Delayed Trait Development and the Convergent Evolution of Shell Kinesis in Turtles Gerardo A

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10-3-2018 Delayed trait development and the convergent evolution of shell kinesis in turtles Gerardo A. Cordero Iowa State University

Kevin Quinteros Iowa State University, [email protected]

Fredric J. Janzen Iowa State University, [email protected]

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Abstract Developmental processes are foundational to clarifying the causes of convergent evolution. Here, we show how a key convergently evolving trait is slowly “acquired” in growing turtles. Adaptive morphological change tends to originate late in turtle ontogeny, owing to design constraints imposed by the shell. We investigated this trend by examining derived patterns of shell formation associated with the multiple (≥ 8) origins of shell closure (kinesis) in smallbodied turtles. Using box turtles as a model, we demonstrate that the flexible hinge joint required for shell kinesis differentiates gradually and via extensive repatterning of shell tissue. Disproportionate changes in shell shape and size substantiate that this transformation is a delayed ontogenetic response (3-5 years post-hatching) to structural alterations that arise in embryogenesis. These findings exemplify that the translation of genotype to phenotype may reach far beyond embryonic life stages. Thus, the temporal scope for developmental origins of adaptive morphological change might be broader than generally understood. We propose that delayed trait differentiation via tissue repatterning might facilitate phenotypic diversification and innovation that otherwise would not arise due to developmental constraints.

Disciplines Animal Sciences | Ecology and Evolutionary Biology | Genetics | Population Biology

Comments This is a manuscript of an article published as Cordero, Gerardo A., Kevin Quinteros, and Fredric J. Janzen. "Delayed trait development and the convergent evolution of shell kinesis in turtles." Proc. R. Soc. B 285 (2018): 20181585. doi: 10.1098/rspb.2018.1585. Posted with permission.

This article is available at Iowa State University Digital Repository: https://lib.dr.iastate.edu/eeob_ag_pubs/305

Convergent phenotypes develop slowly in turtles with shell kinesis

Gerardo Antonio Corderoa*, Kevin Quinterosa, Fredric J. Janzena

a Department of Ecology, Evolution, and Organismal Biology, 251 Bessey Hall; Phone: 515

294-3522; Fax: 515 294-1337

Corresponding Author: Gerardo Antonio Cordero; Email: [email protected]

Summary

Developmental processes are foundational to clarifying the causes of convergent evolution.

Here, we show how a key convergently evolving trait is slowly “acquired” in growing turtles.

Adaptive morphological change tends to originate late in turtle ontogeny, owing to design constraints imposed by the shell. We investigated this trend by examining derived patterns of shell formation associated with the multiple (≥ 8) origins of shell closure (kinesis) in small- bodied turtles. Using box turtles as a model, we demonstrate that the flexible hinge joint required for shell kinesis differentiates gradually and via extensive repatterning of shell tissue.

Disproportionate changes in shell shape and size substantiate that this transformation is a delayed ontogenetic response (3-5 years post-hatching) to structural alterations that arise in embryogenesis. These findings exemplify that the translation of genotype to phenotype may reach far beyond embryonic life stages. Thus, the temporal scope for developmental origins of adaptive morphological change might be broader than generally understood. We propose that delayed trait differentiation via tissue repatterning might facilitate phenotypic diversification and innovation that otherwise would not arise due to developmental constraints. 1. Introduction

Similar environmental selective pressures often lead to similar adaptive phenotypes in species that do not share a recent common ancestor, i.e. convergent evolution [1-4]. This fascinating trend provides an opportunity to evaluate the extent to which evolution is predictable and repeatable [3, 4]. Indeed, major advances in formulating a theoretical framework for the study of convergent evolution were achieved in recent years [5, 6].

However, hypotheses targeting the role of developmental processes are rarely addressed (but see [7-10]).

Although the primacy of natural selection is unequivocal in explaining convergent evolution, developmental processes may ultimately limit the number of evolvable character states [3]. For instance, the likelihood of trait convergence might be higher in species groups that exhibit morphological stasis owing to design limitations imposed by underlying developmental processes [11, 12]. The atypical turtle body plan is an outstanding model to examine such a phylogenetic pattern [13, 14].

