and Passmore 1988; Telford and Dyson 1988; Dyson et al. 1992; Jennions et al. 1995), and as such there may be an overlooked role for mutual mate-choice in hyperoliids in which SUPPLEMENTARY MATERIAL females locate males by call and/or pheromones and Data available from the Dryad Digital Repository: males assess color patterns of approaching females. http://dx.doi.org/10.5061/dryad.1740n0h. Although it can be tempting to equate sexually dimorphic traits such as dichromatism with sexual selection, several alternative mechanisms may FUNDING also contribute to female-biased dichromatism in Molecular data collection by DMP was funded by hyperoliids. For instance, aposematism is a widespread a National Science Foundation Doctoral Dissertation anti-predator mechanism in frogs that is typically Improvement Grant (DEB: 1311006), an Ecological, accompanied by the presence of skin toxins (reviewed in Evolutionary, and Conservation Genomics Research Toledo and Haddad 2009; Rojas 2017) such as alkaloids Award presented to DMP by the American Genetic (Daly 1995). Sex-specific differences in chemical defense Association, a National Science Foundation grant (DEB: have been documented in some frogs (Saporito et al. 1202609) awarded to DCB, a UC Berkeley President’s 2010; Jeckel et al. 2015), but in several dichromatic Postdoctoral Fellowship awarded to RCB, and by the hyperoliid species neither sex contained alkaloids in Museum of Vertebrate Zoology. their skin (Portik et al. 2015). This finding suggests that either aposematism is an unlikely explanation for ornate coloration or that hyperoliids have evolved ACKNOWLEDGMENTS novel compounds for chemical defense. Female-biased We thank the following institutions for accessioning dichromatism has also been tied to sex-role reversal field collections and for facilitating loan access: in fishes and birds (Roede 1972; Oring 1982; Berglund California Academy of Sciences, Cornell University et al. 1986a,b; Eens and Pinxten 2000), in which females Museum of Vertebrates, Institut National de Recherche compete more intensely than males for access to en Sciences Exactes et Naturelles, Museum für mates. This mechanism seems unlikely for hyperoliids Naturkunde, Berlin, Museìum National d’Histoire because males in both monochromatic and dichromatic Naturelle, Museum of Comparative Zoology,Museum of species form dense choruses and compete intensely Vertebrate Zoology, National Museum Prague, Natural to attract females, often engaging in combat (Telford History Museum London, North Carolina Museum of 1985; Backwell and Passmore 1990). Finally, sexual Natural Sciences, Port Elizabeth Museum, Senckenberg niche partitioning (Selander 1966; Shine 1989) can result Natural History Collections Dresden, South African in sexual dichromatism when the sexes use different Institute for Aquatic Biodiversity, South African

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National Biodiversity Institute, The Field Museum, AmphibiaWeb. 2019. Available at: amphibiaweb.org. University of Trento Museum of Science, University of Texas at El Paso California, Berkeley. Accessed May 2018. Biodiversity Collections, and the Zoological Natural Andersson M. 1994. Sexual selection. Princeton (NJ): Princeton University Press. History Museum, Addis Ababa University. All authors Backwell P.R.Y., Passmore N.I. 1990. Aggressive interactions and express thanks to the many government agencies, intermale spacing in choruses of the leaf-folding frog, Afrixalus ministries, and departments that issued research permits delicatus. S. Afr. J. Zool. 25:133–137. and provided the access necessary to conduct their Baele G., Lemey P., Bedford T., Rambaut A., Suchard M.A., Alekseyenko A.V. 2012. Improving the accuracy of demographic individual ﬁeld work in sub-Saharan Africa. This and molecular clock model comparison while accommodating

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[16:55 25/9/2019 Sysbio-OP-SYSB190025.tex] Page: 875 859–875 Phrynomantis microps Microhylidae Breviceps adspersus Brevicipitidae Hemisus marmoratus Hemisotidae Leptopelis rufus Cardioglossa gracilis Arthtroleptis poecilinotus Leptodactylodon ovatus Arthroleptidae Nyctibates corrugatus Trichobatrachus robustus Scotobleps gabonicus Acanthixalus spinosus Acanthixalus sonjae Semnodactylus wealli Paracassina obscura Paracassina kounhiensis Kassininae Phlyctimantis maculata Phlyctimantis verrucosus Phlyctimantis boulengeri Phlyctimantis leonardi Kassina kuvangensis Kassina fusca Kassina lamottei Kassina senegalensis Kassina decorata Kassina maculosa Kassina arboricola Hyperoliidae Kassina cochranae Opisthothylax immaculatus Afrixalus enseticola Tachycnemis seychellensis Heterixalus luteostriatus Malagasy-Seychelles Heterixalus alboguttatus Species Afrixalus vibekensis Afrixalus weidholzi Afrixalus lacustris Afrixalus dorsimaculatus 1 Afrixalus dorsimaculatus 2 Afrixalus knysae Afrixalus spinifrons Hyperoliinae Afrixalus delicatus Afrixalus sylvaticus Afrixalus brachycnemis Afrixalus laevis Afrixalus lacteus Afrixalus fornasini Afrixalus wittei Afrixalus osorioi Afrixalus paradorsalis paradorsalis 2 Afrixalus paradorsalis paradorsalis 1 Afrixalus paradorsalis manengubensis Afrixalus nigeriensis Afrixalus vittiger 1 Afrixalus vittiger 2 Afrixalus dorsalis 3 Afrixalus dorsalis 1 Afrixalus quadrivittatus 1 Afrixalus quadrivittatus 2 Afrixalus dorsalis 2 Afrixalus fulvovittatus 2 Afrixalus fulvovittatus 1 Morerella cyanophthalma Cryptothylax greshoffii Hyperolius semidiscus Hyperolius parkeri Hyperolius pusillus Hyperolius adspersus Hyperolius pseudoargus Hyperolius fusciventris ssp. Hyperolius fusciventris burtoni Hyperolius bolifambae Hyperolius ocellatus ocellatus Hyperolius ocellatus purpurescens Clade 1 Hyperolius sp. nov. ‘Kupe’ Hyperolius camerunensis Hyperolius riggenbachi Hyperolius guttulatus 2 Hyperolius guttulatus 1 Hyperolius phantasticus Hyperolius pardalis Hyperolius steindachneri Hyperolius argus Hyperolius bocagei Hyperolius lamottei Hyperolius tuberculatus Hyperolius dintelmanni Hyperolius parallelus parallelus Hyperolius parallelus marginatus Hyperolius parallelus alborufus Hyperolius parallelus pyrrhodictyon Hyperolius marmoratus taeniatus Hyperolius mariae Hyperolius reesi Hyperolius marmoratus marmoratus Hyperolius marmoratus swynnertoni Hyperolius nitidulus 1 Hyperolius spatzi Hyperolius nitidulus 2 Hyperolius viridiflavus ngorongoriensis Hyperolius viridiflavus ommatostictus Hyperolius viridiflavus pantherinus Hyperolius viridiflavus glandicolor Hyperolius viridiflavus pitmani Hyperolius viridiflavus ssp. Hyperolius viridiflavus destefanii Hyperolius viridiflavus variablilis Hyperolius koehleri Hyperolius tuberilinguis Hyperolius mosaicus Hyperolius endjami 2 Clade 2 Hyperolius endjami 1 Hyperolius cystocandicans Hyperolius jackie Hyperolius discodactylus Hyperolius lateralis Hyperolius castaneus Hyperolius frontalis Hyperolius platyceps Hyperolius raymondii Hyperolius langi Hyperolius kuligae Hyperolius ademetzi Hyperolius tanneri Hyperolius minitissimus Hyperolius burgessi Hyperolius spinigularis Hyperolius torrentis Hyperolius chlorosteus Hyperoliid Species Tree Analysis (ASTRAL-III) Hyperolius occidentalis Hyperolius sylvaticus Hyperolius baumanni 1,047 exons Hyperolius picturatus 561,180 bp Hyperolius bobirensis Hyperolius concolor concolor Hyperolius concolor guttatus Hyperolius concolor ibadanensis Branch Length Scale Hyperolius montanus Hyperolius pictus Hyperolius substriatus 2.0 Coalescent Units Hyperolius stictus Hyperolius mitchelli Hyperolius rubrovermiculatus Hyperolius quinquevittatus Node Support Key Hyperolius kivuensis 1 Hyperolius kivuensis 3 quartet support > 0.9; multilocus bootstrap > 70% Hyperolius kivuensis 2 Hyperolius balfouri balfouri quartet support < 0.9; multilocus bootstrap > 70% Hyperolius balfouri viridistriatus Hyperolius veithi quartet support > 0.9; multilocus bootstrap < 70% Hyperolius cinnamomeoventris 2 Hyperolius cinnamomeoventris 1 quartet support < 0.9; multilocus bootstrap < 70% Hyperolius olivaceus Hyperolius thomensis Hyperolius molleri Phrynomantis microps Hemisus sudanensis Hemisus marmoratus Balebreviceps hillmani Breviceps mossambicus Breviceps adspersus Breviceps montanus Breviceps fuscus Breviceps gibbosus Breviceps macrops Breviceps namaquensis Breviceps branchi Spelaeophryne methneri Probreviceps loveridgei Probreviceps uluguruensis Probreviceps durirostris Probreviceps sp. Nguru Probreviceps sp. Rubeho Probreviceps sp. Kigogo Probreviceps sp. Udzungwa Probreviceps rungwensis Probreviceps macrodactylus Callulina shengena Callulina laphami Callulina dawida Callulina kanga Callulina sp lowland Callulina sp Rubeho Callulina hanseni Callulina meteora Callulina stanleyi Callulina kisiwamsitu Callulina kreffti Nyctibates corrugatus Scotobleps gabonicus Astylosternus laticephalus Astylosternus occidentalis Trichobatrachus robustus Astylosternus diadematus Astylosternus schioetzi Astylosternus batesi Leptodactylodon mertensi Leptodactylodon erythrogaster Leptodactylodon perreti Leptodactylodon axillaris Leptodactylodon polyacanthus Leptodactylodon bicolor Leptodactylodon bueanus Leptodactylodon ornatus Leptodactylodon boulengeri Leptodactylodon ventrimarmoratus Leptodactylodon ovatus Leptopelis parkeri Leptopelis macrotis Leptopelis millsoni Leptopelis rufus Leptopelis argenteus Leptopelis yaldeni Leptopelis vannutellii Leptopelis susanae Leptopelis gramineus Leptopelis kivuensis Leptopelis ocellatus Leptopelis spiritusnoctis Leptopelis viridis Leptopelis aubryi Leptopelis natalensis Leptopelis palmatus Leptopelis calcaratus Leptopelis brevirostris Leptopelis notatus Leptopelis modestus Leptopelis flavomaculatus Leptopelis nordequatorialis Leptopelis bocagii Leptopelis vermiculatus Leptopelis aubryioides Leptopelis boulengeri Leptopelis fiziensis Leptopelis karissimbensis Cardioglossa melanogaster Cardioglossa gracilis Cardioglossa elegans Cardioglossa leucomystax Cardioglossa occidentalis Cardioglossa pulchra Cardioglossa oreas Cardioglossa manengouba Cardioglossa escalerae Cardioglossa trifasciata Cardioglossa annulata Cardioglossa nigromaculata Arthroleptis schubotzi Arthroleptis xenodactyloides Arthroleptis fichika Arthroleptis sylvaticus Arthroleptis taeniatus Arthroleptis reichei Arthroleptis affinis Arthroleptis nikeae Arthroleptis tanneri Arthroleptis francei Arthroleptis stenodactylus Arthroleptis perreti Arthroleptis palava Arthroleptis variabilis Arthroleptis adelphus Arthroleptis brevipes Arthroleptis bioko Arthroleptis poecilonotus Acanthixalus spinosus Acanthixalus sonjae Semnodactylus wealii Paracassina obscura Paracassina kounhiensis Phlyctimantis maculata Phlyctimantis keithae Phlyctimantis verrucosus Phlyctimantis boulengeri Phlyctimantis leonardi Kassina kuvangensis Kassina fusca Kassina lamottei Kassina schioetzi Kassina cassinoides Kassina senegalensis Kassina decorata Kassina maculosa Kassina arboricola Kassina cochranae Opisthothylax immaculatus Afrixalus enseticola Tachycnemis seychellensis Heterixalus madagascariensis Heterixalus boettgeri Heterixalus alboguttatus Heterixalus punctatus Heterixalus andrakata Heterixalus tricolor Heterixalus variabilis Heterixalus luteostriatus Heterixalus betsileo Heterixalus carbonei Heterixalus rutenbergi Afrixalus vibekensis Afrixalus weidholzi Afrixalus lacustris Afrixalus dorsimaculatus 1 Afrixalus morerei Afrixalus dorsimaculatus 2 Afrixalus knysae Afrixalus spinifrons Afrixalus delicatus Afrixalus brachycnemis Afrixalus stuhlmanni Afrixalus sylvaticus Afrixalus laevis Afrixalus lacteus Afrixalus fornasini Afrixalus equatorialis Afrixalus wittei Afrixalus osorioi Afrixalus paradorsalis paradorsalis 2 Afrixalus paradorsalis paradorsalis 1 Afrixalus paradorsalis manengubensis Afrixalus nigeriensis Afrixalus vittiger 1 Afrixalus vittiger 2 Afrixalus dorsalis 3 Afrixalus dorsalis 1 Afrixalus quadrivittatus 1 Afrixalus quadrivittatus 2 Afrixalus dorsalis 2 Afrixalus fulvovittatus 2 Afrixalus fulvovittatus 1 Morerella cyanophthalma Cryptothylax greshoffii Hyperolius semidiscus Hyperolius lupiroensis Hyperolius parkeri Hyperolius viridis Hyperolius pusillus Hyperolius benguellensis Hyperolius inyangae Hyperolius nasutus Hyperolius dartevellei Hyperolius rwandae Hyperolius friedemanni Hyperolius howelli Hyperolius acuticeps Hyperolius poweri Hyperolius igbettensis Hyperolius jacobseni Hyperolius adspersus Hyperolius pseudoargus Hyperolius fusciventris ssp. Hyperolius fusciventris burtoni Hyperolius bolifambae Hyperolius ocellatus ocellatus Hyperolius ocellatus purpurescens Hyperolius sp. nov. ‘Kupe’ Hyperolius camerunensis Hyperolius riggenbachi Hyperolius guttulatus 2 Hyperolius guttulatus 1 Hyperolius phantasticus Hyperolius pardalis Hyperolius steindachneri Hyperolius argus Hyperolius bocagei Hyperolius lamottei Hyperolius tuberculatus Hyperolius dintelmanni Hyperolius parallelus parallelus Hyperolius parallelus marginatus Hyperolius parallelus alborufus Hyperolius parallelus pyrrhodictyon Hyperolius marmoratus taeniatus Hyperolius marmoratus marmoratus Hyperolius marmoratus swynnertoni Hyperolius mariae Hyperolius reesi Hyperolius nitidulus 1 Hyperolius spatzi Hyperolius nitidulus 2 Hyperolius viridiflavus pitmani Hyperolius viridiflavus spp. Hyperolius viridiflavus destefanii Hyperolius viridiflavus variablilis Hyperolius viridiflavus ngorongoriensis Hyperolius viridiflavus ommatostictus Hyperolius viridiflavus pantherinus Hyperolius viridiflavus glandicolor Hyperolius koehleri Hyperolius tuberilinguis Hyperolius mosaicus Hyperolius endjami 2 Hyperolius endjami 1 Hyperolius cystocandicans Hyperolius jackie Hyperolius discodactylus Hyperolius lateralis Hyperolius castaneus Hyperolius frontalis Hyperolius platyceps Hyperolius cinereus Hyperolius raymondi Hyperolius langi Hyperolius kuligae Hyperolius ademetzi Hyperolius tanneri Hyperolius minitissimus Hyperolius burgessi Hyperolius spinigularis Hyperolius torrentis Hyperolius chlorosteus Hyperolius occidentalis Hyperolius sylvaticus Hyperolius baumanni Hyperolius picturatus Hyperolius zonatus Hyperolius bobirensis Hyperolius concolor concolor Hyperolius concolor guttatus Hyperolius concolor ibadanensis Hyperolius montanus Hyperolius pictus Hyperolius substriatus Hyperolius stictus Hyperolius mitchelli Hyperolius rubrovermiculatus Hyperolius quinquevittatus Hyperolius kivuensis 1 Hyperolius kivuensis 3 Hyperolius kivuensis 2 Hyperolius balfouri balfouri Hyperolius balfouri viridistriatus Hyperolius veithi Hyperolius cinnamomeoventris 2 Hyperolius cinnamomeoventris 1 Hyperolius olivaceus Hyperolius thomensis Hyperolius molleri

