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. 2024 Oct 25;44(1):24–35. doi: 10.1002/zoo.21876

Captive Breeding Reveals Insights Into the Ecology and Reproductive Biology of 11 Little‐Known Malagasy Frog Species

Justin Claude Rakotoarisoa 1, Andolalao Rakotoarison 2, Solonirina Rasoanantenaina 1, Eric Robsomanitrandrasana 3, Samina Sidonie Sam Edmonds 1, Jeanne Soamiarimampionona 1, Edupsie Tsimialomanana 1, Sebastian Wolf 1, Devin Edmonds 1,4,5,✉
PMCID: PMC11802484  PMID: 39449579

ABSTRACT

Amphibians are facing an extinction crisis, with ex situ programs increasingly being used as a tool for their conservation. However, conservation efforts are often limited because we do not understand the ecological, behavioral, and life history traits of many amphibian species. Here, we report on the seasonal breeding patterns, egg‐laying behavior, clutch size, and development of 11 frog species maintained at a conservation breeding facility in Andasibe, Madagascar. The frogs exhibited diverse breeding strategies aligned with life history theory. Counting the eggs in 1239 egg masses across these 11 species, we found endotrophic microhylids and terrestrial‐breeding species had the smallest clutch size yet completed metamorphosis quickly, whereas species that laid eggs above or in water with exotrophic larvae had larger clutch sizes and took longer to develop. Most reproduction in captivity occurred during the warm, rainy season and followed seasonal patterns in temperature variation. Yet, Mantidactylus betsileanus bred throughout the year, and Heterixalus betsileo required additional environmental stimuli to trigger reproduction. Notably, we confirmed that Gephyromantis mitsinjo lays eggs on land with tadpoles developing terrestrially within jelly, a behavior previously theorized but which until now remained unobserved. Such observations show how captive breeding programs can be used to gain valuable data on the life history traits of species that are otherwise challenging to observe in nature. Our findings can be used to assess threats to closely related species, helping inform conservation efforts in a country harboring exceptional amphibian species richness and endemism.

Keywords: amphibian, conservation, ex situ, life history, Madagascar

Amphibian conservation efforts are limited by a lack of knowledge about their life history traits. At a conservation breeding facility in Andasibe, Madagascar, we observed diverse breeding patterns and egg numbers among 11 frog species, revealing links between reproductive strategies and environmental factors. These findings provide valuable data for assessing threats to related species and informing conservation in a biodiversity hotspot.

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Summary

  • Captive breeding can reveal ecological insights into poorly known species.

  • We observed diverse reproductive strategies for 11 frog species bred in captivity in Madagascar.

  • These data help to assess threats and inform conservation efforts.

1. Introduction

Over 40% of assessed amphibian species are at risk of extinction, making Amphibia the most threatened vertebrate class (Luedtke et al. 2023). Many species are facing threats that cannot immediately be addressed in nature, requiring ex situ conservation measures to safeguard them from extinction and facilitate reintroductions, population augmentation, and conservation research programs (Zippel et al. 2011; Johnson and Mendelson 2022). Despite the need, only a small proportion of threatened amphibian species are part of conservation breeding programs, with common nonthreatened species dominating zoological collections (Dawson et al. 2016; Jacken, Rödder, and Ziegler 2020). Consequently, ex situ conservation efforts continue to be necessary for the species facing threats that are not currently being mitigated in time to prevent their extinction.

Knowledge about the natural history of a species is often required for enacting ex situ conservation efforts (Michaels, Gini, and Preziosi 2014). However, we know little about the basic ecological traits of most amphibians, especially in the parts of the world harboring the greatest diversity. At the same time, this knowledge gap presents an opportunity for captive breeding programs, as captivity can reveal ecological insights that otherwise would be difficult to observe in nature (Ortiz, Almeida‐Reinoso, and Coloma 2013; Tapley and Girgin 2015; Edwards, Bungard, and Griffiths 2022). Such observations can have important conservation implications. For example, reproductive and larval development modes can be strong indicators of extinction risk (Becker and Loyola 2008; Fontana et al. 2021); yet for many species, we have limited information about their breeding habits.

Madagascar is a biodiversity hotspot with nearly unmatched frog species endemism and richness (Antonelli et al. 2022). Despite the country's rich anuran diversity, its endemic frogs face high extinction risk, with 46.7% of assessed species categorized as threatened by the IUCN (2023). Indeed, amphibians in Madagascar face the most threats per species of any vertebrate class (Ralimanana et al. 2022). Primary threats include habitat loss from subsistence agriculture, logging, and urbanization, as well as climate change, emerging infectious diseases, overcollection for the international pet trade, and invasive species (Andreone et al. 2021). As such, Madagascar is a global priority area for amphibian research and conservation (Nori, Villalobos, and Loyola2018; Nori, Loyola, and Villalobos 2020; Ferreira et al. 2023).