The turtle’s shell forms via changes in the skeletal architecture of tetrapods that have profoundly influenced turtle diversification over the last 210 million years [15, 16]. As a consequence of shell development, turtles are the only tetrapod to feature bone sutures in the thoracic region analogous to the ones commonly found in the cranium [17]. These sutures have been co-opted repeatedly to give rise to an assortment of functional shell adaptions [15,

18-20]. Beginning in the early Cretaceous, hinge sutures that enable shell closure via muscle- induced movement of the shell (i.e. kinesis) have evolved independently in multiple lineages

[15, 21-29]. Adaptive hypotheses on this convergence generally invoke enhancement of antipredator defense in terrestrial habitats or in shallow water (Fig. 1) [23, 29-34], as well as avoidance of dehydration while in terrestrial burrows [35]. Shell kinesis comprises a suite of musculoskeletal traits that develop in a taxon- specific manner [16, 23-25, 27, 36]. Even so, gradual differentiation of one or two hinge joints during post-embryonic stages is common to all kinetic-shelled lineages [37-39]. To explain delayed hinge differentiation, morphologists proposed that boundaries of ectodermal plates (i.e. scutes) and underlying bones must align as these shell elements grow and shift their positions (Fig. 1) [23, 40, 41]. Furthermore, mechanical strain exerted by muscles during shell closure might prevent sutural fusion of adjacent bones, leading to the formation of elastic connective tissue [23, 41]. This complex transformation, however, has not been examined. In general, post-embryonic tissue changes are challenging to describe in long-lived organisms, e.g. humans [42].

We investigated developmental processes underlying the convergence of shell hinges in turtles with shell kinesis. We first quantified evolutionary origins of kinesis and tested the hypothesis that small-bodied turtles with terrestrial habits are more likely to feature kinetic hinges [34, 43, 44]. Then, using box turtles (Terrapene) as a model, we tested the prediction that fusion of a precursor hinge suture is delayed and elastic tissue forms as kinesis becomes functional in juveniles [23], hereafter ‘heterochronic’ model. Alternatively, this suture might fuse and undergo repatterning in conjunction with elastic tissue formation, hereafter

‘repatterning’ model. In either case, we expected associated shell structures (scutes and buttresses) to also undergo repatterning [23, 37]. Lastly, we tested whether kinetic shells exhibit disproportionate size and shape changes during ontogeny [29, 43].

2. Materials and methods

(a) Comparative phylogenetic analyses

We performed comparative analyses in the phytools R package [45]. Using stochastic character mapping [46], we reconstructed ancestral character states of shell kinesis on the most recent molecular phylogeny of turtles (N = 292 species) [47]. We simulated character state histories 1,000 times (along branches and on nodes) with sampling conditioned on an equal-rates model (Markov k-state 1) for state transitions. We then computed simulated averages for absent-to-present and present-to-absent transitions to evaluate hypothetical independent origins of shell kinesis, as well as potential reversals.

We tested the hypothesis that body size, i.e. carapace length (CL), is shorter in kinetic- shelled species with a simulation-based phylogenetic analysis of variance (pANOVA) [48].

We evaluated the correlation of habitat type (aquatic/terrestrial) and shell type

(akinetic/kinetic) with Pagel’s test for discrete characters [49], assuming that transitions in one character depended on the other, and vice versa, and that all rates for state transitions were different. To do so, we optimized model fitting with the geiger ‘fitDiscrete’ function and ran two separate Pagel’s tests: one based on the standard aquatic/terrestrial turtle dichotomy

[44], and another that included semiaquatic Kinosternon as terrestrial (following refs. [50-

52]). All character state data appear in Table S1 of the electronic supplementary material.

(b) Shell tissue preparation

We prepared samples following standard histological protocols for bone tissue [53].

We first fixed hatchling carcasses, obtained from a previous study [16], in 10% buffered formalin before dehydrating them in an increasing ethanol series. We stained tissue of representative lineages with the most common forms of plastral (ventral shell) kinesis in alizarin red to confirm the absence of hinges. We then focused on emydid lineages with plastral kinesis–Terrapene ornata and Emys (Emydoidea) blandingii [16]–and akinetic- shelled Chrysemys picta and Glyptemys insculpta. See the electronic supplementary material for further details on specimens used. We skeletonized preserved adult C. picta, E. blandingii, and T. ornata in the ISU herpetological collection to be scanned with a NextEngine 3D surface scanner to compare 3D models of shell morphology. We also dissected formalin-fixed plastron tissue and stained it in hematoxylin and eosin solution to examine immature sutures in hatchling T. ornata. We further decalcified adult plastron tissue in 5% formic acid and stained it with Verhoeff-van

Gieson solution to compare elastic fiber variation in sutures of T. ornata versus G. insculpta.