80 70 60 50 40 30 20 10 0 Cretaceous Paleocene Eocene Oligocene Miocene Plio-Pleistocene Monochromatic Root Prior Monochromatic Root Prior Monochromatic Root Prior Monochromatic Root Prior Relaxed Local Clock Strict Clock Relaxed Local Clock Strict Clock Mk1 Model Mk1 Model Mk2 Model Mk2 Model

FIXED FIXED FIXED FIXED Phrynomantis_microps Phrynomantis_microps Phrynomantis_microps Phrynomantis_microps Hemisus_sudanensis Hemisus_sudanensis Hemisus_sudanensis Hemisus_sudanensis Hemisus_marmoratus Hemisus_marmoratus Hemisus_marmoratus Hemisus_marmoratus Balebreviceps_hillmani Balebreviceps_hillmani Balebreviceps_hillmani Balebreviceps_hillmani Breviceps_mossambicus Breviceps_mossambicus Breviceps_mossambicus Breviceps_mossambicus Breviceps_adspersus Breviceps_adspersus Breviceps_adspersus Breviceps_adspersus Breviceps_montanus Breviceps_montanus Breviceps_montanus Breviceps_montanus Breviceps_fuscus Breviceps_fuscus Breviceps_fuscus Breviceps_fuscus Breviceps_gibbosus Breviceps_gibbosus Breviceps_gibbosus Breviceps_gibbosus Breviceps_macrops Breviceps_macrops Breviceps_macrops Breviceps_macrops Breviceps_namaquensis Breviceps_namaquensis Breviceps_namaquensis Breviceps_namaquensis Breviceps_branchi Breviceps_branchi Breviceps_branchi Breviceps_branchi Spelaeophryne_methneri Spelaeophryne_methneri Spelaeophryne_methneri Spelaeophryne_methneri Probreviceps_loveridgei Probreviceps_loveridgei Probreviceps_loveridgei Probreviceps_loveridgei Probreviceps_uluguruensis Probreviceps_uluguruensis Probreviceps_uluguruensis Probreviceps_uluguruensis Probreviceps_durirostris Probreviceps_durirostris Probreviceps_durirostris Probreviceps_durirostris Probreviceps_sp_Nguru Probreviceps_sp_Nguru Probreviceps_sp_Nguru Probreviceps_sp_Nguru Probreviceps_sp_Rubeho Probreviceps_sp_Rubeho Probreviceps_sp_Rubeho Probreviceps_sp_Rubeho Probreviceps_sp_Kigogo Probreviceps_sp_Kigogo Probreviceps_sp_Kigogo Probreviceps_sp_Kigogo Probreviceps_sp_Udzungwa Probreviceps_sp_Udzungwa Probreviceps_sp_Udzungwa Probreviceps_sp_Udzungwa Probreviceps_rungwensis Probreviceps_rungwensis Probreviceps_rungwensis Probreviceps_rungwensis Probreviceps_macrodactylus Probreviceps_macrodactylus Probreviceps_macrodactylus Probreviceps_macrodactylus Callulina_shengena Callulina_shengena Callulina_shengena Callulina_shengena Callulina_laphami Callulina_laphami Callulina_laphami Callulina_laphami Callulina_dawida Callulina_dawida Callulina_dawida Callulina_dawida Callulina_kanga Callulina_kanga Callulina_kanga Callulina_kanga Callulina_sp_lowland Callulina_sp_lowland Callulina_sp_lowland Callulina_sp_lowland Callulina_sp_Rubeho Callulina_sp_Rubeho Callulina_sp_Rubeho Callulina_sp_Rubeho Callulina_hanseni Callulina_hanseni Callulina_hanseni Callulina_hanseni Callulina_meteora Callulina_meteora Callulina_meteora Callulina_meteora Callulina_stanleyi Callulina_stanleyi Callulina_stanleyi Callulina_stanleyi Callulina_kisiwamsitu Callulina_kisiwamsitu Callulina_kisiwamsitu Callulina_kisiwamsitu Callulina_kreffti Callulina_kreffti Callulina_kreffti Callulina_kreffti Nyctibates_corrugatus Nyctibates_corrugatus Nyctibates_corrugatus Nyctibates_corrugatus Scotobleps_gabonicus Scotobleps_gabonicus Scotobleps_gabonicus Scotobleps_gabonicus Astylosternus_laticephalus Astylosternus_laticephalus Astylosternus_laticephalus Astylosternus_laticephalus Astylosternus_occidentalis Astylosternus_occidentalis Astylosternus_occidentalis Astylosternus_occidentalis Trichobatrachus_robustus Trichobatrachus_robustus Trichobatrachus_robustus Trichobatrachus_robustus Astylosternus_diadematus Astylosternus_diadematus Astylosternus_diadematus Astylosternus_diadematus Astylosternus_schioetzi Astylosternus_schioetzi Astylosternus_schioetzi Astylosternus_schioetzi Astylosternus_batesi Astylosternus_batesi Astylosternus_batesi Astylosternus_batesi Leptodactylodon_mertensi Leptodactylodon_mertensi Leptodactylodon_mertensi Leptodactylodon_mertensi Leptodactylodon_erythrogaster Leptodactylodon_erythrogaster Leptodactylodon_erythrogaster Leptodactylodon_erythrogaster Leptodactylodon_perreti Leptodactylodon_perreti Leptodactylodon_perreti Leptodactylodon_perreti Leptodactylodon_axillaris Leptodactylodon_axillaris Leptodactylodon_axillaris Leptodactylodon_axillaris Arthroleptidae Leptodactylodon_polyacanthus Arthroleptidae Leptodactylodon_polyacanthus Arthroleptidae Leptodactylodon_polyacanthus Arthroleptidae Leptodactylodon_polyacanthus Leptodactylodon_bicolor Leptodactylodon_bicolor Leptodactylodon_bicolor Leptodactylodon_bicolor Leptodactylodon_bueanus Leptodactylodon_bueanus Leptodactylodon_bueanus Leptodactylodon_bueanus Leptodactylodon_ornatus Leptodactylodon_ornatus Leptodactylodon_ornatus Leptodactylodon_ornatus Leptodactylodon_boulengeri Leptodactylodon_boulengeri Leptodactylodon_boulengeri Leptodactylodon_boulengeri Leptodactylodon_ventrimarmoratus Leptodactylodon_ventrimarmoratus Leptodactylodon_ventrimarmoratus Leptodactylodon_ventrimarmoratus Leptodactylodon_ovatus Leptodactylodon_ovatus Leptodactylodon_ovatus Leptodactylodon_ovatus Leptopelis_parkeri Leptopelis_parkeri Leptopelis_parkeri Leptopelis_parkeri Leptopelis_macrotis Leptopelis_macrotis Leptopelis_macrotis Leptopelis_macrotis Leptopelis_millsoni Leptopelis_millsoni Leptopelis_millsoni Leptopelis_millsoni Leptopelis_rufus Leptopelis_rufus Leptopelis_rufus Leptopelis_rufus Leptopelis_argenteus Leptopelis_argenteus Leptopelis_argenteus Leptopelis_argenteus Leptopelis_yaldeni Leptopelis_yaldeni Leptopelis_yaldeni Leptopelis_yaldeni Leptopelis_vannutellii Leptopelis_vannutellii Leptopelis_vannutellii Leptopelis_vannutellii Leptopelis_susanae Leptopelis_susanae Leptopelis_susanae Leptopelis_susanae Leptopelis_gramineus Leptopelis_gramineus Leptopelis_gramineus Leptopelis_gramineus Leptopelis_kivuensis Leptopelis_kivuensis Leptopelis_kivuensis Leptopelis_kivuensis Leptopelis_ocellatus Leptopelis_ocellatus Leptopelis_ocellatus Leptopelis_ocellatus Leptopelis_spiritusnoctis Leptopelis_spiritusnoctis Leptopelis_spiritusnoctis Leptopelis_spiritusnoctis Leptopelis_viridis Leptopelis_viridis Leptopelis_viridis Leptopelis_viridis Leptopelis_aubryi Leptopelis_aubryi Leptopelis_aubryi Leptopelis_aubryi Leptopelis_natalensis Leptopelis_natalensis Leptopelis_natalensis Leptopelis_natalensis Leptopelis_palmatus Leptopelis_palmatus Leptopelis_palmatus Leptopelis_palmatus Leptopelis_calcaratus Leptopelis_calcaratus Leptopelis_calcaratus Leptopelis_calcaratus Leptopelis_brevirostris Leptopelis_brevirostris Leptopelis_brevirostris Leptopelis_brevirostris Leptopelis_notatus Leptopelis_notatus Leptopelis_notatus Leptopelis_notatus Leptopelis_modestus Leptopelis_modestus Leptopelis_modestus Leptopelis_modestus Leptopelis_flavomaculatus Leptopelis_flavomaculatus Leptopelis_flavomaculatus Leptopelis_flavomaculatus Leptopelis_nordequatorialis Leptopelis_nordequatorialis Leptopelis_nordequatorialis Leptopelis_nordequatorialis Leptopelis_bocagii Leptopelis_bocagii Leptopelis_bocagii Leptopelis_bocagii Leptopelis_vermiculatus Leptopelis_vermiculatus Leptopelis_vermiculatus Leptopelis_vermiculatus Leptopelis_aubryioides Leptopelis_aubryioides Leptopelis_aubryioides Leptopelis_aubryioides Leptopelis_boulengeri Leptopelis_boulengeri Leptopelis_boulengeri Leptopelis_boulengeri Leptopelis_fiziensis Leptopelis_fiziensis Leptopelis_fiziensis Leptopelis_fiziensis Leptopelis_karissimbensis Leptopelis_karissimbensis Leptopelis_karissimbensis Leptopelis_karissimbensis Cardioglossa_melanogaster Cardioglossa_melanogaster Cardioglossa_melanogaster Cardioglossa_melanogaster Cardioglossa_gracilis Cardioglossa_gracilis Cardioglossa_gracilis Cardioglossa_gracilis Cardioglossa_elegans Cardioglossa_elegans Cardioglossa_elegans Cardioglossa_elegans Cardioglossa_leucomystax Cardioglossa_leucomystax Cardioglossa_leucomystax Cardioglossa_leucomystax Cardioglossa_occidentalis Cardioglossa_occidentalis Cardioglossa_occidentalis Cardioglossa_occidentalis Cardioglossa_pulchra Cardioglossa_pulchra Cardioglossa_pulchra Cardioglossa_pulchra Cardioglossa_oreas Cardioglossa_oreas Cardioglossa_oreas Cardioglossa_oreas Cardioglossa_manengouba Cardioglossa_manengouba Cardioglossa_manengouba Cardioglossa_manengouba Cardioglossa_escalerae Cardioglossa_escalerae Cardioglossa_escalerae Cardioglossa_escalerae Cardioglossa_trifasciata Cardioglossa_trifasciata Cardioglossa_trifasciata Cardioglossa_trifasciata Cardioglossa_annulata Cardioglossa_annulata Cardioglossa_annulata Cardioglossa_annulata Cardioglossa_nigromaculata Cardioglossa_nigromaculata Cardioglossa_nigromaculata Cardioglossa_nigromaculata Arthroleptis_schubotzi Arthroleptis_schubotzi Arthroleptis_schubotzi Arthroleptis_schubotzi Arthroleptis_xenodactyloides Arthroleptis_xenodactyloides Arthroleptis_xenodactyloides Arthroleptis_xenodactyloides Arthroleptis_fichika Arthroleptis_fichika Arthroleptis_fichika Arthroleptis_fichika Arthroleptis_sylvaticus Arthroleptis_sylvaticus Arthroleptis_sylvaticus Arthroleptis_sylvaticus Arthroleptis_taeniatus Arthroleptis_taeniatus Arthroleptis_taeniatus Arthroleptis_taeniatus Arthroleptis_reichei Arthroleptis_reichei Arthroleptis_reichei Arthroleptis_reichei Arthroleptis_affinis Arthroleptis_affinis Arthroleptis_affinis Arthroleptis_affinis Arthroleptis_nikeae Arthroleptis_nikeae Arthroleptis_nikeae Arthroleptis_nikeae Arthroleptis_tanneri Arthroleptis_tanneri Arthroleptis_tanneri Arthroleptis_tanneri Arthroleptis_francei Arthroleptis_francei Arthroleptis_francei Arthroleptis_francei Arthroleptis_stenodactylus Arthroleptis_stenodactylus Arthroleptis_stenodactylus Arthroleptis_stenodactylus Arthroleptis_perreti Arthroleptis_perreti Arthroleptis_perreti Arthroleptis_perreti Arthroleptis_palava Arthroleptis_palava Arthroleptis_palava Arthroleptis_palava Arthroleptis_variabilis Arthroleptis_variabilis Arthroleptis_variabilis Arthroleptis_variabilis Arthroleptis_adelphus Arthroleptis_adelphus Arthroleptis_adelphus Arthroleptis_adelphus Arthroleptis_brevipes Arthroleptis_brevipes Arthroleptis_brevipes Arthroleptis_brevipes Arthroleptis_bioko Arthroleptis_bioko Arthroleptis_bioko Arthroleptis_bioko Arthroleptis_poecilonotus Arthroleptis_poecilonotus Arthroleptis_poecilonotus Arthroleptis_poecilonotus Acanthixalus_spinosus Acanthixalus_spinosus Acanthixalus_spinosus Acanthixalus_spinosus Acanthixalus_sonjae Acanthixalus_sonjae Acanthixalus_sonjae Acanthixalus_sonjae Semnodactylus_wealii Semnodactylus_wealii Semnodactylus_wealii Semnodactylus_wealii Paracassina_obscura Paracassina_obscura Paracassina_obscura Paracassina_obscura Paracassina_kounhiensis Paracassina_kounhiensis Paracassina_kounhiensis Paracassina_kounhiensis Phlyctimantis_maculata Phlyctimantis_maculata Phlyctimantis_maculata Phlyctimantis_maculata Phlyctimantis_keithae Phlyctimantis_keithae Phlyctimantis_keithae Phlyctimantis_keithae Phlyctimantis_verrucosus Phlyctimantis_verrucosus Phlyctimantis_verrucosus Phlyctimantis_verrucosus Phlyctimantis_boulengeri Phlyctimantis_boulengeri Phlyctimantis_boulengeri Phlyctimantis_boulengeri Phlyctimantis_leonardi Phlyctimantis_leonardi Phlyctimantis_leonardi Phlyctimantis_leonardi Kassina_kuvangensis Kassina_kuvangensis Kassina_kuvangensis Kassina_kuvangensis Kassina_fusca Kassina_fusca Kassina_fusca Kassina_fusca Kassina_lamottei Kassina_lamottei Kassina_lamottei Kassina_lamottei Kassina_schioetzi Kassina_schioetzi Kassina_schioetzi Kassina_schioetzi Kassina_cassinoides Kassina_cassinoides Kassina_cassinoides Kassina_cassinoides Kassina_senegalensis Kassina_senegalensis Kassina_senegalensis Kassina_senegalensis Kassina_decorata Kassina_decorata Kassina_decorata Kassina_decorata Kassina_maculosa Kassina_maculosa Kassina_maculosa Kassina_maculosa Kassina_arboricola Kassina_arboricola Kassina_arboricola Kassina_arboricola Kassina_cochranae Kassina_cochranae Kassina_cochranae Kassina_cochranae Opisthothylax_immaculatus Opisthothylax_immaculatus Opisthothylax_immaculatus Opisthothylax_immaculatus Afrixalus_enseticola Afrixalus_enseticola Afrixalus_enseticola Afrixalus_enseticola Hyperoliidae Tachycnemis_seychellensis Hyperoliidae Tachycnemis_seychellensis Hyperoliidae Tachycnemis_seychellensis Hyperoliidae Tachycnemis_seychellensis Heterixalus_madagascariensis Heterixalus_madagascariensis Heterixalus_madagascariensis Heterixalus_madagascariensis Heterixalus_boettgeri Heterixalus_boettgeri Heterixalus_boettgeri Heterixalus_boettgeri Heterixalus_alboguttatus Heterixalus_alboguttatus Heterixalus_alboguttatus Heterixalus_alboguttatus Heterixalus_punctatus Heterixalus_punctatus Heterixalus_punctatus Heterixalus_punctatus Heterixalus_andrakata Heterixalus_andrakata Heterixalus_andrakata Heterixalus_andrakata Heterixalus_tricolor Heterixalus_tricolor Heterixalus_tricolor Heterixalus_tricolor Heterixalus_variabilis Heterixalus_variabilis Heterixalus_variabilis Heterixalus_variabilis Heterixalus_luteostriatus Heterixalus_luteostriatus Heterixalus_luteostriatus Heterixalus_luteostriatus Heterixalus_betsileo Heterixalus_betsileo Heterixalus_betsileo Heterixalus_betsileo Heterixalus_carbonei Heterixalus_carbonei Heterixalus_carbonei Heterixalus_carbonei Heterixalus_rutenbergi Heterixalus_rutenbergi Heterixalus_rutenbergi Heterixalus_rutenbergi Afrixalus_vibekensis Afrixalus_vibekensis Afrixalus_vibekensis Afrixalus_vibekensis Afrixalus_weidholzi Afrixalus_weidholzi Afrixalus_weidholzi Afrixalus_weidholzi Afrixalus_lacustris Afrixalus_lacustris Afrixalus_lacustris Afrixalus_lacustris Afrixalus_dorsimaculatus_1 Afrixalus_dorsimaculatus_1 Afrixalus_dorsimaculatus_1 Afrixalus_dorsimaculatus_1 Afrixalus_morerei Afrixalus_morerei Afrixalus_morerei