Running captive breeding programs within the native range of target species reduces biosecurity risks, helps garner local support, and can have important political and economic benefits (Gagliardo et al. 2008; Mendelson 2018). Thus, in 2010, the community‐run Malagasy conservation organization Association Mitsinjo began working with the national government and the IUCN Amphibian Specialist Group of Madagascar to establish an amphibian breeding facility in‐country (Edmonds et al. 2012). The initial motivation for the project was to build ex situ conservation capacity in Madagascar ahead of the anticipated threat of the amphibian chytrid fungus Batrachochytrium dendrobatidis (Andreone et al. 2008; Lötters et al. 2011; Bletz et al. 2015). Elsewhere, the emergence of B. dendrobatidis required a rapid response from the ex situ conservation community before there was capacity within local institutions to implement breeding programs (Lee et al. 2006; Poole 2008). Accordingly, we launched Mitsinjo's breeding facility with three objectives: (1) build ex situ conservation capacity in Madagascar, (2) establish captive assurance populations of locally threatened frog species, and (3) conduct captive husbandry research on poorly known species from diverse ecological guilds. Today, the facility serves mainly to sustain a survival assurance colony of the Endangered golden mantella frog (Mantella aurantiaca), with thousands of captive‐bred offspring released into the created habitat to help mitigate the impact of a nearby nickel and cobalt mine (Rakotonanahary et al. 2017; Edmonds et al. 2022).

Here, we report results from maintaining 11 additional Malagasy frog species at Mitsinjo's breeding facility. Specifically, we report novel observations on their breeding phenology, clutch size variability, preferred oviposition sites, and the development stages and timing of metamorphosis. Our results demonstrate how captive breeding can be used to gain valuable insights into the natural history of understudied species, with conservation implications for the threatened species that the common frog species we maintained served as analogs for.

2. Methods

2.1. Study Area

We conducted the study at Association Mitsinjo's amphibian conservation breeding facility in Analamazaotra Forest Station near the village of Andasibe, Madagascar (GPS coordinates: 18.938 S, 48.411 E). The environment is characterized by a warm rainy season during November–March and a cooler winter during April–October, with regional rainfall peaking in the wettest month of January and lowest in September. The area supports exceptional amphibian species richness, with nearly 90 Madagascar‐endemic frog species recorded within the forest station and adjacent Andasibe‐Mantadia National Park (Vieites et al. 2009).

2.2. Species Selection and Collection of Founders

We selected species at the start of the project due to their local abundance, nonthreatened status, and representation of varied ecological guilds (semi‐aquatic, arboreal, phytotelmic species living in water‐filled plant structures, etc.), collecting all founders within Analamazaotra Forest Station. Among the first frogs established in captivity in 2011 were Blommersia blommersae, Boophis pyrrhus, Heterixalus betsileo, and Mantidactylus betsileanus. In May 2013, we added Guibemantis pulcher, collected as an egg mass (33 eggs) found on a Pandanus plant near the breeding facility. In February 2016, we collected additional species for captive husbandry research including Anodonthyla pollicaris, Boophis bottae, Gephyromantis mitsinjo, Guibemantis ambakoana, Platypelis barbouri, and Plethodontohyla mihanika, as well as additional Bl. blommersae (Figure 1). Several of these new species were recommended as analogs for closely related threatened species that may need ex situ conservation assistance in the future (Edmonds et al. 2022; Table 1). After collection, we housed frogs individually in an isolated quarantine room separate from the rest of the breeding facility. Project authorization and collection permits were received from the Ministère de l'Environnement et du Développement Durable (No. 145/MEF/SG/DGF/DVRN/SGFF and No. 30‐12/MEF/SG/DGF/DVRN/SGFF).

Figure 1.

Figure 1

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Eleven frog species bred in captivity during 2011–2021 at the Mitsinjo conservation breeding facility near Andasibe, Madagascar: (A) Platypelis barbouri, (B) Anodonthyla pollicaris, (C) Blommersia blommersae, (D) Boophis bottae, (E) Boophis pyrrhus, (F) Gephyromantis mitsinjo, (G) Guibemantis pulcher, (H) Guibemantis ambakoana, (I) Plethodontohyla mihanika, (J) Heterixalus betsileo, and (K) Mantidactylus betsileanus.

Table 1.

Frog species bred at Mitsinjo's amphibian facility during 2011–2021 (excluding Mantella aurantiaca, see Edmonds et al. 2015) and the potentially threatened species they served as an analog for.