To ascertain whether disproportionate plastron growth occurs in kinetic-shelled species, we examined log-transformed measurements of plastron length (PL) and CL in live- trapped adults and juveniles of C. picta, E. blandingii, and T. ornata from Thomson, Illinois

[54]. Using digital calipers, we recorded shell measurements in embryos (stages 17-22) and hatchlings preserved in previous studies [16, 55]. We evaluated PL variation in relation to CL with a general linear model, which included a species by PL interaction term to account for interspecific variation in PL growth rates. The plastron may occlude the carapace once the plastral hinge is functional [23], thus we expected a 1:1 PL-to-CL relationship in adults of kinetic-shelled species. In emydid turtles, PL/CL generally approaches one (i.e. isometry)

[56], thus an ANOVA was suitable to test mean differences in this ratio [57].

Following Myers et al. [58], we quantified post-hatching plastron shape by digitizing

12 fixed homologous landmarks in a subsample of museum specimens. We placed landmarks in the same position in all photographed specimens (with a ruler for scaling) to minimize distortion using tpsDig [59]. We then performed generalized Procrustes analysis to remove non-shape variation using tpsRelw [60]. For comparison, we first superimposed landmarks and translated them to a shared origin, followed by rescaling to units of centroid size. We subsequently rotated them to minimize the sums-of-squares differences among all landmark configurations [61]. After orthogonal projection into a linear tangent space, aligned Procrustes shape coordinates depicted shape variation, which we represented as thin-plate spline deformation grids using tpsSpline [62].

We explored shape variation with a principal component analysis and regressed the first PC axis against log PL to test whether plastron deformation increased disproportionately during post-hatching growth in kinetic-shelled emydids. We evaluated interspecific differences in shape deformation, represented by the first PC axis, with a general linear model

(as above). We conducted these statistical analyses using base functions of R [63].

3. Results

(a) Multiple origins of shell kinesis

On average 9.30 shell kinesis absent-to-present transitions arose in Pelusios, Lissemys,

Kinosternidae, Emys, Terrapene, Kinixys, Cyclemys, and Cuora (95% HPD = 8-11; 1,000 simulations) (Fig. 1d; Fig. S1). In contrast, only 0.77 present-to-absent transitions were indicated (95% HPD = 0-3) (Fig. S1). The presence of shell kinesis was positively correlated with habitat terrrestriality in a model that included semiaquatic Kinosternon as terrestrial

(Pagel’s correlation test: P = 0.0009, likelihood ratio = 18.6), otherwise the correlation was not supported (P = 0.324; likelihood ratio = 4.66) (Fig. 1d). Further, species with shell kinesis were smaller (CL: 206.5 mm ± 9.07 SE, N = 68) than akinetic-shelled species (CL: 441.8 mm

± 23.7 SE, N = 223) (pANOVA: F = 63.4, P = 0.008; permutations = 1,000) (Fig. 1d-e).

(b) Repatterning of shell tissue

By hatching, a plastral hinge is absent in lineages representing diverse types of plastral kinesis (Emydidae, Kinosternidae, Pelomedusidae; Fig. S2). In T. ornata, a proper hyoplastral-hypoplastral bone suture forms by 3 yr post-hatching in the location of the incipient hinge joint (Fig. 2a-b). However, this suture is progressively repatterned as the plastral hinge becomes functional 3-5 yr post-hatching: interdigitating bony processes are reduced and replaced with fibrous connective tissue (Fig. 2c-e; Movie S1).

In adult T. ornata, reduced plastral buttresses permit movement of the anterior and posterior plastral lobes. In E. blandingii, plastral buttresses are partially reduced and thus kinesis is limited to the anterior lobe. By contrast, adult akinetic-shelled C. picta features fully formed buttresses. This trend is mirrored in hatchlings, though buttress reduction occurs via bone remodeling and connective tissue forms as kinetic-shelled T. ornata grows (Fig. 3a-d).