Afrixalus_morerei Afrixalus_dorsimaculatus_2 Afrixalus_dorsimaculatus_2 Afrixalus_dorsimaculatus_2 Afrixalus_dorsimaculatus_2 Afrixalus_knysae Afrixalus_knysae Afrixalus_knysae Afrixalus_knysae Afrixalus_spinifrons Afrixalus_spinifrons Afrixalus_spinifrons Afrixalus_spinifrons Afrixalus_delicatus Afrixalus_delicatus Afrixalus_delicatus Afrixalus_delicatus Afrixalus_brachycnemis Afrixalus_brachycnemis Afrixalus_brachycnemis Afrixalus_brachycnemis Afrixalus_stuhlmanni Afrixalus_stuhlmanni Afrixalus_stuhlmanni Afrixalus_stuhlmanni Afrixalus_sylvaticus Afrixalus_sylvaticus Afrixalus_sylvaticus Afrixalus_sylvaticus Afrixalus_laevis Afrixalus_laevis Afrixalus_laevis Afrixalus_laevis Afrixalus_lacteus Afrixalus_lacteus Afrixalus_lacteus Afrixalus_lacteus Afrixalus_fornasini Afrixalus_fornasini Afrixalus_fornasini Afrixalus_fornasini Afrixalus_equatorialis Afrixalus_equatorialis Afrixalus_equatorialis Afrixalus_equatorialis Afrixalus_wittei Afrixalus_wittei Afrixalus_wittei Afrixalus_wittei Afrixalus_osorioi Afrixalus_osorioi Afrixalus_osorioi Afrixalus_osorioi Afrixalus_paradorsalis_paradorsalis_2 Afrixalus_paradorsalis_paradorsalis_2 Afrixalus_paradorsalis_paradorsalis_2 Afrixalus_paradorsalis_paradorsalis_2 Afrixalus_paradorsalis_paradorsalis_1 Afrixalus_paradorsalis_paradorsalis_1 Afrixalus_paradorsalis_paradorsalis_1 Afrixalus_paradorsalis_paradorsalis_1 Afrixalus_paradorsalis_manengubensis Afrixalus_paradorsalis_manengubensis Afrixalus_paradorsalis_manengubensis Afrixalus_paradorsalis_manengubensis Afrixalus_nigeriensis Afrixalus_nigeriensis Afrixalus_nigeriensis Afrixalus_nigeriensis Afrixalus_vittiger_1 Afrixalus_vittiger_1 Afrixalus_vittiger_1 Afrixalus_vittiger_1 Afrixalus_vittiger_2 Afrixalus_vittiger_2 Afrixalus_vittiger_2 Afrixalus_vittiger_2 Afrixalus_dorsalis_3 Afrixalus_dorsalis_3 Afrixalus_dorsalis_3 Afrixalus_dorsalis_3 Afrixalus_dorsalis_1 Afrixalus_dorsalis_1 Afrixalus_dorsalis_1 Afrixalus_dorsalis_1 Afrixalus_quadrivittatus_1 Afrixalus_quadrivittatus_1 Afrixalus_quadrivittatus_1 Afrixalus_quadrivittatus_1 Afrixalus_quadrivittatus_2 Afrixalus_quadrivittatus_2 Afrixalus_quadrivittatus_2 Afrixalus_quadrivittatus_2 Afrixalus_dorsalis_2 Afrixalus_dorsalis_2 Afrixalus_dorsalis_2 Afrixalus_dorsalis_2 Afrixalus_fulvovittatus_2 Afrixalus_fulvovittatus_2 Afrixalus_fulvovittatus_2 Afrixalus_fulvovittatus_2 Afrixalus_fulvovittatus_1 Afrixalus_fulvovittatus_1 Afrixalus_fulvovittatus_1 Afrixalus_fulvovittatus_1 Morerella_cyanophthalma Morerella_cyanophthalma Morerella_cyanophthalma Morerella_cyanophthalma Cryptothylax_greshoffii Cryptothylax_greshoffii Cryptothylax_greshoffii Cryptothylax_greshoffii Hyperolius_semidiscus Hyperolius_semidiscus Hyperolius_semidiscus Hyperolius_semidiscus Hyperolius_lupiroensis Hyperolius_lupiroensis Hyperolius_lupiroensis Hyperolius_lupiroensis Hyperolius_parkeri Hyperolius_parkeri Hyperolius_parkeri Hyperolius_parkeri Hyperolius_viridis Hyperolius_viridis Hyperolius_viridis Hyperolius_viridis Hyperolius_pusillus Hyperolius_pusillus Hyperolius_pusillus Hyperolius_pusillus Hyperolius_benguellensis Hyperolius_benguellensis Hyperolius_benguellensis Hyperolius_benguellensis Hyperolius_inyangae Hyperolius_inyangae Hyperolius_inyangae Hyperolius_inyangae Hyperolius_nasutus Hyperolius_nasutus Hyperolius_nasutus Hyperolius_nasutus Hyperolius_dartevellei Hyperolius_dartevellei Hyperolius_dartevellei Hyperolius_dartevellei Hyperolius_rwandae Hyperolius_rwandae Hyperolius_rwandae Hyperolius_rwandae Hyperolius_friedemanni Hyperolius_friedemanni Hyperolius_friedemanni Hyperolius_friedemanni Hyperolius_howelli Hyperolius_howelli Hyperolius_howelli Hyperolius_howelli Hyperolius_acuticeps Hyperolius_acuticeps Hyperolius_acuticeps Hyperolius_acuticeps Hyperolius_poweri Hyperolius_poweri Hyperolius_poweri Hyperolius_poweri Hyperolius_igbettensis Hyperolius_igbettensis Hyperolius_igbettensis Hyperolius_igbettensis Hyperolius_jacobseni Hyperolius_jacobseni Hyperolius_jacobseni Hyperolius_jacobseni Hyperolius_adspersus Hyperolius_adspersus Hyperolius_adspersus Hyperolius_adspersus Hyperolius_pseudoargus Hyperolius_pseudoargus Hyperolius_pseudoargus Hyperolius_pseudoargus Hyperolius_fusciventris_burtoni Hyperolius_fusciventris_burtoni Hyperolius_fusciventris_burtoni Hyperolius_fusciventris_burtoni Hyperolius_fusciventris_ssp Hyperolius_fusciventris_ssp Hyperolius_fusciventris_ssp Hyperolius_fusciventris_ssp Hyperolius_bolifambae Hyperolius_bolifambae Hyperolius_bolifambae Hyperolius_bolifambae Hyperolius_ocellatus_ocellatus Hyperolius_ocellatus_ocellatus Hyperolius_ocellatus_ocellatus Hyperolius_ocellatus_ocellatus Hyperolius_ocellatus_purpurescens Hyperolius_ocellatus_purpurescens Hyperolius_ocellatus_purpurescens Hyperolius_ocellatus_purpurescens Hyperolius_sp_nov Hyperolius_sp_nov Hyperolius_sp_nov Hyperolius_sp_nov Hyperolius_camerunensis Hyperolius_camerunensis Hyperolius_camerunensis Hyperolius_camerunensis Hyperolius_riggenbachi Hyperolius_riggenbachi Hyperolius_riggenbachi Hyperolius_riggenbachi Hyperolius_guttulatus_2 Hyperolius_guttulatus_2 Hyperolius_guttulatus_2 Hyperolius_guttulatus_2 Hyperolius_guttulatus_1 Hyperolius_guttulatus_1 Hyperolius_guttulatus_1 Hyperolius_guttulatus_1 Hyperolius_phantasticus Hyperolius_phantasticus Hyperolius_phantasticus Hyperolius_phantasticus Hyperolius_pardalis Hyperolius_pardalis Hyperolius_pardalis Hyperolius_pardalis Hyperolius_steindachneri Hyperolius_steindachneri Hyperolius_steindachneri Hyperolius_steindachneri Hyperolius_argus Hyperolius_argus Hyperolius_argus Hyperolius_argus Hyperolius_bocagei Hyperolius_bocagei Hyperolius_bocagei Hyperolius_bocagei Hyperolius_lamottei Hyperolius_lamottei Hyperolius_lamottei Hyperolius_lamottei Hyperolius_tuberculatus Hyperolius_tuberculatus Hyperolius_tuberculatus Hyperolius_tuberculatus Hyperolius_dintelmanni Hyperolius_dintelmanni Hyperolius_dintelmanni Hyperolius_dintelmanni Hyperolius_parallelus_parallelus Hyperolius_parallelus_parallelus Hyperolius_parallelus_parallelus Hyperolius_parallelus_parallelus Hyperolius_parallelus_marginatus Hyperolius_parallelus_marginatus Hyperolius_parallelus_marginatus Hyperolius_parallelus_marginatus Hyperolius_parallelus_alborufus Hyperolius_parallelus_alborufus Hyperolius_parallelus_alborufus Hyperolius_parallelus_alborufus Hyperolius_parallelus_pyrrhodictyon Hyperolius_parallelus_pyrrhodictyon Hyperolius_parallelus_pyrrhodictyon Hyperolius_parallelus_pyrrhodictyon Hyperolius_marmoratus_taeniatus Hyperolius_marmoratus_taeniatus Hyperolius_marmoratus_taeniatus Hyperolius_marmoratus_taeniatus Hyperolius_marmoratus_marmoratus Hyperolius_marmoratus_marmoratus Hyperolius_marmoratus_marmoratus Hyperolius_marmoratus_marmoratus Hyperolius_marmoratus_swynnertoni Hyperolius_marmoratus_swynnertoni Hyperolius_marmoratus_swynnertoni Hyperolius_marmoratus_swynnertoni Hyperolius_mariae Hyperolius_mariae Hyperolius_mariae Hyperolius_mariae Hyperolius_reesi Hyperolius_reesi Hyperolius_reesi Hyperolius_reesi Hyperolius_nitidulus_1 Hyperolius_nitidulus_1 Hyperolius_nitidulus_1 Hyperolius_nitidulus_1 Hyperolius_spatzi Hyperolius_spatzi Hyperolius_spatzi Hyperolius_spatzi Hyperolius_nitidulus_2 Hyperolius_nitidulus_2 Hyperolius_nitidulus_2 Hyperolius_nitidulus_2 Hyperolius_viridiflavus_pitmani Hyperolius_viridiflavus_pitmani Hyperolius_viridiflavus_pitmani Hyperolius_viridiflavus_pitmani Hyperolius_viridiflavus_ssp Marginal Likelihood Estimate Hyperolius_viridiflavus_ssp Marginal Likelihood Estimate Hyperolius_viridiflavus_ssp Marginal Likelihood Estimate Hyperolius_viridiflavus_ssp Marginal Likelihood Estimate Hyperolius_viridiflavus_destefanii Hyperolius_viridiflavus_destefanii Hyperolius_viridiflavus_destefanii Hyperolius_viridiflavus_destefanii Hyperolius_viridiflavus_variablilis Hyperolius_viridiflavus_variablilis Hyperolius_viridiflavus_variablilis Hyperolius_viridiflavus_variablilis Hyperolius_viridiflavus_ngorongoriensis Hyperolius_viridiflavus_ngorongoriensis Hyperolius_viridiflavus_ngorongoriensis Hyperolius_viridiflavus_ngorongoriensis Hyperolius_viridiflavus_ommatostictus Hyperolius_viridiflavus_ommatostictus Hyperolius_viridiflavus_ommatostictus Hyperolius_viridiflavus_ommatostictus -85.10 Hyperolius_viridiflavus_pantherinus -92.12 Hyperolius_viridiflavus_pantherinus -81.73 Hyperolius_viridiflavus_pantherinus -81.83 Hyperolius_viridiflavus_pantherinus Hyperolius_viridiflavus_glandicolor Hyperolius_viridiflavus_glandicolor Hyperolius_viridiflavus_glandicolor Hyperolius_viridiflavus_glandicolor Hyperolius_koehleri Hyperolius_koehleri Hyperolius_koehleri Hyperolius_koehleri Hyperolius_tuberilinguis Hyperolius_tuberilinguis Hyperolius_tuberilinguis Hyperolius_tuberilinguis Hyperolius_mosaicus Hyperolius_mosaicus Hyperolius_mosaicus Hyperolius_mosaicus Hyperolius_endjami_2 Hyperolius_endjami_2 Hyperolius_endjami_2 Hyperolius_endjami_2 Hyperolius_endjami_1 Hyperolius_endjami_1 Hyperolius_endjami_1 Hyperolius_endjami_1 Hyperolius_cystocandicans Hyperolius_cystocandicans Hyperolius_cystocandicans Hyperolius_cystocandicans Hyperolius_jackie Hyperolius_jackie Hyperolius_jackie Hyperolius_jackie Hyperolius_discodactylus Hyperolius_discodactylus Hyperolius_discodactylus Hyperolius_discodactylus Hyperolius_lateralis Hyperolius_lateralis Hyperolius_lateralis Hyperolius_lateralis Hyperolius_castaneus Hyperolius_castaneus Hyperolius_castaneus Hyperolius_castaneus Estimated State Changes Hyperolius_frontalis Estimated State Changes Hyperolius_frontalis Estimated State Changes Hyperolius_frontalis Estimated State Changes Hyperolius_frontalis Hyperolius_platyceps Hyperolius_platyceps Hyperolius_platyceps Hyperolius_platyceps Hyperolius_cinereus Hyperolius_cinereus Hyperolius_cinereus Hyperolius_cinereus Hyperolius_raymondi Hyperolius_raymondi Hyperolius_raymondi Hyperolius_raymondi Hyperolius_langi 0 -> 1: 14.7 [9 - 18] Hyperolius_langi 0 -> 1: 3.0 [1 - 8] Hyperolius_langi 0 -> 1: 2.6 [1 - 6] Hyperolius_langi 0 -> 1: 16.5 [9 - 22] Hyperolius_kuligae Hyperolius_kuligae Hyperolius_kuligae Hyperolius_kuligae Hyperolius_ademetzi Hyperolius_ademetzi Hyperolius_ademetzi Hyperolius_ademetzi Hyperolius_tanneri Hyperolius_tanneri Hyperolius_tanneri Hyperolius_tanneri Hyperolius_minitissimus 1 -> 0: 7.2 [3 - 11] Hyperolius_minitissimus 1 -> 0: 25.6 [17 - 32] Hyperolius_minitissimus 1 -> 0: 26.2 [17 - 31] Hyperolius_minitissimus 1 -> 0: 8.9 [4 - 16] Hyperolius_burgessi Hyperolius_burgessi Hyperolius_burgessi Hyperolius_burgessi Hyperolius_spinigularis Hyperolius_spinigularis Hyperolius_spinigularis Hyperolius_spinigularis Hyperolius_torrentis Hyperolius_torrentis Hyperolius_torrentis Hyperolius_torrentis Hyperolius_chlorosteus Hyperolius_chlorosteus Hyperolius_chlorosteus Hyperolius_chlorosteus Hyperolius_occidentalis Hyperolius_occidentalis Hyperolius_occidentalis Hyperolius_occidentalis Hyperolius_sylvaticus Hyperolius_sylvaticus Hyperolius_sylvaticus Hyperolius_sylvaticus Hyperolius_baumanni Hyperolius_baumanni Hyperolius_baumanni Hyperolius_baumanni Hyperolius_picturatus Hyperolius_picturatus Hyperolius_picturatus Hyperolius_picturatus Hyperolius_zonatus Hyperolius_zonatus Hyperolius_zonatus Hyperolius_zonatus Hyperolius_bobirensis Hyperolius_bobirensis Hyperolius_bobirensis Hyperolius_bobirensis Hyperolius_concolor_concolor Hyperolius_concolor_concolor Hyperolius_concolor_concolor Hyperolius_concolor_concolor Hyperolius_concolor_guttatus Hyperolius_concolor_guttatus Hyperolius_concolor_guttatus Hyperolius_concolor_guttatus Hyperolius_concolor_ibadanensis Hyperolius_concolor_ibadanensis Hyperolius_concolor_ibadanensis Hyperolius_concolor_ibadanensis Hyperolius_montanus Hyperolius_montanus Hyperolius_montanus Hyperolius_montanus Posterior Hyperolius_pictus Posterior Hyperolius_pictus Posterior Hyperolius_pictus Posterior Hyperolius_pictus Hyperolius_substriatus Hyperolius_substriatus Hyperolius_substriatus Hyperolius_substriatus Probability Hyperolius_stictus Probability Hyperolius_stictus Probability Hyperolius_stictus Probability Hyperolius_stictus Monochromatic Hyperolius_mitchelli Monochromatic Hyperolius_mitchelli Monochromatic Hyperolius_mitchelli Monochromatic Hyperolius_mitchelli 1.0 Hyperolius_rubrovermiculatus 1.0 Hyperolius_rubrovermiculatus 1.0 Hyperolius_rubrovermiculatus 1.0 Hyperolius_rubrovermiculatus 0.9 Hyperolius_quinquevittatus 0.9 Hyperolius_quinquevittatus 0.9 Hyperolius_quinquevittatus 0.9 Hyperolius_quinquevittatus Hyperolius_kivuensis_1 Hyperolius_kivuensis_1 Hyperolius_kivuensis_1 Hyperolius_kivuensis_1 0.8 Hyperolius_kivuensis_3 0.8 Hyperolius_kivuensis_3 0.8 Hyperolius_kivuensis_3 0.8 Hyperolius_kivuensis_3 Dichromatic Hyperolius_kivuensis_2 Dichromatic Hyperolius_kivuensis_2 Dichromatic Hyperolius_kivuensis_2 Dichromatic Hyperolius_kivuensis_2 0.7 Hyperolius_balfouri_balfouri 0.7 Hyperolius_balfouri_balfouri 0.7 Hyperolius_balfouri_balfouri 0.7 Hyperolius_balfouri_balfouri 0.6 Hyperolius_balfouri_viridistriatus 0.6 Hyperolius_balfouri_viridistriatus 0.6 Hyperolius_balfouri_viridistriatus 0.6 Hyperolius_balfouri_viridistriatus Hyperolius_veithi Hyperolius_veithi Hyperolius_veithi Hyperolius_veithi Hyperolius_cinnamomeoventris_2 Hyperolius_cinnamomeoventris_2 Hyperolius_cinnamomeoventris_2 Hyperolius_cinnamomeoventris_2 Hyperolius_cinnamomeoventris_1 Hyperolius_cinnamomeoventris_1 Hyperolius_cinnamomeoventris_1 Hyperolius_cinnamomeoventris_1 Hyperolius_olivaceus Hyperolius_olivaceus Hyperolius_olivaceus Hyperolius_olivaceus Hyperolius_thomensis Hyperolius_thomensis Hyperolius_thomensis Hyperolius_thomensis Hyperolius_molleri Hyperolius_molleri Hyperolius_molleri Hyperolius_molleri