Analog species bred at Mitsinjo's facility Threatened species
Anodonthyla pollicaris (EN) A. emilei (EN), A. nigrigularis (EN), A. theoi (CR)a
Blommersia blommersae (LC) —
Boophis bottae (LC) Bo. ankarafensis (CR), Bo. baetkei (CR)a
Boophis pyrrhus (LC) Bo. feonnyala (EN), Bo. narinsi (EN), Bo. piperatus (EN)
Gephyromantis mitsinjo (LC)b Ge. mafy (CR)a
Guibemantis ambakona (NA) Gu. annulatus (EN), Gu. punctatus (CR), Gu. wattersoni (EN)
Guibemantis pulcher (LC) Gu. annulatus (EN), Gu. punctatus (CR), Gu. wattersoni (EN)
Heterixalus betsileo (LC) —
Mantidactylus betsileanus (LC) M. kortei (EN),b M. noralottae (CR),b M. riparius (EN)b
Platypelis barbouri (LC) Pla. mavomavo (EN),a Pla. olgae (EN)a
Plethodontohyla mihanika (LC) Ple. fonetana (EN), Ple. guentheri (EN)
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Abbreviations: CR, critically endangered; EN, endangered; NA, not assessed by the IUCN Red List and no recommendation for status when described; VU, vulnerable.

a

Threatened species recommended by the 2016 Amphibian Ark Conservation Needs Assessment for Madagascar (https://www.conservationneeds.org). The IUCN Red List category from the most recent assessment is provided in parentheses after the species name.

b

For newly described potentially threatened species that have not been assessed by the IUCN, we note the unofficial Red List category recommended in the species description.

2.3. Captive Environment

We established groups of 4–10 individuals in glass terraria ranging in size from 45 cm × 30 cm × 45 cm to 80 cm × 60 cm × 40 cm (Table 2; Figure 2A–D). All terraria were outfitted with screen covers and drains. The facility's biosecurity protocol limited material inside enclosures to items that could be completely dried and/or chemically disinfected. Pea‐sized gravel or bare‐bottom glass with dried leaves scattered on top were used as substrates. We provided live plants for structure and egg deposition sites, grown in net pots with gravel and sphagnum moss. Leaf litter was collected from the surrounding forest. We washed leaf litter and sphagnum moss in water and allowed them to completely dry in the sun for several days before entering the facility.

Table 2.

Summary of the frog species maintained at the breeding facility, the enclosure size as length x depth x height in centimeters, the number of founders originally collected (M = male; F = female; U = unknown), and their reproductive mode. The number in parentheses following the dimensions of the terrarium is the number of different enclosures housing separate breeding groups originally established by the founders.

Species Tank size (groups) M F U Reproductive mode
Anodonthyla pollicaris 60 × 40 × 47 (1) 4 1 2 Eggs in water in phytotelmata; endotrophic larvae
Blommersia blommersae 45 × 30 × 45 (3) 19 8 0 Eggs laid on plants above lentic water; exotrophic larvae
Boophis bottae 80 × 60 × 40 (1) 8 2 0 Aquatic floating eggs in lotic water; exotrophic larvae
Boophis pyrrhus 80 × 60 × 40 (1) 3 1 0 Aquatic floating eggs in lotic water; exotrophic larvae
Gephyromantis mitsinjo 60 × 40 × 40 (1) 3 1 4 Terrestrial eggs; endotrophic larvae
Guibemantis ambakoana 45 × 30 × 45 (2) 4 3 0 Eggs laid on leaves of Pandanus plants; larvae in phytotelmata
Guibemantis pulcher 45 × 30 × 45 (2) 9 7 0 Eggs laid on leaves of Pandanus plants; larvae in phytotelmata
Heterixalus betsileo 45 × 30 × 45 (1) 3 3 0 Aquatic floating eggs in lentic water; exotrophic larvae
Mantidactylus betsileanus 60 × 40 × 47 (2) 7 4 0 Terrestrial eggs laid near water; exotrophic larvae
Platypelis barbouri 60 × 40 × 47 (1) 4 3 2 Eggs in water in phytotelmata; endotrophic larvae
Plethodontohyla mihanika 60 × 40 × 40 (1) 3 3 0 Eggs in water in phytotelmata; endotrophic larvae
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Figure 2.

Figure 2

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Representative enclosures used to maintain frogs at the breeding facility in Madagascar. The terrariums in the photos housed: (A) Guibemantis pulcher, (B) Mantidactylus betsileanus, (C) Plethodontohyla mihanika, and (D) Boophis pyrrhus.

All enclosures had a water source. For M. betsileanus, we provided a water area that took up 30%–50% of the terrarium footprint. For other species, we used a water dish which was changed daily. Water was sourced from the Ambatomandondona Dam inside Andasibe National Park. The pH of incoming water ranged from 6.0 to 7.2, measured with a colorimetric aquarium test kit. We heated incoming water with a solar water heater before entering the facility to reduce the risk of introducing B. dendrobatidis, and then cooled the water in a holding tank before use.

In addition to gravel, potted plants, and a water area, we provided refugia for terrestrial frogs in the form of broken terra cotta flowerpots, PVC plastic pipe segments cut in half, and coconut huts. For species known to reproduce in tree holes, we provided bamboo ~4–6 cm in diameter, film canisters suction‐cupped to the side of the enclosure, and/or a wooden branch with a hole drilled in the top. To better access the inside of bamboo tubes, which proved useful breeding sites for A. pollicaris, Pla. barbouri, and Ple. mihanika, we cut out a small door fixed with an elastic band (Figure 2C). For the two Guibemantis species, which are phytotelmic and live in Pandanus screw pines, we provided plants with a similar structure to Pandanus species, including bromeliads in the genus Aechmea, Callisia fragrans, and artificial bromeliads made by Exo Terra. Segments of PVC plastic pipe positioned at angles leaning against the glass served as perches for arboreal species. Every 1–2 months, we removed frogs from their enclosures, wiped down the sides of the terrariums, and flushed the tanks with water to prevent waste buildup.

No supplemental heating or cooling was provided because all species were native to the forest surrounding the facility. Thus, the ambient temperature inside the building was comparable to outside. However, we used large fans to circulate air in the frog room during the warmest part of the day. A natural photoperiod was provided with ambient light through the windows, along with supplemental fluorescent tubes placed above the terraria and set on timers for 11–12 h per day. During 2011–2012, we did not have reliable electricity at the breeding facility so only ambient lighting from windows regulated photoperiod.

2.4. Feeding Regimens

We fed frogs cultured live foods from Andasibe, mostly fruit flies and crickets (five different species; see Edmonds et al. 2012). Smaller frogs were also fed collembolans. We occasionally collected earthworms outside the facility to supplement the diet of M. betsileanus when crickets were running low. We fed frogs two to four times per week, offering more food during the warm rainy season and smaller amounts during the winter. Juvenile frogs were fed daily. We coated fruit flies with nutritional supplements before feeding, usually Repashy Calcium Plus or Herpetal Amphib. Crickets were fed a range of seasonally available fruits and vegetables along with ground dry freshwater shrimp as a protein source to improve their nutritional value (Finke 2003).

2.5. Tadpole Care

Tadpole care techniques varied by reproductive mode and larvae type. We fed the exotrophic tadpoles (tadpoles that rely on an external food source rather than yolk reserves) of Bl. blommersae, Bo. bottae, Bo. pyrrhus, H. betsileo, and M. betsileanus a mixture of spirulina powder, ground dried freshwater shrimp, and aquarium fish foods (Tetra Min and Bassleer Biofish Food brands). We also experimented with feeding dried greens to M. betsileanus (see Soamiarimampionona et al. 2015). The endotrophic (nonfeeding) microhylids (A. pollicaris, Pla. barbouri, and Ple. mihanika) and Ge. mitsinjo (which has no free‐swimming tadpole), did not require food during the larval stage. Tadpoles were raised either in glass aquariums or plastic bins. Lentic species that breed in nonflowing water like Bl. blommersae were raised in groups of up to around 30 individuals in 16‐L bare‐bottom plastic bins filled with 10–14 L of water. The stream‐breeding Boophis species were maintained in groups of up to 40 (though usually < 10) within 22‐L glass aquaria equipped with sponge filters powered by air pumps for mechanical filtration. We changed 50%–80% of the water in aquariums one to two times per week and spot‐cleaned waste daily using a turkey baster. The water temperature typically ranged 16°C–22°C, depending on the time of year. The microhylids were allowed to develop in the enclosures of adults until metamorphosis, whereas for Ge. mitsinjo, we removed the eggs when found and placed them on gravel and moist sphagnum moss in a plastic cup until they completed metamorphosis within egg jelly.

2.6. Data Collection and Analyses

We checked terrariums daily for eggs as part of our regular maintenance routine. To standardize the data collection process, we used a paper form with the following fields: species, date eggs found, date eggs counted, location eggs found, number of eggs, and number of fertile eggs. In addition to documenting reproductive events, we also recorded information related to egg development such as the number of tadpoles that hatched from an egg mass, the date the first tadpole hatched from an egg, the number of tadpoles that died before metamorphosis, the date the first tadpole completed metamorphosis, the date the last tadpole completed metamorphosis, and the total number that completed metamorphosis. Additionally, during 2018–2021, we also recorded the minimum and maximum 24‐h air temperature using a digital thermometer, which was checked each morning and its memory cleared after.