8 0 7 0 6 0 5 0 4 0 3 0 2 0 1 0 0 Ma 8 0 7 0 6 0 5 0 4 0 3 0 2 0 1 0 0 Ma 8 0 7 0 6 0 5 0 4 0 3 0 2 0 1 0 0 Ma 8 0 7 0 6 0 5 0 4 0 3 0 2 0 1 0 0 Ma L

p s Leptopelis_karissimbensis

Cardioglossa_melanogaster t u o Leptopelis_vermiculatus

Leptopelis_aubryioides t

a l

Leptopelis_boulengeri e

Cardioglossa_occidentalis i u

Cardioglossa_leucomystax s r

t L c s

Cardioglossa_manengouba u

Cardioglossa_gracilis t Cardioglossa_elegans _ e a Leptopelis_fiziensis

s o n p

r m

e i

o t o o

Cardioglossa_pulchra r v p Cardioglossa_nigromaculata d e

r a

Cardioglossa_escalerae e

l f m b l q

Arthroleptis_xenodactyloidesCardioglossa_trifasciata i s _ _ _ Cardioglossa_oreas u

_ s s

s a i i

i l l l b

Cardioglossa_annulata t e e o e

o p p r p c

i o o

a a o t t t

l g i p p Arthroleptis_schubotzi p

i e e

e L L L Leptopelis_notatus Leptopelis_calcaratus Leptopelis_palmatus Leptopelis_natalensis Arthroleptis_taeniatus Leptopelis_viridis Arthroleptis_fichika Leptopelis_aubryi Leptopelis_spiritusnoctis Arthroleptis_sylvaticus Leptopelis_ocellatus Leptopelis_kivuensis Leptopelis_susanae Leptopelis_gramineus Leptopelis_vannutellii Leptopelis_yaldeni Arthroleptis_affinis Leptopelis_argenteus Arthroleptis_reichei Leptopelis_rufus Leptopelis_macrotis Arthroleptis_nikeae Arthroleptis_stenodactylus Leptopelis_millsoni Arthroleptis_francei Leptopelis_parkeri Leptodactylodon_ventrimarmoratus Arthroleptis_tanneri Leptodactylodon_ovatus Leptodactylodon_boulengeri Leptodactylodon_ornatus Arthroleptis_perreti Leptodactylodon_bueanus Arthroleptis_variabilis Leptodactylodon_polyacanthus Leptodactylodon_bicolor Arthroleptis_adelphusArthroleptis_palava Leptodactylodon_perreti Leptodactylodon_axillaris Arthroleptis_poecilonotus Arthroleptis_brevipes Leptodactylodon_erythrogaster Leptodactylodon_mertensi Astylosternus_schioetzi Astylosternus_batesi Arthroleptis_bioko Acanthixalus_sonjae Astylosternus_diadematus Acanthixalus_spinosus Trichobatrachus_robustus Astylosternus_laticephalus Paracassina_kounhiensisSemnodactylus_wealii Astylosternus_occidentalis Scotobleps_gabonicus Paracassina_obscura Nyctibates_corrugatus Phlyctimantis_maculata Callulina_kisiwamsitu Phlyctimantis_leonardi Callulina_kreffti Phlyctimantis_boulengeri Callulina_stanleyi Phlyctimantis_verrucosus Callulina_hanseni Callulina_meteora Phlyctimantis_keithae Callulina_sp_Rubeho Kassina_kuvangensis Callulina_sp_lowland Callulina_kanga Callulina_dawida Kassina_fusca Kassina_lamottei Callulina_shengena Kassina_cassinoides Callulina_laphami Probreviceps_rungwensis Kassina_schioetzi Kassina_senegalensis Probreviceps_macrodactylus Probreviceps_sp_Udzungwa Kassina_cochranae Probreviceps_sp_Kigogo Kassina_arboricola Probreviceps_sp_Rubeho Kassina_maculosa Probreviceps_durirostris Probreviceps_sp_Nguru Opisthothylax_immaculatusKassina_decorata Probreviceps_loveridgei Afrixalus_enseticola Probreviceps_uluguruensis Tachycnemis_seychellensis Spelaeophryne_methneri Heterixalus_madagascariensis Breviceps_namaquensis Breviceps_branchi Heterixalus_alboguttatus Breviceps_macrops Heterixalus_boettgeri Breviceps_gibbosus Heterixalus_punctatus Breviceps_fuscus Heterixalus_andrakata Breviceps_montanus Heterixalus_variabilis Breviceps_mossambicus Breviceps_adspersus Heterixalus_tricolor Balebreviceps_hillmani Heterixalus_luteostriatus Hemisus_sudanensis Heterixalus_betsileo Hemisus_marmoratus Heterixalus_rutenbergi Phrynomantis_microps Heterixalus_carbonei Afrixalus_spinifrons Hyperolius_balfouri_1 Afrixalus_knysae Hyperolius_balfouri_2 Afrixalus_delicatus Hyperolius_kivuensis_2 Hyperolius_kivuensis_1 Afrixalus_brachycnemis Hyperolius_kivuensis_3 Afrixalus_sylvaticus Hyperolius_quinquevittatus Afrixalus_stuhlmanni Hyperolius_cinnamomeoventris_2 Afrixalus_vibekensis Hyperolius_cinnamomeoventris_1 Afrixalus_weidholzi Hyperolius_veithi Afrixalus_lacustris Hyperolius_thomensis Hyperolius_molleri Afrixalus_dorsimaculatus_1 Hyperolius_cinnamomeoventris_3 Hyperolius_mitchelli Afrixalus_dorsimaculatus_2Afrixalus_morerei Hyperolius_rubrovermiculatus Afrixalus_lacteus Hyperolius_sp_nov_2 Afrixalus_laevis Hyperolius_pictus Afrixalus_fornasini Hyperolius_substriatus Hyperolius_montanus Afrixalus_equatorialis Afrixalus_osorioi Hyperolius_concolor_1 Afrixalus_wittei Hyperolius_concolor_3 Hyperolius_concolor_2 Hyperolius_zonatus Afrixalus_paradorsalis_3 Hyperolius_bobirensis Afrixalus_paradorsalis_1 Hyperolius_baumanni Afrixalus_paradorsalis_2 Hyperolius_picturatus Afrixalus_nigeriensis Afrixalus_vittiger_2 Hyperolius_sylvaticus Hyperolius_chlorosteus Afrixalus_vittiger_1 Hyperolius_occidentalis Afrixalus_dorsalis_1 Hyperolius_torrentis Afrixalus_dorsalis_3 Hyperolius_burgessi Hyperolius_spinigularis Hyperolius_minitissimus Afrixalus_quadrivittatus_2 Hyperolius_tanneri Afrixalus_quadrivittatus_1Afrixalus_dorsalis_2 Hyperolius_cinereus Hyperolius_raymondi Hyperolius_platyceps Afrixalus_fulvovittatus_1 Hyperolius_kuligae Afrixalus_fulvovittatus_2 Hyperolius_ademetzi Cryptothylax_greshoffii Hyperolius_langi Hyperolius_discodactylus Morerella_cyanophthalmaHyperolius_semidiscusHyperolius_parkeri Hyperolius_lateralis Hyperolius_castaneus Hyperolius_frontalis Hyperolius_lupiroensisHyperolius_pusillus Hyperolius_jackie Hyperolius_viridis Hyperolius_cystocandicans Hyperolius_endjami_2 Hyperolius_endjami_1 Hyperolius_inyangae Hyperolius_mosaicus Hyperolius_tuberilinguis Hyperolius_koehleri Hyperolius_nasutus Hyperolius_viridiflavus_destefanii Hyperolius_viridiflavus_variablilis Hyperolius_benguellensis Hyperolius_viridiflavus_sp Hyperolius_dartevellei Hyperolius_viridiflavus_pitmani Hyperolius_rwandae Hyperolius_viridiflavus_pantherinus Hyperolius_viridiflavus_glandicolor Hyperolius_viridiflavus_ommatostictus Hyperolius_viridiflavus_ngorongoriensis Hyperolius_howelli Hyperolius_spatzi Hyperolius_friedemanniHyperolius_acuticeps Hyperolius_nitidulus_2 Hyperolius_nitidulus_1 Hyperolius_marmoratus_marmoratus Hyperolius_marmoratus_swynnertoni Hyperolius_mariae Hyperolius_reesi Hyperolius_marmoratus_taeniatus H H