Most egg masses were culled to prevent captive populations from exceeding the capacity of the breeding facility. To stop egg development, we placed egg masses in a 5% ethanol solution and then gradually added 70% or 90% ethanol with a pipette once we counted the number of eggs that were fertile. We considered eggs fertile if we observed a division of the ovum and culled them before stage 17 in Gosner (1960).

While compiling data on clutch size, we excluded two egg masses from Bl. blommersae that numbered 208 and 222 eggs and appeared to be multiple egg masses from different females. We also excluded 10 M. betsileanus egg masses numbering < 8 eggs that were not fertile and may have been leftover eggs in the terrarium from a previous spawning. While compiling data on time to metamorphosis, we omitted all tadpoles from M. betsileanus egg masses used in a pilot study on a captive diet because several tadpoles took more than 1 year to finish metamorphosis (presumably, they were not provided with optimal conditions, see Soamiarimampionona et al. 2015). We entered data into Microsoft Access and summarized them for visualization using package tidyr in R version 4.3.2 (Wickham et al. 2019; R Core Team 2022).

3. Results

3.1. Reproduction and Breeding Phenology

We recorded data on 1272 total captive breeding events from all 11 frog species. We had the most success breeding M. betsileanus (86.7% of all egg masses recorded), which bred consistently throughout all months of the year. Most other species exhibited highly seasonal reproduction (Figure 3), though Gu. ambakoana laid a small number of infertile egg masses during the cooler months of June–September. Three species (Bo. bottae, Gu. pulcher, and H. betsileo) only spawned one time each, all during the warm rainy season. However, the eggs from Gu. pulcher were infertile. Notably, after 6 years of maintaining H. betsileo without breeding success, we placed animals in a rain chamber made from a plastic box, aquarium pump, and irrigation hose, and soon found fertilized eggs. All other species bred without immediate environmental stimuli. Instead, breeding aligned with the natural ambient seasonal variation in temperature (Figure 3).

Figure 3.

Figure 3

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Total number of monthly egg masses for eight species during 2018–2021 overlayed with the monthly mean high (red points) and mean low (blue points) temperature in the facility. The error bars on the points are +/− 1 standard deviation. Mantidactylus betsileanus was excluded from the plot because the large number of breeding events overshadowed all other species. Boophis bottae and Guibemantis pulcher are not shown because they were only bred once in 2017 before we began consistently recording daily minimum and maximum temperatures.

3.2. Egg Mass Variability

We counted the number of eggs in 1239 egg masses (Table 3; Figure 4). Ge. mitsinjo had the smallest mean clutch size, followed by the microhylids and Guibemantis species (Figure 4). Species that spawned in water, such as Bo. bottae, Bo. pyrrhus, and H. betsileo, laid the most eggs. Bl. blommersae and Bo. pyrrhus exhibited the greatest within‐species clutch size variability (Figure 4).

Table 3.

Percentage of eggs fertilized during captive breeding events from April 2011 to May 2021. To prevent captive populations from exceeding the capacity of the facility, we regularly stopped egg development by placing the eggs in alcohol after counting the number of fertilized eggs. Clutches represent the number of egg masses for which we counted all eggs. Saved is the number of clutches that were allowed to develop into tadpoles after noting whether the eggs were fertile or not. Total is the total number of eggs in all clutches. Fertile is the total number of fertilized eggs. Percent is the Total column divided by the Fertile column × 100. Offspring is the total number of tadpoles that completed metamorphosis from all breeding events for the species.

Species Clutches Saved Fertile Total Percent Offspring
Anodonthyla pollicaris 25 3 554 731 75.8 151
Blommersia blommersae 49 1 1669 3753 44.5 0
Boophis bottae 1 1 113a 336 33.6a 8
Boophis pyrrhus 18 5 698 1486 47.0 72
Gephyromantis mitsinjo 52 6 166 509 32.6 42
Guibemantis ambakoana 19 0 20 371 5.4 0
Guibemantis pulcher 1 — 0 33 0.0 0
Heterixalus betsileo 1 1 33a 121 27.3a 10
Mantidactylus betsileanus 1054 20 22,446 35,780 62.7 79
Platypelis barbouri 10 1 38 306 12.4 24
Plethodontohyla mihanika 9 5 332 473 70.2 131
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a

We did not count the number of fertilized eggs but instead the number of tadpoles that hatched.

Figure 4.

Figure 4

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Density plots of eggs per egg mass by species to visualize clutch size variability. Boophis bottae, Guibemantis pulcher, and Hetreixalus betsileo were only bred once, and data are not included in this figure.

3.3. Oviposition Substrates and Locations

Egg masses were laid in different locations and on different substrates depending on species (Figures 5 and 6). The two Boophis species and H. betsileo laid eggs in water, except for one egg mass from Bo. pyrrhus which we found partially adhered to the glass of the terrarium. Terrestrial and semi‐aquatic frogs used a range of locations. Bl. blommersae attached most eggs to the underside of live plant leaves, and Ge. mitsinjo and M. betslieanus deposited most eggs on land under refugia such as dried leaves, pieces of terra cotta flowerpots, or half‐cut PVC plastic pipe segments. More than half of Gu. ambakona egg masses were laid on the terrarium floor under leaf litter rather than on plants used to mimic Pandanus screw pines. We found a single infertile egg mass from the closely related Gu. pulcher on the glass bottom of the terrarium. The three microhylids bred exclusively within water‐filled holes, most often segments of bamboo, but also film canisters attached to the side of the terrarium with a suction cup. The eggs of the microhylids were laid in the water, and the males of all three species stayed with the eggs and larvae in the holes through metamorphosis.

Figure 5.

Figure 5

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Egg masses and developing larvae of frogs bred in captivity at the breeding facility in Madagascar: (A) Blommersia blommersae, (B) Boophis pyrrhus, (C) Guibemantis ambakoana (infertile), (D) adult male Plethodontohyla mihanika inside bamboo with metamorphosing juveniles, (E) Gephyromantis mitsinjo, (F) Mantidactylus betsileanus, (G, H) Anodonthyla pollicaris, and (I) Platypelis barbouri adult frogs with eggs inside a film canister.

Figure 6.

Figure 6

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Location of egg masses as a proportion of total egg masses for each species. Glass = the glass of the terrarium, either on the side or bottom where there was no substrate; Gravel = small stones about 1 cm in diameter, which served as a substrate in enclosures; Hole = inside a film canister, piece of bamboo, or hole carved into wood; Leaf = on the leaf of a live plant; Refuge = under a shelter such as a coconut hut, half‐cut PVC plastic pipe, or broken terra cotta flowerpot; Water = in a water dish or water area of the terrarium. The sample size n at the bottom of each bar is the number of egg masses with a recorded location.

3.4. Larval Development and Time to Metamorphosis

The eggs of A. pollicaris, Pla. barbouri, and Ple. mihanika took a relatively long time to hatch into tadpoles yet completed metamorphosis rapidly (Figures 7 and 8). Pla. barbouri completed metamorphosis in as little as 17 days after the eggs were laid, and A. pollicaris and Ple. mihanika in 20 days (Figure 8). Ge. mitsinjo, which does not have an aquatic larval stage (Figure 9), also developed quickly, taking a minimum of 22 days to complete development. In contrast, some slow‐developing individual Bo. pyrrhus tadpoles required nearly 300 days to complete metamorphosis, with the fastest‐developing Bo. pyrrhus tadpole still taking 126 days (Figure 8). Several exceptionally slow M. betsileanus tadpoles also took a long time (nearly 300 days) to complete metamorphosis, though most M. betsileanus completed metamorphosis in < 4 months.

Figure 7.

Figure 7

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Mean number of eggs per egg mass by species +/− 1 standard deviation (top) and the median minimum number of days until a tadpole breaks free from the egg mass +/− the minimum and maximum (bottom). The sample size n is the number of egg masses where eggs were counted (top) and where the date of hatching was recorded (bottom). The sample size and species on the x‐axis vary between plots because we did not record days to hatching for all species or all egg masses found.

Figure 8.

Figure 8

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Range in days to metamorphosis from the date eggs were found in terrariums until the last tadpole in a clutch had completely absorbed its tail. The sample size is the total number of frogs that completed metamorphosis for each species and where we recorded the first and last date of metamorphosis for the clutch. Note the sample size for Mantidactylus betsileanus here differs from that in Table 1 because for one egg mass in 2013, we recorded the dates at which tadpoles completed metamorphosis but not the number of eggs or number fertilized. †No aquatic larval stage.

Figure 9.

Figure 9

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The tadpoles of Gephyromantis mitsinjo developing in egg jelly on land.