y Hyperolius_adspersus y

s s

p Hyperolius_jacobseni p

Hyperolius_poweri

u u

l e

e t

e r

r a

Hyperolius_igbettensis l o o

l n

i l l

a i i

u r

g r

s a

s a _ _ Hyperolius_pseudoargus p

_ m

Hyperolius_fusciventris_1 a Hyperolius_bolifambae _

u r

Hyperolius_fusciventris_2 s

l Hyperolius_ocellatus_2 a a

Hyperolius_sp_nov_1 u

l Hyperolius_ocellatus_1 l l

l l l

l Hyperolius_argus e

e l

a l Hyperolius_riggenbachi l l

Hyperolius_pardalis u r

u a Hyperolius_bocagei Hyperolius_lamottei s

s r Hyperolius_guttulatus_1 _ _ Hyperolius_camerunensis p Hyperolius_guttulatus_2 a

a p

Hyperolius_phantasticus _ p

Hyperolius_dintelmanni y Hyperolius_steindachneri _ b

Hyperolius_tuberculatus

u s o

h u

r i

u l

f o

e u r

p s

c e

t p

y y

H n Monochromatism: Hidden State Absent Dichromatism: Hidden State Absent Monochromatism: Hidden State Present Dichromatism: Hidden State Present Lineage Tissue ID Country/Island Locality Locality Museum Number Institution Acanthixalus sonjae MORAS1 Cote d'Ivoire 05°49'59.9''N, W 007°23'08.1''W Tai National Park ZMB only tissue Museum für Naturkunde Berlin Acanthixalus sonjae MORAS2 Cote d'Ivoire 05°49'59.9''N, W 007°23'08.1''W Tai National Park ZMB only tissue Museum für Naturkunde Berlin Acanthixalus spinosus JMD 890 Gabon Parc National de l'Ivindo Pending Museum für Naturkunde, Berlin Afrixalus "brachycnemis" FMNH 274855 Malawi -12.11386, 33.72457 Vipya Plateau, Luwawa FMNH 274855 The Field Museum Afrixalus "brachycnemis" FMNH 274898 Malawi -16.01366, 35.65175 base of Mulanje Massif FMNH 274898 The Field Museum Afrixalus "brachycnemis" MVZ 265826 Mozambique -15.384275, 37.071958 Mt. Namuli, Zambezia Province MVZ 265826 Museum of Vertebrate Zoology Afrixalus "delicatus" FMNH 274871 Malawi -16.04831, 35.71076 base of Mulanje Massif FMNH 274871 The Field Museum Afrixalus "delicatus" GPN032 Mozambique 18.66485°S, 34.80526°E Gorongosa National Park Pending Museum für Naturkunde Berlin Afrixalus "delicatus" JH14 AA1 South Africa 28° 5'48"S, 32°16'26"E Malala, Hluhluwe, KwaZulu-Natal province Afrixalus "delicatus" MVZ 226254 Kenya -3.33333, 39.86667 Arabuko Sokoke Forest, Coast Province MVZ 226254 Museum of Vertebrate Zoology Afrixalus dorsalis 1 CAS 253855 Cameroon 4.95539, 9.86786 Manjo, Mt. Kupe, Littoral Region CAS 253855 California Academy of Sciences Afrixalus dorsalis 1 MVZ 234690 Cameroon 6.27555, -1.0081 Nguti, Mamfe, Sud-Ouest Region MVZ 234690 Museum of Vertebrate Zoology Afrixalus dorsalis 2 CUMV 15070 Gabon 01.80976°S, 09.38770°E Iguéla, Ogooue-Maritime Province CUMV 15070 Cornell University Museum of Vertebrates Afrixalus dorsalis 2 CUMV 15071 Gabon 01.80976°S, 09.38770°E Iguéla, Ogooue-Maritime Province CUMV 15071 Cornell University Museum of Vertebrates Afrixalus dorsalis 2 MBUR 3454 Republic of Congo 02 25 07.4 S, 12 45 16.5 E 3 km E of Tsinguidi ZMB 56734 Museum für Naturkunde Berlin Afrixalus dorsalis 3 MVZ 244954 Ghana 5.29004, -2.63957 Ankasa Natl Park, Western Region MVZ 244954 Museum of Vertebrate Zoology Afrixalus dorsalis 3 UWBM 5580 Ghana 6.72604 N, 01.52750 W Kumasi, Ashanti Region UWBM 5580 Burke Museum of Natural History and Culture Afrixalus dorsimaculatus 1 T676 Tanzania S04 48'45.1, E038 30'12.9" West Usambaras, Mazumbai Forest Reserve MW 01977 Natural History Museum, London Afrixalus dorsimaculatus 1 T677 Tanzania S04 48'45.1, E038 30'12.9" West Usambaras, Mazumbai Forest Reserve MW 01979 Natural History Museum, London Afrixalus dorsimaculatus 2 MVZ 265820 Mozambique -12.86737656, 35.17335137 Serra Jeci, Niassa Province MVZ 265820 Museum of Vertebrate Zoology Afrixalus enseticola TJC 873 Ethiopia 6.63893, 39.73375 Oromia, Bale Pending Zoological Natural History Museum, Addis Ababa University Afrixalus enseticola TJC 874 Ethiopia 6.63893, 39.73376 Oromia, Bale Pending Zoological Natural History Museum, Addis Ababa University Afrixalus fornasini FMNH 274842 Malawi -11.98449, 34.04609 Nkhata Bay FMNH 274842 The Field Museum Afrixalus fornasini MVZ 233800 Kenya -3.333, 39.867 Arabuko Sokoke Forest, Coast Province MVZ 233800 Museum of Vertebrate Zoology Afrixalus fulvovittatus 1 CAS 253641 Cameroon 05⁰56'07", 10⁰07'41" Nsongwa, Bamenda, Nord-Ouest Region CAS 253641 California Academy of Sciences Afrixalus fulvovittatus 2 CAS 253392 Cameroon 2.59461, 14.02827 Ngwoyla, Est Region CAS 253392 California Academy of Sciences Afrixalus fulvovittatus 2 CUMV 14918 Gabon 00.51121°N, 12.80278°E Ivindo National Park, Ogooue-Ivindo Province CUMV 14918 Cornell University Museum of Vertebrates Afrixalus fulvovittatus 2 JMD 847 Gabon Parc National de l'Ivindo Pending Museum für Naturkunde, Berlin Afrixalus knysae WC10-125 South Africa -33.95022, 23.60150 Western Cape, The Cove, Natures Valley PEM T534 Port Elizabeth Museum Afrixalus lacteus CAS 253841 Cameroon 4º 57' 33.98" N, 9º 51' 54.22" E Manengouba II, Mt. Manengouba, Littoral Region CAS 253841 California Academy of Sciences Afrixalus lacteus MH 0615 Cameroon 5.02594, 9.7629 Mt. Manengouba, Ebonemin ZMB 78975 Museum für Naturkunde Berlin Afrixalus lacustris CAS 256130 Uganda 0.4508 N, 32.9493 E Mabira Forest Reserve, Mukono District CAS 256130 California Academy of Sciences Afrixalus laevis CAS 253804 Cameroon 4.32228, 9.06635 Small Koto, Mt. Cameroon, Sud-Ouest Region CAS 253804 California Academy of Sciences Afrixalus laevis CAS 253845 Cameroon 4º 57' 33.98" N, 9º 51' 54.22" E Manengouba II, Mt. Manengouba, Littoral Region CAS 253845 California Academy of Sciences Afrixalus nigeriensis UWBM 5603 Ghana 6.24246 N, 0.55710 W Atewa Hills, Eastern Region UWBM 5603 Burke Museum of Natural History and Culture Afrixalus osorioi CAS 256140 Uganda 0.43226 N, 32.96107 E Mabira Forest Reserve, Mukono District CAS 256140 California Academy of Sciences Afrixalus osorioi MVZ 233823 Kenya 0.2663333, 34.80675 Kakamega Forest, Western Province MVZ 233823 Museum of Vertebrate Zoology Afrixalus paradorsalis manengubensis CAS 254144 Cameroon 4⁰57.636, 9⁰39.222 Edib Village, Sud-Ouest Region CAS 254144 California Academy of Sciences Afrixalus paradorsalis paradorsalis 1 CAS 249907 Cameroon 5°43.493' N, 9°20.963' E Mamfe, Sud-Ouest Region CAS 249907 California Academy of Sciences Afrixalus paradorsalis paradorsalis 2 CUMV 14900 Gabon 00.45358°N, 10.27809°E Monts de Cristal National Park, Woleu-Ntem Province CUMV 14900 Cornell University Museum of Vertebrates Afrixalus paradorsalis paradorsalis 2 MBUR 03778 Republic of Congo 02 25 14.9 S, 12 52 20.1 E 16 km E of Tsinguidi IRSEN 0369 Institut national de Recherche en Sciences Exactes et Naturelles Afrixalus quadrivittatus 1 CAS 250032 Cameroon 6°10.517' N, 10°04.474' E Lower Bafut, Nord-Ouest Region CAS 250032 California Academy of Sciences Afrixalus quadrivittatus 2 CAS 256142 Uganda 0.43226 N, 32.96107 E Mabira Forest Reserve, Mukono District CAS 256141 California Academy of Sciences Afrixalus spinifrons WC-DNA-832 South Africa -31.65386, 29.50683 Eastern Cape, Silaka Nature Reserve PEM A10559 Port Elizabeth Museum Afrixalus sylvaticus MVZ 234560 Kenya -4.2664167, 39.3405 Shimba Hills, Coast Province MVZ 234560 Museum of Vertebrate Zoology Afrixalus vibekensis T01.35 Cote d'Ivoire 05°49'59.9''N, W 007°23'08.1''W Tai National Park ZMB 78449 Museum für Naturkunde Berlin Afrixalus vittiger 1 MVZ 249415 Ghana 8.42942, 0.56544 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 249415 Museum of Vertebrate Zoology Afrixalus vittiger 2 UWBM 5615 Ghana 10.42018 N, 2.07402 W Gbele Reserve, Upper West Region UWBM 5615 Burke Museum of Natural History and Culture Afrixalus weidholzi MVZ 249448 Ghana 8.42942, 0.56544 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 249448 Museum of Vertebrate Zoology Afrixalus wittei ELI 376 DRC S7.3382, E28.0088 Luvua River near Kiambi, Katanga Province UTEP 21795 University of Texas at El Paso Biodiversity Collections Afrixalus wittei RB11-D028 Zambia -12.25386111, 25.34419444 Kalumbila airstrip pools, Kalumbila, Solwezi District SAIAB 185816 South African Institute for Aquatic Biodiversity Arthtroleptis poecilinotus CAS 253319 Cameroon 2.90072, 13.90334 Zoulabot II, Est Region CAS 253319 California Academy of Sciences Breviceps adspersus MVZ 265908 Mozambique -21.996, 35.323 Vilanculos, Inhambane Province MVZ 265908 Museum of Vertebrate Zoology Cardioglossa gracilis CAS 254213 Cameroon 5.15301, 10.53249 Bangangte, Ouest Region CAS 254213 California Academy of Sciences Cryptothylax greshoffii CUMV 15021 Gabon 00.49986°N, 12.80182°E Ivindo National Park, Ogooue-Ivindo Province CUMV 15021 Cornell University Museum of Vertebrates Cryptothylax greshoffii JMD 842 Gabon Parc National de l'Ivindo Pending Museum für Naturkunde, Berlin Cryptothylax greshoffii MVZ 234714 Cameroon 2.96625, 12.0816 Sangmelima, Sud Region MVZ 234714 Museum of Vertebrate Zoology Hemisus marmoratus MVZ 249304 Ghana 8.28022, 0.53053 Nkwanta, Volta Region MVZ 249304 Museum of Vertebrate Zoology Heterixalus alboguttatus MVZ 241451 Madagascar -21.25, 47.45 Ranomafana MVZ 241451 Museum of Vertebrate Zoology Heterixalus luteostriatus MVZ 238721 Madagascar -21.89067, 46.76333 SW Ambalavao, Fianarantsoa MVZ 238721 Museum of Vertebrate Zoology Hyperolius ademetzi CAS 253623 Cameroon 05⁰51.344', 10⁰09.512' Akum-Santa, Bamenda, Nord-Ouest Region CAS 253623 California Academy of Sciences Hyperolius ademetzi vg05-C020 Cameroon 6.2038, 10.4596 Lake Oku, Nord-Ouest Region NMP6V 73384 National Museum, Prague Hyperolius adspersus CAS 254255 Cameroon 5.10863, 10.61671 Bangangte, Ouest Region CAS 254255 California Academy of Sciences Hyperolius argus FMNH 274326 Tanzania S 04.82645°, E 038.78822° Mtai Forest, East Usambara Mountains FMNH 274326 The Field Museum Hyperolius argus MVZ 234112 Kenya -3.1999167, 40.0077 Lake Mbaratumo Ziwa, Kakuyuni, Coast Province MVZ 234112 Museum of Vertebrate Zoology Hyperolius balfouri balfouri CAS 256187 Uganda 0.44423 N, 32.97309 E Mabira Forest Reserve, Mukono District CAS 256187 California Academy of Sciences Hyperolius balfouri balfouri MVZ 238699 Kenya 0.042222222, 34.74694444 Buyangu, Kakamega District, Western Province MVZ 238699 Museum of Vertebrate Zoology Hyperolius balfouri viridistriatus CAS 253644 Cameroon 05⁰56'07", 10⁰07'41" Nsongwa, Bamenda, Nord-Ouest Region CAS 253644 California Academy of Sciences Hyperolius balfouri viridistriatus CAS 254229 Cameroon 5.15301, 10.53249 Bangangte, Ouest Region CAS 254229 California Academy of Sciences Hyperolius baumanni MVZ 244982 Ghana 8.3226, 0.55622 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 244982 Museum of Vertebrate Zoology Hyperolius baumanni MVZ 249308 Ghana 8.40584, 0.59387 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 249308 Museum of Vertebrate Zoology Hyperolius bobirensis UWBM 5651 Ghana 6.24246 N, 0.55710 W Atewa Hills, Eastern Region UWBM 5651 Burke Museum of Natural History and Culture Hyperolius bocagei MTD-TD 14272 Angola S 6,78086 E 16,17629 Quimbele, Uíge Province MTD 48946 Senckenberg Natural History Collections Dresden Hyperolius bolifambae CAS 253509 Cameroon 3.17376, 12.52714 Mekas, Dja Reserve, Sud Region CAS 253509 California Academy of Sciences Hyperolius bolifambae CAS 253883 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253883 California Academy of Sciences Hyperolius burgessi CAS 169942 Tanzania 5 5 60 S, 38 37 60 W Amani, East Usambara Mtns CAS 169942 California Academy of Sciences Hyperolius burgessi MVZ 233942 Tanzania -5.1000333, 38.6266333 Amani Nature Reserve, Amani, Usambara Mountains MVZ 233942 Museum of Vertebrate Zoology Hyperolius camerunensis CAS 253935 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253935 California Academy of Sciences Hyperolius camerunensis CAS 253937 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253937 