4. Discussion

To our knowledge, of the 11 frog species we report on, only Bo. pyrrhus and M. betsileanus have been described breeding in captivity previously (Scheld et al. 2013; Soamiarimampionona et al. 2015; Edmonds et al. 2016). Institutions outside Madagascar have also maintained Gu. pulcher and H. betsileo, though without recorded breeding success (Vences, Hauswaldt, and Glaw 2011; Peš and Peš 2022; Ziegler et al. 2022). The only other reproductive data available are from fieldwork, where one or two egg masses of the species we bred or a close relative were discovered incidentally (Blommers‐Schlösser 1975, 1979; Vences et al. 2003; Fenolio et al. 2007). Among our most notable observations is that Ge. mitsinjo lays eggs terrestrially with nonfeeding tadpoles hatching from eggs and developing on land within egg jelly. Randrianiaina et al. (2011) first noted tadpoles remaining in egg jelly in Ge. feomborona (as Ge. aff. blanci) from an egg mass found in the field near a calling male. Here, we confirm Ge. mitsinjo exhibits the same reproductive strategy. Taken together, our results highlight how captive programs for amphibians are a valuable tool for gaining knowledge about breeding frequency, fecundity, larval development, and related life history traits of poorly known amphibian species.

The short duration of metamorphosis and small clutch size observed in species with nonfeeding tadpoles agrees with amphibian life history patterns. The tree‐hole breeding microhylids and Ge. mitsinjo completed metamorphosis quicker than other feeding tadpole species. As small water‐filled tree holes and terrestrial habitats are prone to desiccation, the rapid development of nonfeeding tadpoles reflects a reproductive strategy that minimizes risk in an unpredictable environment (Duellman 1989; Rudolf and Rödel 2007). Relatedly, our observation of Ge. mitsinjo laying the fewest eggs per mass aligns with their mode of reproduction; terrestrial‐breeding species tend to lay small numbers of eggs (Furness et al. 2022). Another common trait of frogs with endotrophic larvae is parental care (Altig 2003). Indeed, we observed egg‐guarding in male A. pollicaris, Pla. barbouri, and Ple. mihanika, a behavior also recorded in the field (Blommers‐Schlösser 1975; Vences et al. 2003). However, we did not document additional parental care behaviors as has been theorized for Madagascar's tree‐hole breeding microhylids. Future research in captivity could remove adults from larvae or eggs and monitor development to further explore questions about parental care considering Blommers‐Schlösser (1975) noted eggs molding in the field after collecting an egg‐guarding male from a hole.

While our observations of reproduction in the microhylids and Ge. mitsinjo might have been expected, the oviposition sites chosen by the Guibemantis species were surprising. The two Guibemantis we maintained are phytotelmic, living and reproducing within the leaf axils of Pandanus screw pines (Glaw and Vences 2007; Gabriel et al. 2024). However, we found more than half of the egg masses on the glass bottom of the tank or under leaf litter. One likely explanation is that the captive environment we provided did not have adequate egg laying sites. The single fertile clutch from Gu. ambakoana was laid in the leaf axil of an artificial bromeliad made of plastic. Considering all but one egg mass was infertile, we suggest continued captive husbandry research into frogs of the subgenus Pandanusicola.

The founders of the captive populations were identified based only on morphology. In fact, several of the species we bred have only recently been described. As such, without genetic verification, some of our observations could conceivably correspond to other morphologically similar sympatric species. For example, Ple. mihanika is told apart from Ple. notosticta by having hind legs that extend past the nostril when bent forward (Vences et al. 2003), but this is challenging to evaluate in live animals. Similarly, M. betsileanus and M. katae are only distinguishable by femoral gland size and call (Scherz et al. 2022), and as such some of the female frogs at the breeding facility could have plausibly been M. katae. Furthermore, Gu. ambakoana, Gu. rianasoa, and Gu. methueni, as well as Bl. blommersae and Bl. sarotra, are superficially similar in coloration (Glaw and Vences 2002; Gabriel et al. 2024). Breeding programs in Panama have found cryptic species diversity within captive populations using molecular techniques (Crawford et al. 2013). Should captive survival assurance populations be needed for the threatened species our frogs served as analogs for, we recommend founders be genetically identified.

Temperature is one of the main exogenous stimuli driving seasonal reproductive cycles (Oseen and Wassersug 2002). Yet, aside from Xenopus laevis (Kalk 1960), most research on amphibian breeding cues has focused on species from temperate regions where seasons are more pronounced (Borah, Renthlei, and Trivedi 2019). The species we bred reproduced mainly during December–March, which agrees with field observations of peak amphibian activity and temperatures in Andasibe (Heinermann et al. 2015). Interestingly, this pattern contrasts with the breeding behavior of some closely related taxa at lower elevations. For example, A. boulengeri is known to breed in the winter month of July near Toamasina (Blommers‐Schlösser 1975) and Anodonthyla spp. calls can be heard year‐round in Madagascar's lowland forests, leading to the belief the genus is not seasonal in its reproduction (Vences et al. 2010). However, captive A. pollicaris exposed to natural seasonal temperature variation in Andasibe bred mainly during December–May. In contrast, M. betsileanus bred every month of the year with no apparent seasonal pattern. Temperature influences the breeding cycles of many species, but others may be triggered by additional stimuli, especially rainfall (Green 2017, Güell and Warkentin 2023). For example, even after years of exposure to seasonal temperature changes, H. bestileo required the added environmental stimuli of a rain chamber, highlighting how some frog species need immediate cues to act as behavioral triggers in captivity on top of seasonal temperature cycles (Linhoff, Germano, and Molinia 2022).

Increasingly, life history traits are being used to assess threats to data‐deficient amphibian species and identify priorities for conservation (Crump 2015; González del Pliego et al. 2019). Our findings of oviposition sites and reproductive modes can contribute to threat assessments for understudied closely related taxa. For example, we made a second observation of tadpoles hatching from eggs but remaining in egg jelly within the nominate Gephyromantis subgenus, a group of frogs for which reproduction has never been observed before. Terrestrial breeding amphibian species are especially sensitive to the threat of climate change because their eggs and larvae are prone to desiccation and decreased precipitation reduces soil moisture (Cummins et al. 2019). Accordingly, data‐deficient species related to Ge. mitsinjo like Ge. mafifeo and threatened species like Ge. boulengeri, Ge. cornucopia, and Ge. mafy (Miralles et al. 2023) may well be at risk because they likely breed the same way. In addition to assessing threats, our results can be used to inform conservation strategies through population modeling, which requires estimates of fecundity (Schwartz et al. 2017; Zipkin and Saunders 2018). However, we lack data on egg number variability for most Malagasy frog species. The data provided here on clutch size can be used as a starting point for population models of threatened taxa when species‐specific fecundity estimates are unavailable.

In many ways, amphibians are ideally suited for ex situ conservation programs but limited knowledge about their natural history often inhibits program success (Michaels, Gini, and Preziosi 2014, Tapley et al. 2015). We selected several species to serve as analogs for learning about threatened taxa, yet just because a species is threatened does not make it well‐suited for a conservation breeding program. Factors like an institution's geographic proximity to the native range, biosecurity risks, and knowledge of the species' biology are all important to assess (Carrillo, Johnson, and Mendelson 2015; Bradfield, Tapley, and Johnson 2023). Increasingly, zoological institutions are using field research to inform the captive care of species in ex situ programs, but as we showed, the connection between in situ and ex situ also works in reverse. By studying the ecological traits and behavior of poorly known species in captivity, we can gain valuable information for assessing threats, modeling populations, and informing conservation.

Ethics Statement

Project authorization and collection permits were received from the Ministère de l'Environnement et du Développement Durable (No. 145/MEF/SG/DGF/DVRN/SGFF and No. 30‐12/MEF/SG/DGF/DVRN/SGFF). The study adheres to the ARRIVE guidelines and applies to national legislation in Madagascar.

Acknowledgments

We are grateful to the many individuals who helped us with the project, especially R. Dolch, H. Kurrer, R. Gagliardo, A. Pessier, and J. Pramuk, and to volunteers including S. Boks, L. Harding, J. Heinermann, S. Sutor, G. Strait, and M. Ward. We thank J. E. A. Fanirihasimbolatiana, J. D. Rabarinirina, and J. B. Lahinirina for their work producing live foods and organizing data. Staff from Woodland Park Zoo, Amphibian Ark, Durrell Wildlife Conservation Trust, Colchester Zoo, and Chester Zoo kindly ran training workshops in Andasibe for staff. Jersey Zoo, J. Dawson, and Toledo Zoo facilitated training abroad. F. Rabemananjara, N. Rabibisoa, C. Randrianantoandro, and other Amphibian Specialist Group members offered helpful input on study design and data collection methods during project assessment trips. We thank H. Gabriel for offering a second opinion on the identification of the Guibemantis species, as well as F. Andreone, A. Crottini, S. Gehring, J. Köhler, F. Glaw, J. Glos, C. Hutter, R. Lehtinen, A. Raselimanana, C. Raxworthy, J. Reimann, G. Rosa, M. Scherz, M. Vences, and D. Vallan for participating in the 2015 Amphibian Ark Conservation Needs Assessment to identify analog species. Financial and material support during 2010–2021 were received from Ambatovy Minerals S.A., the Amphibian Ark Seed Grant, the Association of Zoos and Aquariums Conservation Endowment Fund, American Frog Day, BIOPAT—Patrons for Biodiversity, the Cleveland Zoological Society Africa Seed Grant, Conservation International, Dendrobatidae Nederland, Deutsche Gesellschaft für Herpetologie und Terrarienkunde, Durrell Wildlife Conservation Trust, Josh's Frogs, Thoiry Zoo, Toronto Zoo, Tree Walkers International, Understory Enterprises, and Woodland Park Zoo.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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Associated Data

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Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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