California Academy of Sciences Hyperolius castaneus CAS 256022 Uganda 1.05315 S, 29.76138 E Bwindi Impenetrable Forest, Kabale District CAS 256022 California Academy of Sciences Hyperolius castaneus EBG 1706 DRC S3.36255, E28.87713 Magunda, Itombwe Plateau, South Kivu UTEP 20652 University of Texas at El Paso Biodiversity Collections Hyperolius chlorosteus FO86 Guinea N 08°31.499', W 08°56.204' Simandou, Pic de Fon area ZMB 80739 Museum für Naturkunde Berlin Hyperolius chlorosteus JP0136 Sierra Leone 8º30.035' N, 11º08.800'W Nimni Forest Reserve ZMB 80738 Museum für Naturkunde Berlin Hyperolius cinnamomeoventris 1 CAS 256160 Uganda 0.43226 N, 32.96107 E Mabira Forest Reserve, Mukono District CAS 256160 California Academy of Sciences Hyperolius cinnamomeoventris 1 MVZ 233830 Kenya 0.2663333, 34.80675 Kakamega Forest, Western Province MVZ 233830 Museum of Vertebrate Zoology Hyperolius cinnamomeoventris 2 CAS 253471 Cameroon 2.93994, 11.97633 Sangmelima, Sud Region CAS 253471 California Academy of Sciences Hyperolius cinnamomeoventris 2 CUMV 14953 Gabon 00.51121°N, 12.80278°E Ivindo National Park, Ogooue-Ivindo Province CUMV 14953 Cornell University Museum of Vertebrates Hyperolius concolor concolor MVZ 244987 Ghana 8.3226, 0.55621 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 244987 Museum of Vertebrate Zoology Hyperolius concolor guttatus CAS 253874 Cameroon 4.95539, 9.86786 Manjo, Mt. Kupe, Littoral Region CAS 253874 California Academy of Sciences Hyperolius concolor guttatus CAS 253914 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253914 California Academy of Sciences Hyperolius concolor ibadanensis CAS 249915 Cameroon 5°43.493' N, 9°20.963' E Mamfe, Sud-Ouest Region CAS 249915 California Academy of Sciences Hyperolius cystocandicans MVZ 233908 Kenya -1.1241667, 36.67375 Tigoni, Nairobi Province MVZ 233908 Museum of Vertebrate Zoology Hyperolius cystocandicans MVZ 234120 Kenya -1.2446833, 36.65705 Ondiri Swamp, Kikuyu, Nairobi Province MVZ 234120 Museum of Vertebrate Zoology Hyperolius dintelmanni CAS 253991 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253991 California Academy of Sciences Hyperolius dintelmanni CAS 254135 Cameroon 4⁰57.636, 9⁰39.222 Edib Village, Sud-Ouest Region CAS 254135 California Academy of Sciences Hyperolius discodactylus CAS 256074 Uganda 1º 4' 26.4" S, 29º 45' 57.6" E Bwindi Impenetrable Forest, Kabale District CAS 256074 California Academy of Sciences Hyperolius discodactylus JMD 633 Rwanda Nyungwe ZMB 78951 Museum für Naturkunde, Berlin Hyperolius drewesi CAS 219127 Sao Tome and Principe 1 39 7.7 N, 7 24 57.8 E Chada, Principe Id CAS 219127 California Academy of Sciences Hyperolius endjami 1 CUMV 15239 Bioko Island 3.4673166667, 8.6411 Moeri, Bioko Id CUMV 15239 Cornell University Museum of Vertebrates Hyperolius endjami 2 vg09-275 Cameroon 4.96344, 9.67492 Edib Village, Bakossi Mountains, Sud-Ouest Region NMP6V 74622/1 National Museum, Prague Hyperolius endjami 2 vg09-276 Cameroon 4.96344, 9.67492 Edib Village, Bakossi Mountains, Sud-Ouest Region NMP6V 74622/2 National Museum, Prague Hyperolius frontalis CAS 201987 Uganda 0 58 42 S, 29 41 42 E Bwindi Impenetrable National Park, Rukungiri Dist. CAS 201987 California Academy of Sciences Hyperolius frontalis CAS 256039 Uganda 1.04903 S, 29.79025 E Bwindi Impenetrable Forest, Kabale District CAS 256039 California Academy of Sciences Hyperolius fusciventris burtoni MVZ 245012 Ghana 5.28434, -2.64861 Ankasa Natl Park, Western Region MVZ 245012 Museum of Vertebrate Zoology Hyperolius fusciventris ssp. CAS 254005 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 254005 California Academy of Sciences Hyperolius guttulatus 1 vg10-C43/09 Cameroon 4.96413, 8.907116 Mundemba, Sud-Ouest Region NMP6V 74552 National Museum, Prague Hyperolius guttulatus 2 UWBM 5695 Ghana 6.83673 N, 1.72202 W Barekese Watershed, Ashanti Region UWBM 5695 Burke Museum of Natural History and Culture Hyperolius jackie JMD 671 Rwanda Nyungwe ZMB 77477 Museum für Naturkunde, Berlin Hyperolius jackie JMD 675 Rwanda Nyungwe ZMB 77480 Museum für Naturkunde, Berlin Hyperolius kivuensis 1 CAS 256162 Uganda 0.43226 N, 32.96107 E Mabira Forest Reserve, Mukono District CAS 256162 California Academy of Sciences Hyperolius kivuensis 1 JMD 628 Rwanda Rukarara Pending Museum für Naturkunde, Berlin Hyperolius kivuensis 2 RB11-D025 Zambia -12.25386111, 25.34419444 Kalumbila airstrip pools, Kalumbila, Solwezi District SAIAB 185850 South African Institute for Aquatic Biodiversity Hyperolius kivuensis 2 RB11B-343 Zambia -9.362555556, 30.12366667 Fields of Mporokoso SAIAB 98724 South African Institute for Aquatic Biodiversity Hyperolius kivuensis 3 FMNH 274889 Malawi -14.24392, 35.39216 Mtai Forest, East Usambara Mountains FMNH 274889 The Field Museum Hyperolius kivuensis 3 FMNH 274940 Malawi -14.24392, 35.39216 Mtai Forest, East Usambara Mountains FMNH 274940 The Field Museum Hyperolius koehleri MH 0536 Cameroon 5.00983, 9.8568 Mt. Manengouba, near summit ZMB 82052 Museum für Naturkunde Berlin Hyperolius koehleri MH 0550 Cameroon 5.01291, 9.7682 Mt. Manengouba, Ebonemin ZMB 82053 Museum für Naturkunde Berlin Hyperolius kuligae CAS 253675 Cameroon 05⁰55'34", 10⁰07'32" Nsongwa, Bamenda, Nord-Ouest Region CAS 253675 California Academy of Sciences Hyperolius kuligae CUMV 14931 Gabon 00.51121°N, 12.80278°E Ivindo National Park, Ogooue-Ivindo Province CUMV 14931 Cornell University Museum of Vertebrates Hyperolius lamottei CAS 230152 Sierra Leone 8 5 24 N, 12 26 51.6 W Moyamba, Moyamba District CAS 230152 California Academy of Sciences Hyperolius lamottei CAS 230153 Sierra Leone 08 05.40 N, 12 26.86 W Moyamba, Moyamba District CAS 230153 California Academy of Sciences Hyperolius langi CAS 256132 Uganda 0.4508 N, 32.9493 E Mabira Forest Reserve, Mukono District CAS 256132 California Academy of Sciences Hyperolius lateralis CAS 256158 Uganda 0.43226 N, 32.96107 E Mabira Forest Reserve, Mukono District CAS 256158 California Academy of Sciences Hyperolius lateralis JMD 652 Rwanda Butare Pending Museum für Naturkunde, Berlin Hyperolius lateralis MVZ 233858 Kenya 0.2663333, 34.80675 Kakamega Forest, Western Province MVZ 233858 Museum of Vertebrate Zoology Hyperolius mariae FMNH 274379 Tanzania S 04.82645°, E 038.78822° Mtai Forest, East Usambara Mountains FMNH 274379 The Field Museum Hyperolius mariae MVZ 233862 Tanzania -6.6067583, 38.8202167 S Mlandizi, Dar es Salaam Region MVZ 233862 Museum of Vertebrate Zoology Hyperolius marmoratus marmoratus RSP193 South Africa -29.9396667, 30.9935837 Bluff Nature Reserve, KwaZulu-Natal SANBI 5878 South African National Biodiversity Institute Hyperolius marmoratus swynnertoni GPN036 Mozambique 18.66485°S, 34.80526°E Gorongosa National Park ZMB not yet accessioned Museum für Naturkunde Berlin Hyperolius marmoratus taeniatus 2010.0185 Mozambique 10°42’08.47”S, 40°12’24.46”E Nhica do Rovuma, Cabo Delgado Province MNHN 2010.185 Muséum National d’Histoire Naturelle Hyperolius marmoratus taeniatus MVZ 266048 Mozambique -15.384275, 37.071958 Mt. Namuli, Zambezia Province MVZ 266048 Museum of Vertebrate Zoology Hyperolius minitissimus FMNH 274290 Tanzania S 08.3295°, E 035.939983° Ivalla River, Kilolo FMNH 274290 The Field Museum Hyperolius minitissimus T2969 Tanzania -8.3721, 35.98133 Kihanga MUSE 11024 Trento Museum of Science Hyperolius mitchelli MVZ 233890 Tanzania -6.6067583, 38.8202167 S Mlandizi, Dar es Salaam Region MVZ 233890 Museum of Vertebrate Zoology Hyperolius mitchelli MVZ 233891 Tanzania -6.6067583, 38.8202167 S Mlandizi, Dar es Salaam Region MVZ 233891 Museum of Vertebrate Zoology Hyperolius molleri CAS 218849 Sao Tome and Principe 0 17 52.8 N, 6 43 49.3 E Caxueira, Sao Tome Id CAS 218849 California Academy of Sciences Hyperolius montanus SL 131 Kenya Hyperolius mosaicus CAS 253533 Cameroon 3.17916, 12.53352 Mekas, Dja Reserve, Sud Region CAS 253533 California Academy of Sciences Hyperolius mosaicus JMD 938 Gabon Parc National des Monts de Cristal Pending Museum für Naturkunde, Berlin Hyperolius nitidulus 1 BJE 3203 Cameroon 6°32.917' N, 10°45.599' E Nkambe, Nord-Ouest Region MCZ A148082 Museum of Comparative Zoology Hyperolius nitidulus 1 CAS 249894 Cameroon 5°58.063' N, 10°02.382' E Bamenda, Nord-Ouest Region CAS 249894 California Academy of Sciences Hyperolius nitidulus 1 CAS 254241 Cameroon 5.15301, 10.53249 Bangangte, Ouest Region CAS 254241 California Academy of Sciences Hyperolius nitidulus 1 CAS 254242 Cameroon 5.15301, 10.53249 Bangangte, Ouest Region CAS 254242 California Academy of Sciences Hyperolius nitidulus 2 MVZ 245049 Ghana 5.88026, 0.03782 Shai Hills Preserve, Accra Region MVZ 245049 Museum of Vertebrate Zoology Hyperolius nitidulus 2 MVZ 249370 Ghana 9.25988, -1.86001 Mole Natl Park, Northern Region MVZ 249370 Museum of Vertebrate Zoology Hyperolius occidentalis B017 Guinea Boké Prefecture, Kalaboui ZMB 74879 Museum für Naturkunde Berlin Hyperolius occidentalis B090 Guinea Boké Prefecture, Kamsar, village Filima near Taigbe East ZMB 74880 Museum für Naturkunde Berlin Hyperolius ocellatus ocellatus CAS 254074 Cameroon 4º 50' 59.1" N, 9º 46' 18.5874" E Manjo, Mt. Kupe, Littoral Region CAS 254074 California Academy of Sciences Hyperolius ocellatus ocellatus CUMV 15205 Bioko Island 03 28.039, 08 38.466 Moeri, Bioko Id CUMV 15205 Cornell University Museum of Vertebrates Hyperolius ocellatus purpurescens CAS 253588 Cameroon 3.17334, 12.52902 Mekas, Dja Reserve, Sud Region CAS 253588 California Academy of Sciences Hyperolius olivaceus CUMV 15088 Gabon 01.80976°S, 09.38770°E Iguéla, Ogooue-Maritime Province CUMV 15088 Cornell University Museum of Vertebrates Hyperolius olivaceus CUMV 15089 Gabon 01.80976°S, 09.38770°E Iguéla, Ogooue-Maritime Province CUMV 15089 Cornell University Museum of Vertebrates Hyperolius parallelus alborufus RB11-D189 Zambia -12.18691667, 25.21452778 Enterprise camp, Solwezi District SAIAB 185835 South African Institute for Aquatic Biodiversity Hyperolius parallelus marginatus MI11-181 Zambia -12.28622222, 30.6215 Nkondo camp SAIAB 99247 South African Institute for Aquatic Biodiversity Hyperolius parallelus parallelus AMB 7989 Namibia 18 02 04.7S, 20 51 41.0 E Shamvura Camp, Kavango Region MCZ A148568 Museum of Comparative Zoology Hyperolius parallelus pyrrhodictyon RB11B-306 Zambia -15.60325, 28.26433333 Eureka camping site, Lusaka SAIAB 98764 South African Institute for Aquatic Biodiversity Hyperolius pardalis CAS 253428 Cameroon 2.59461, 14.02827 Ngwoyla, Est Region CAS 253428 California Academy of Sciences Hyperolius pardalis vg10-144 Cameroon 2.44217, 15.43078 Mambele, Est Region NMP6V 74701/2 National Museum, Prague Hyperolius parkeri FMNH 274401 Tanzania S 05.1133° E, E 038.75257° Magorotto Forest, East Usambara FMNH 274401 The Field Museum Hyperolius parkeri MVZ 233913 Kenya 04 15.042 S, 39 20.654 E Shimba Hills, Coast Province MVZ 233913 Museum of Vertebrate Zoology Hyperolius phantasticus CUMV 14898 Gabon 00.45358°N, 10.27809°E Monts de Cristal National Park, Woleu-Ntem Province CUMV 14898 Cornell University Museum of Vertebrates Hyperolius phantasticus CUMV 14899 Gabon 00.45358°N, 10.27809°E Monts de Cristal National Park, Woleu-Ntem Province CUMV 14899 Cornell University Museum of Vertebrates Hyperolius picturatus MVZ 252512 Ghana 6.27555, -1.0081 Mamang River Forest Reserve, Eastern Region MVZ 252512 Museum of Vertebrate Zoology Hyperolius picturatus UWBM 5723 Ghana 6.24246 N, 0.55710 W Atewa Hills, Eastern Region UWBM 5723 Burke Museum of Natural History and Culture Hyperolius pictus FMNH 274426 Tanzania S 08.09955°, E 035.89848° Dabaga, Kilolo FMNH 274426 The Field Museum Hyperolius pictus FMNH 274427 Tanzania S 08.09955°, E 035.89848° Dabaga, Kilolo FMNH 274427 The Field Museum Hyperolius platyceps CAS 249967 Cameroon 4°36.693' N, 12°13.525' E W Ndombam, Centre Region CAS 249967 California Academy of Sciences Hyperolius platyceps CG15-084 Republic of Congo -3.2923, 16.1623 Talangai NMP6V 75685/1 National Museum, Prague Hyperolius platyceps CG15-085 Republic of Congo -3.2923, 16.1623 Talangai NMP6V 75685/2 National Museum, Prague Hyperolius platyceps CUMV 15094 Gabon 01.77104°S, 09.36941°E Iguéla, Ogooue-Maritime Province CUMV 15094 Cornell University Museum of Vertebrates Hyperolius platyceps MBUR 03627 Republic of Congo 02 12 38.3 S, 12 49 56.6 E 1.6 km NE of Lehala, about 9 km NNE of Mayoko Poste IRSEN 0411 Institut national de Recherche en Sciences Exactes et Naturelles Hyperolius pseudoargus FMNH 274495 Tanzania S 08.51464°, E 035.4540583° Fox Farm, Mufindi FMNH 274495 The Field Museum Hyperolius pusillus MVZ 233930 Tanzania -6.6067583, 38.8202167 S Mlandizi, Dar es Salaam Region MVZ 233930 Museum of Vertebrate Zoology Hyperolius pusillus MVZ 234123 Kenya -3.15845, 39.07105 Tsavo East Natl Park, Coast Province MVZ 234123 Museum of Vertebrate Zoology Hyperolius quinquevittatus FMNH 275139 Malawi -11.66565, 33.7613 Vipya Plateau, Luwawa FMNH 275139 The Field Museum Hyperolius raymondi ANG-0047 Angola -7.74422, 19.95467 Lunda Norte, Lagoa Carumbo PEM A10048 Port Elizabeth Museum Hyperolius reesi FMNH 274348 Tanzania S 07.8421° , E 034.87814° Mang'ula Village, base of Udzungwa Mountains FMNH 274348 The Field Museum Hyperolius reesi FMNH 274349 Tanzania S 07.8421° , E 034.87814° Mang'ula Village, base of Udzungwa Mountains FMNH 274349 The Field Museum Hyperolius riggenbachi CAS 249896 Cameroon 6°00.855' N, 10°16.216' E Bamenda, Nord-Ouest Region CAS 249896 California Academy of Sciences Hyperolius riggenbachi CAS 250002 Cameroon 6°32.917' N, 10°45.599' E Nkambe, Nord-Ouest Region CAS 250002 California Academy of Sciences Hyperolius riggenbachi vg09-092 Cameroon 7.74049, 12.72305 Tchabal Gangdaba, Nord Region NMP6V 74576/1 National Museum, Prague Hyperolius rubrovermiculatus MVZ 233887 Tanzania 05 6.002 S, 38 37.598 E Amani Nature Reserve, Amani, Usambara Mountains MVZ 233887 Museum of Vertebrate Zoology Hyperolius rubrovermiculatus MVZ 233893 Kenya -3.333, 39.867 Arabuko Sokoke Forest, Coast Province MVZ 233893 Museum of Vertebrate Zoology Hyperolius rubrovermiculatus MVZ 233958 Kenya -4.2664167, 39.3405 Shimba Hills, Coast Province MVZ 233958 Museum of Vertebrate Zoology Hyperolius semidiscus WC-DNA-289 South Africa -32.28392, 28.84885 Eastern Cape, Silaka Nature Reserve PEM A10586 Port Elizabeth Museum Hyperolius semidiscus WC-DNA-876 South Africa -31.65010, 29.50513 Eastern Cape, Dwesa Nature Reserve PEM A10118 Port Elizabeth Museum Hyperolius sp. nov. CAS 253945 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253945 California Academy of Sciences Hyperolius sp. nov. CAS 254177 Cameroon 5.17626, 10.34873 Bangoua, Ouest Region CAS 254177 California Academy of Sciences Hyperolius spatzi SA120 Senegal N 13°09.368', W 12°06.882' Sabodala ZMB 74279 Museum für Naturkunde Berlin Hyperolius spatzi SA154 Senegal N 13°07.259', W 12°07.622' Sabodala ZMB 74285 Museum für Naturkunde Berlin Hyperolius spinigularis MVZ 266050 Mozambique -15.384275, 37.071958 Mt. Namuli, Zambezia Province MVZ 266050 Museum of Vertebrate Zoology Hyperolius steindachneri ANG-0112 Angola -8.33847, 20.24250 Lunde Norte, Capaia PEM A10029 Port Elizabeth Museum Hyperolius stictus MZ_1 Mozambique -10.76161, 40.42922 Cabo Delgabo Province, Palma PEM A11704 Port Elizabeth Museum Hyperolius stictus MZ_2 Mozambique -10.76161, 40.42922 Cabo Delgabo Province, Palma PEM A11705 Port Elizabeth Museum Hyperolius substriatus CAS 169105 Tanzania 4 47 60 S, 38 30 0 W Mazumbai Forest Reserve, West Usamabara Mtns CAS 169105 California Academy of Sciences Hyperolius substriatus MVZ 266019 Mozambique -12.84388577, 35.17817779 Serra Jeci, Niassa Province MVZ 266019 Museum of Vertebrate Zoology Hyperolius substriatus MVZ 266026 Mozambique -15.384275, 37.071958 Mt. Namuli, Zambezia Province MVZ 266026 Museum of Vertebrate Zoology Hyperolius sylvaticus MVZ 245057 Ghana 5.28434, -2.64861 Ankasa Natl Park, Western Region MVZ 245057 Museum of Vertebrate Zoology Hyperolius tanneri FMNH 274288 Tanzania S 04.829116° , E 038.51273° Shume Magamba, West Usambara FMNH 274288 The Field Museum Hyperolius tanneri FMNH 274289 Tanzania S 04.74783°, E 038.29743° Mazumbi, West Usambara FMNH 274289 The Field Museum Hyperolius thomensis CAS 233470 Sao Tome and Principe 0 16 34 N, 6 36 20 E Bom Sucesso, Sao Tome Id CAS 233470 California Academy of Sciences Hyperolius thomensis CAS 233471 Sao Tome and Principe 0 16 34 N, 6 36 20 E Bom Sucesso, Sao Tome Id CAS 233471 California Academy of Sciences Hyperolius torrentis MVZ 249336 Ghana 8.33383, 0.59449 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 249336 Museum of Vertebrate Zoology Hyperolius torrentis MVZ 249402 Ghana 8.43424, 0.60552 Kyabobo Natl Park, Togo Hills, Volta Region MVZ 249402 Museum of Vertebrate Zoology Hyperolius tuberculatus CAS 253400 Cameroon 2.59461, 14.02827 Ngwoyla, Est Region CAS 253400 California Academy of Sciences Hyperolius tuberculatus CAS 254219 Cameroon 5.15301, 10.53249 Bangangte, Ouest Region CAS 254219 California Academy of Sciences Hyperolius tuberculatus CUMV 14908 Gabon 00.62120°N, 10.40763°E Monts de Cristal National Park, Woleu-Ntem Province CUMV 14908 Cornell University Museum of Vertebrates Hyperolius tuberilinguis FMNH 274970 Malawi -16.04831, 35.71076 base of Mulanje Massif FMNH 274970 The Field Museum Hyperolius tuberilinguis MVZ 234126 Kenya -3.1699667, 39.8628333 Kakoneni, Coast Province MVZ 234126 Museum of Vertebrate Zoology Hyperolius veithi JK 702 DRC -2.7531 20.3781 Hyperolius viridiflavus destefanii TJC 476 Ethiopia 7.09845, 38.4164 Oromia, Awassa Pending Zoological Natural History Museum, Addis Ababa University Hyperolius viridiflavus destefanii TJC 591 Ethiopia 6.58604, 38.38871 Oromia, Awassa Pending Zoological Natural History Museum, Addis Ababa University Hyperolius viridiflavus glandicolor CAS 191491 Kenya 3 30 41.4 S, 38 15 7.8 E Taita Hills, Taita Dist. CAS 191491 California Academy of Sciences Hyperolius viridiflavus ngorongoriensis CAS 191455 Tanzania 3 22 24 S, 35 51 18 E Mto-Wa-Mbu, Monduli Dist. CAS 191455 California Academy of Sciences Hyperolius viridiflavus ommatostictus CAS 191407 Tanzania 3 21 33 S, 36 50 14.4 E Ngare Sero, Arumeru Dist. CAS 191407 California Academy of Sciences Hyperolius viridiflavus pantherinus CAS 191374 Kenya 0 9 18 S, 37 0 36 E Naro Moru Lodge, Laikipia Dist. CAS 191374 California Academy of Sciences Hyperolius viridiflavus pitmani CAS 202293 Uganda 0 59 12.9 S, 29 38 3.5 E Bwindi Impenetrable National Park, Rukungiri Dist. CAS 202293 California Academy of Sciences Hyperolius viridiflavus ssp. JMD 564 Rwanda Lac Muhazi Pending Museum für Naturkunde, Berlin Hyperolius viridiflavus ssp. JMD 691 Rwanda Marais Rugesi Pending Museum für Naturkunde, Berlin Hyperolius viridiflavus variablilis CAS 256267 Uganda 0.4293 N, 33.1958 E Jinja, Jinja District CAS 256267 California Academy of Sciences Kassina arboricola UWBM 5746 Ghana 6.23899 N, 0.55824 W Atewa Hills, Eastern Region UWBM 5746 Burke Museum of Natural History and Culture Kassina arboricola UWBM 5747 Ghana 6.23899 N, 0.55824 W Atewa Hills, Eastern Region UWBM 5747 Burke Museum of Natural History and Culture Kassina cochranae FO 35 Guinea 08 31.861' N, 08 54.361' W Simandou, Pic de Fon area ZMB 75797 Museum für Naturkunde Berlin Kassina cochranae FO 36 Guinea 08 31.861' N, 08 54.361' W Simandou, Pic de Fon area ZMB 75797 Museum für Naturkunde Berlin Kassina decorata CAS 253990 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253990 California Academy of Sciences Kassina fusca FD75 Guinea 11 46.073 N, 12 40.844' W Fouta Djalon, Foret de Nialama, next to village Linsan ZMB 75806 Museum für Naturkunde Berlin Kassina kuvangensis WC12-A125 Angola -13.71617, 17.09661 Caundo Cubango, Cuelei River PEM T571 Port Elizabeth Museum Kassina lamottei K14 Cote d'Ivoire 05°50'03.5''N, 007°20'57.0''W Tai National Park ZMB only tissue Museum für Naturkunde Berlin Kassina maculosa 05-C068 Cameroon 6.13263, 10.17813 Bamo Forest, Nord-Ouest Region NMP6V 73396 National Museum, Prague Kassina senegalensis CAS 256210 Uganda 0.44423 N, 32.97309 E Mabira Forest Reserve, Mukono District CAS 256210 California Academy of Sciences Kassina senegalensis FMNH 274853 Malawi -16.04831, 35.71076 base of Mulanje Massif FMNH 274853 The Field Museum Kassina senegalensis JWH10-A115 Zambia -11.83111111, 31.42580556 Kalumbila Forest SAIAB 98458 South African Institute for Aquatic Biodiversity Kassina senegalensis MCZ F 38743 South Africa 22° 42' 23" S, 29° 49' 44" E Limpopo Province Kassina senegalensis QQ0678 Malawi -16.02456, 35.49902 Mt. Mulanje, Dambo SAIAB 96453 South African Institute for Aquatic Biodiversity Kassina senegalensis UWBM 5765 Ghana 8.29326 N, 2.28347 W Bui National Park, Brong-Ahafo Region UWBM 5765 Burke Museum of Natural History and Culture Kassina senegalensis WC-DNA-394 South Africa -31.82296, 29.30773 Eastern Cape, Hluleka Nature Reserve PEM A9962 Port Elizabeth Museum Kassina senegalensis WC12-A014 Angola -14.67458, 17.73544 Caundo Cubango, Menongue PEM T574 Port Elizabeth Museum Leptodactylodon ovatus CAS 253933 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253933 California Academy of Sciences Leptopelis rufus CAS 254132 Cameroon 4.9606, 9.6686 Edib Village, Sud-Ouest Region CAS 254132 California Academy of Sciences Morerella cyanophthalma Ba04.2 Cote d'Ivoire N 05°25', W 04°03’ Banco NP ZMB 71589 (paratype) Museum für Naturkunde Berlin Morerella cyanophthalma Ba04.3 Cote d'Ivoire N 05°25', W 04°03’ Banco NP ZMB 71590 (paratype) Museum für Naturkunde Berlin Nyctibates corrugatus CAS 253794 Cameroon 4.32676, 9.06609 Small Koto, Mt. Cameroon, Sud-Ouest Region CAS 253794 California Academy of Sciences Opisthothylax immaculatus CUMV 15167 Bioko Island 03 27.181, 08 30.410 Batete, Bioko Id CUMV 15167 Cornell University Museum of Vertebrates Opisthothylax immaculatus JMD 858 Gabon Parc National de l'Ivindo Pending Museum für Naturkunde, Berlin Opisthothylax immaculatus MVZ 234817 Cameroon 2.96625, 12.0816 Sangmelima, Sud Region MVZ 234817 Museum of Vertebrate Zoology Paracassina kounhiensis TJC 869 Ethiopia 6.63893, 39.73374 Oromia, Bale Pending Zoological Natural History Museum, Addis Ababa University Paracassina obscura T1256 Ethiopia 7.24932, 36.2554 Southern Nations, Bonga Town AK 1728 Addis Ababa University Phlyctimantis boulengeri MVZ 245085 Ghana 5.29004, -2.63957 Ankasa Natl Park, Western Region MVZ 245085 Museum of Vertebrate Zoology Phlyctimantis leonardi CAS 253979 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 253979 California Academy of Sciences Phlyctimantis leonardi JMD 929 Gabon Parc National des Monts de Cristal Pending Museum für Naturkunde, Berlin Phlyctimantis leonardi NCSM 76834 Gabon 0.45358 N, 10.27809 E Monts de Cristal National Park, Woleu-Ntem Province NCSM 76834 North Carolina Museum of Natural Sciences Phlyctimantis maculata FMNH 274820 Malawi -16.01366, 35.65175 base of Mulanje Massif FMNH 274820 The Field Museum Phlyctimantis maculata RB10-A111 Mozambique -15.05994444, 40.67088889 Nampula, Lumbo SAIAB 88587 South African Institute for Aquatic Biodiversity Phlyctimantis verrucosus CAS 256033 Uganda 1.04903 S, 29.79025 E Bwindi Impenetrable Forest, Kabale District CAS 256033 California Academy of Sciences Phlyctimantis verrucosus EBG 1774 DRC N00.68335, E29.67143 Virunga Natl Pak near Ndjuma, North Kivu Province UTEP 21796 University of Texas at El Paso Biodiversity Collections Phrynomantis microps MVZ 249480 Ghana 9.259, -1.85416 Mole Natl Park, Northern Region MVZ 249480 Museum of Vertebrate Zoology Scotobleps gabonicus CAS 254015 Cameroon 4.84975, 9.77183 Manjo, Mt. Kupe, Littoral Region CAS 254015 California Academy of Sciences Semnodactylus wealii SVN-005c South Africa -29.3805561, 29.6596394 Kamberg, KwaZulu-Natal SANBI 4627 South African National Biodiversity Institute Tachycnemis seychellensis Glaw 10 Seychelles Islands Mahe Tachycnemis seychellensis Glaw 2 Seychelles Islands Praslin Trichobatrachus robustus CAS 254090 Cameroon 4.94781, 9.69563 Ekenzu, Sud-Ouest Region CAS 254090 California Academy of Sciences Family Lineage Color Score: Monochromatic (0), Dichromatic (1) Arthroleptidae Arthroleptis adelphus 0 Arthroleptidae Arthroleptis affinis 0 Arthroleptidae Arthroleptis bioko 0 Arthroleptidae Arthroleptis brevipes 0 Arthroleptidae Arthroleptis fichika 0 Arthroleptidae Arthroleptis francei 0 Arthroleptidae Arthroleptis nikeae 0 Arthroleptidae Arthroleptis palava 0 Arthroleptidae Arthroleptis perreti 0 Arthroleptidae Arthroleptis poecilonotus 0 Arthroleptidae Arthroleptis reichei 0 Arthroleptidae Arthroleptis schubotzi 0 Arthroleptidae Arthroleptis stenodactylus 0 Arthroleptidae Arthroleptis sylvaticus 0 Arthroleptidae Arthroleptis taeniatus 0 Arthroleptidae Arthroleptis tanneri 0 Arthroleptidae Arthroleptis variabilis 0 Arthroleptidae Arthroleptis xenodactyloides 0 Arthroleptidae Astylosternus batesi 0 Arthroleptidae Astylosternus diadematus 0 Arthroleptidae Astylosternus laticephalus 0 Arthroleptidae Astylosternus occidentalis 0 Arthroleptidae Astylosternus schioetzi 0 Arthroleptidae Cardioglossa annulata 0 Arthroleptidae Cardioglossa elegans 0 Arthroleptidae Cardioglossa escalerae 0 Arthroleptidae Cardioglossa gracilis 0 Arthroleptidae Cardioglossa leucomystax 0 Arthroleptidae Cardioglossa manengouba 0 Arthroleptidae Cardioglossa melanogaster 0 Arthroleptidae Cardioglossa nigromaculata 0 Arthroleptidae Cardioglossa occidentalis 0 Arthroleptidae Cardioglossa oreas 0 Arthroleptidae Cardioglossa pulchra 0 Arthroleptidae Cardioglossa trifasciata 0 Arthroleptidae Leptodactylodon axillaris 0 Arthroleptidae Leptodactylodon bicolor 0 Arthroleptidae Leptodactylodon boulengeri 0 Arthroleptidae Leptodactylodon bueanus 0 Arthroleptidae Leptodactylodon erythrogaster 0 Arthroleptidae Leptodactylodon mertensi 0 Arthroleptidae Leptodactylodon ornatus 0 Arthroleptidae Leptodactylodon ovatus 0 Arthroleptidae Leptodactylodon perreti 0 Arthroleptidae Leptodactylodon polyacanthus 0 Arthroleptidae Leptodactylodon ventrimarmoratus 0 Arthroleptidae Leptopelis argenteus 0 Arthroleptidae Leptopelis aubryi 0 Arthroleptidae Leptopelis aubryioides 0 Arthroleptidae Leptopelis bocagii 0 Arthroleptidae Leptopelis boulengeri 0 Arthroleptidae Leptopelis brevirostris 0 Arthroleptidae Leptopelis calcaratus 0 Arthroleptidae Leptopelis fiziensis 0 Arthroleptidae Leptopelis flavomaculatus 0 Arthroleptidae Leptopelis gramineus 0 Arthroleptidae Leptopelis karissimbensis 0 Arthroleptidae Leptopelis kivuensis 0 Arthroleptidae Leptopelis macrotis 0 Arthroleptidae Leptopelis millsoni 0 Arthroleptidae Leptopelis modestus 0 Arthroleptidae Leptopelis natalensis 0 Arthroleptidae Leptopelis nordequatorialis 0 Arthroleptidae Leptopelis notatus 0 Arthroleptidae Leptopelis ocellatus 0 Arthroleptidae Leptopelis palmatus 0 Arthroleptidae Leptopelis parkeri 0 Arthroleptidae Leptopelis rufus 0 Arthroleptidae Leptopelis spiritusnoctis 0 Arthroleptidae Leptopelis susanae 0 Arthroleptidae Leptopelis vannutellii 0 Arthroleptidae Leptopelis vermiculatus 0 Arthroleptidae Leptopelis viridis 0 Arthroleptidae Leptopelis yaldeni 0 Arthroleptidae Nyctibates corrugatus 0 Arthroleptidae Scotobleps gabonicus 0 Arthroleptidae Trichobatrachus robustus 0 Brevicipitidae Balebreviceps hillmani 0 Brevicipitidae Breviceps adspersus 0 Brevicipitidae Breviceps branchi 0 Brevicipitidae Breviceps fuscus 0 Brevicipitidae Breviceps gibbosus 0 Brevicipitidae Breviceps macrops 0 Brevicipitidae Breviceps montanus 0 Brevicipitidae Breviceps mossambicus 0 Brevicipitidae Breviceps namaquensis 0 Brevicipitidae Callulina dawida 0 Brevicipitidae Callulina hanseni 0 Brevicipitidae Callulina kanga 0 Brevicipitidae Callulina kisiwamsitu 0 Brevicipitidae Callulina kreffti 0 Brevicipitidae Callulina laphami 0 Brevicipitidae Callulina meteora 0 Brevicipitidae Callulina shengena 0 Brevicipitidae Callulina sp lowland 0 Brevicipitidae Callulina sp Rubeho 0 Brevicipitidae Callulina stanleyi 0 Brevicipitidae Probreviceps durirostris 0 Brevicipitidae Probreviceps loveridgei 0 Brevicipitidae Probreviceps macrodactylus 0 Brevicipitidae Probreviceps rungwensis 0 Brevicipitidae Probreviceps sp Kigogo 0 Brevicipitidae Probreviceps sp Nguru 0 Brevicipitidae Probreviceps sp Rubeho 0 Brevicipitidae Probreviceps sp Udzungwa 0 Brevicipitidae Probreviceps uluguruensis 0 Brevicipitidae Spelaeophryne methneri 0 Hemisotidae Hemisus marmoratus 0 Hemisotidae Hemisus sudanensis 0 Hyperoliidae Acanthixalus sonjae 0 Hyperoliidae Acanthixalus spinosus 0 Hyperoliidae Afrixalus brachycnemis 0 Hyperoliidae Afrixalus delicatus 0 Hyperoliidae Afrixalus dorsalis 1 0 Hyperoliidae Afrixalus dorsalis 2 0 Hyperoliidae Afrixalus dorsalis 3 0 Hyperoliidae Afrixalus dorsimaculatus 1 0 Hyperoliidae Afrixalus dorsimaculatus 2 0 Hyperoliidae Afrixalus enseticola 0 Hyperoliidae Afrixalus equatorialis 0 Hyperoliidae Afrixalus fornasini 0 Hyperoliidae Afrixalus fulvovittatus 1 0 Hyperoliidae Afrixalus fulvovittatus 2 0 Hyperoliidae Afrixalus knysae 0 Hyperoliidae Afrixalus lacteus 0 Hyperoliidae Afrixalus lacustris 0 Hyperoliidae Afrixalus laevis 0 Hyperoliidae Afrixalus morerei 0 Hyperoliidae Afrixalus nigeriensis 0 Hyperoliidae Afrixalus osorioi 0 Hyperoliidae Afrixalus paradorsalis manengubensis 0 Hyperoliidae Afrixalus paradorsalis paradorsalis 1 0 Hyperoliidae Afrixalus paradorsalis paradorsalis 2 0 Hyperoliidae Afrixalus quadrivittatus 1 0 Hyperoliidae Afrixalus quadrivittatus 2 0 Hyperoliidae Afrixalus spinifrons 0 Hyperoliidae Afrixalus stuhlmanni 0 Hyperoliidae Afrixalus sylvaticus 0 Hyperoliidae Afrixalus vibekensis 0 Hyperoliidae Afrixalus vittiger 1 0 Hyperoliidae Afrixalus vittiger 2 0 Hyperoliidae Afrixalus weidholzi 0 Hyperoliidae Afrixalus wittei 0 Hyperoliidae Cryptothylax greshoffii 1 Hyperoliidae Heterixalus alboguttatus 1 Hyperoliidae Heterixalus andrakata 0 Hyperoliidae Heterixalus betsileo 0 Hyperoliidae Heterixalus boettgeri 0 Hyperoliidae Heterixalus carbonei 0 Hyperoliidae Heterixalus luteostriatus 0 Hyperoliidae Heterixalus madagascariensis 0 Hyperoliidae Heterixalus punctatus 0 Hyperoliidae Heterixalus rutenbergi 0 Hyperoliidae Heterixalus tricolor 1 Hyperoliidae Heterixalus variabilis 1 Hyperoliidae Hyperolius acuticeps 0 Hyperoliidae Hyperolius ademetzi 1 Hyperoliidae Hyperolius adspersus 0 Hyperoliidae Hyperolius argus 1 Hyperoliidae Hyperolius balfouri balfouri 0 Hyperoliidae Hyperolius balfouri viridistriatus 0 Hyperoliidae Hyperolius baumanni 0 Hyperoliidae Hyperolius benguellensis 0 Hyperoliidae Hyperolius bobirensis 0 Hyperoliidae Hyperolius bocagei 1 Hyperoliidae Hyperolius bolifambae 1 Hyperoliidae Hyperolius burgessi 0 Hyperoliidae Hyperolius camerunensis 1 Hyperoliidae Hyperolius castaneus 0 Hyperoliidae Hyperolius chlorosteus 0 Hyperoliidae Hyperolius cinereus 1 Hyperoliidae Hyperolius cinnamomeoventris 1 1 Hyperoliidae Hyperolius cinnamomeoventris 2 1 Hyperoliidae Hyperolius concolor concolor 1 Hyperoliidae Hyperolius concolor guttatus 1 Hyperoliidae Hyperolius concolor ibadanensis 1 Hyperoliidae Hyperolius cystocandicans 0 Hyperoliidae Hyperolius dartevellei 0 Hyperoliidae Hyperolius dintelmanni 1 Hyperoliidae Hyperolius discodactylus 0 Hyperoliidae Hyperolius endjami 1 1 Hyperoliidae Hyperolius endjami 2 1 Hyperoliidae Hyperolius friedemanni 0 Hyperoliidae Hyperolius frontalis 0 Hyperoliidae Hyperolius fusciventris burtoni 1 Hyperoliidae Hyperolius fusciventris ssp 1 Hyperoliidae Hyperolius guttulatus 1 1 Hyperoliidae Hyperolius guttulatus 2 1 Hyperoliidae Hyperolius howelli 0 Hyperoliidae Hyperolius igbettensis 0 Hyperoliidae Hyperolius inyangae 0 Hyperoliidae Hyperolius jackie 0 Hyperoliidae Hyperolius jacobseni 0 Hyperoliidae Hyperolius kivuensis 1 0 Hyperoliidae Hyperolius kivuensis 2 0 Hyperoliidae Hyperolius kivuensis 3 0 Hyperoliidae Hyperolius koehleri 0 Hyperoliidae Hyperolius kuligae 1 Hyperoliidae Hyperolius lamottei 0 Hyperoliidae Hyperolius langi 1 Hyperoliidae Hyperolius lateralis 0 Hyperoliidae Hyperolius lupiroensis 0 Hyperoliidae Hyperolius mariae 0 Hyperoliidae Hyperolius marmoratus marmoratus 1 Hyperoliidae Hyperolius marmoratus swynnertoni 1 Hyperoliidae Hyperolius marmoratus taeniatus 1 Hyperoliidae Hyperolius minitissimus 0 Hyperoliidae Hyperolius mitchelli 0 Hyperoliidae Hyperolius molleri 0 Hyperoliidae Hyperolius montanus 0 Hyperoliidae Hyperolius mosaicus 0 Hyperoliidae Hyperolius nasutus 0 Hyperoliidae Hyperolius nitidulus 1 1 Hyperoliidae Hyperolius nitidulus 2 1 Hyperoliidae Hyperolius occidentalis 1 Hyperoliidae Hyperolius ocellatus ocellatus 1 Hyperoliidae Hyperolius ocellatus purpurescens 1 Hyperoliidae Hyperolius olivaceus 1 Hyperoliidae Hyperolius parallelus alborufus 1 Hyperoliidae Hyperolius parallelus marginatus 1 Hyperoliidae Hyperolius parallelus parallelus 1 Hyperoliidae Hyperolius parallelus pyrrhodictyon 1 Hyperoliidae Hyperolius pardalis 1 Hyperoliidae Hyperolius parkeri 1 Hyperoliidae Hyperolius phantasticus 1 Hyperoliidae Hyperolius picturatus 1 Hyperoliidae Hyperolius pictus 0 Hyperoliidae Hyperolius platyceps 1 Hyperoliidae Hyperolius poweri 0 Hyperoliidae Hyperolius pseudoargus 0 Hyperoliidae Hyperolius pusillus 0 Hyperoliidae Hyperolius quinquevittatus 0 Hyperoliidae Hyperolius raymondii 1 Hyperoliidae Hyperolius reesi 0 Hyperoliidae Hyperolius riggenbachi 1 Hyperoliidae Hyperolius rubrovermiculatus 0 Hyperoliidae Hyperolius rwandae 0 Hyperoliidae Hyperolius semidiscus 0 Hyperoliidae Hyperolius sp nov 1 Hyperoliidae Hyperolius spatzi 1 Hyperoliidae Hyperolius spinigularis 0 Hyperoliidae Hyperolius steindachneri 1 Hyperoliidae Hyperolius stictus 0 Hyperoliidae Hyperolius substriatus 0 Hyperoliidae Hyperolius sylvaticus 1 Hyperoliidae Hyperolius tanneri 0 Hyperoliidae Hyperolius thomensis 0 Hyperoliidae Hyperolius torrentis 1 Hyperoliidae Hyperolius tuberculatus 1 Hyperoliidae Hyperolius tuberilinguis 1 Hyperoliidae Hyperolius veithi 0 Hyperoliidae Hyperolius viridiflavus destefanii 1 Hyperoliidae Hyperolius viridiflavus glandicolor 1 Hyperoliidae Hyperolius viridiflavus ngorongoriensis 1 Hyperoliidae Hyperolius viridiflavus ommatostictus 1 Hyperoliidae Hyperolius viridiflavus pantherinus 1 Hyperoliidae Hyperolius viridiflavus pitmani 1 Hyperoliidae Hyperolius viridiflavus ssp 1 Hyperoliidae Hyperolius viridiflavus variablilis 1 Hyperoliidae Hyperolius viridis 0 Hyperoliidae Hyperolius zonatus 0 Hyperoliidae Kassina arboricola 0 Hyperoliidae Kassina cassinoides 0 Hyperoliidae Kassina cochranae 0 Hyperoliidae Kassina decorata 0 Hyperoliidae Kassina fusca 0 Hyperoliidae Kassina kuvangensis 0 Hyperoliidae Kassina lamottei 0 Hyperoliidae Kassina maculosa 0 Hyperoliidae Kassina schioetzi 0 Hyperoliidae Kassina senegalensis 0 Hyperoliidae Morerella cyanophthalma 1 Hyperoliidae Opisthothylax immaculatus 0 Hyperoliidae Paracassina kounhiensis 0 Hyperoliidae Paracassina obscura 0 Hyperoliidae Phlyctimantis boulengeri 0 Hyperoliidae Phlyctimantis keithae 0 Hyperoliidae Phlyctimantis leonardi 0 Hyperoliidae Phlyctimantis maculata 0 Hyperoliidae Phlyctimantis verrucosus 0 Hyperoliidae Semnodactylus wealii 0 Hyperoliidae Tachycnemis seychellensis 1 Microhylidae Phrynomantis microps 0