Hinge tissue is highly fibrous, collagen-rich, and covered by a cornified tissue layer (Fig. 3e).

Plastron growth rates varied among species (CL x species interaction: F2, 1953 = 74, P <

0.0001). Terrapene ornata exhibited the highest plastron growth rate (slope = 1.04, 95% CI =

1.02-1.05; N = 221), resulting in a 1:1 PL/CL ratio in adults (1.00 ± 0.004 SE; N = 156) (Fig.

4a-b). Kinetic-shelled E. blandingii featured a similar growth rate (slope = 0.94, 95% CI =

0.93-0.95; N = 136) as akinetic-shelled C. picta (slope = 0.95, 95% CI = 0.94-0.95; N =

1,602), but the adult PL/CL ratio (0.94 ± 0.001 SE; N = 1,365) was higher than in C. picta

(0.92 ± 0.009 SE; N = 70) (Tukey HSD test: P = 0.0002).

Plastron shape shifted with PL during post-hatching growth (ANOVA: F1, 58 = 336, P

< 0.0001), though in a species-specific manner (PL x species: F2, 58 = 18.6, P < 0.0001) (Fig.

4c). In T. ornata (N = 15), plastron shape deformation was greater than in E. blandingii

(Tukey HSD test: P < 0.0001; N = 24) and C. picta (Tukey HSD test: P = 0.001; N = 26). The plastron in E. blandingii and C. picta displayed similar patterns of moderate shape deformation (Fig. 4d-e). By contrast, T. ornata exhibited extensive shape deformation related to a broadening of the anterior plastral lobe (Fig. 4e).

4. Discussion

How the turtle’s shell is patterned in developing embryos has recently come to light

[64, 65], yet many ecologically relevant shell phenotypes emerge long after development in the egg is over [66, 67]. Perhaps this is not surprising as shell development is incomplete in hatchling turtles [68-70]. Nonetheless, documenting progressive changes in the skeletal architecture of such long-lived organisms is challenging. Here, by examining both museum specimens and live-trapped turtles in the wild, we link the convergent evolution of shell kinesis with developmental processes that span both embryonic and post-embryonic life stages.

(a) At least eight independent origins of shell kinesis in extant turtles

Shell kinesis has evolved repeatedly across many fossil and extant turtles since the early Cretaceous, ca. 112 Ma ago [15, 21-29]. Our analyses initially indicated that this complex trait arose independently nine times in extant turtles. However, a more conservative interpretation is that it arose at least eight times. Kinesis is a classic diagnostic feature of

Cyclemys (Geoemydidae) [39], though the phylogeny of Pereira et al. [47] proposes that C. fusca shares a recent common ancestor with akinetic Heosemys and Notochelys. This arrangement would have increased absent-to-present (shell kinesis) transitions by one. Future studies are needed to resolve this discrepancy, but also to clarify the likelihood of reversals to akinesis in geoemydids and emydids. These clades diversified over the last 20 million years and might be predisposed to develop kinesis [71, 72], thus reversals cannot be entirely ruled out [73]. Even so, present-to-absent transitions occurred on average less than once in our character state simulations, suggesting that reversals are rare. Is shell kinesis a common solution to a common ecological problem? Turtles are notably susceptible to predator attacks during juvenile life stages [31, 74], particularly in terrestrial habitats [30]. Thus, predator-driven selection may favor the evolution of shell morphologies that enhance survival [75]. Indeed, kinetic-shelled turtles frequently occupy terrestrial habitats, where the majority of their predators reside, for purposes other than egg laying [37, 50-52, 76, 77]. Although our phylogenetic correlations did not strongly support this well-known trend, we demonstrated that kinetic-shelled species tend to be smaller. These results are congruent with the complex co-evolutionary history of shell morphology and habitat preference in turtles [15, 34, 44, 78, 79]. For instance, some species retained shell kinesis after undergoing terrestrial-to-aquatic reversals [80], and the adaptive value of kinesis may not be entirely related to shell closure in highly aquatic lineages with reduced plastron size, e.g. Sternotherus [81]. Furthermore, the degree of shell closure enabled by kinesis might be associated with the extent to which terrestrial habitats are used [23]. A promising approach to disentangling these intriguing relationships is to integrate ecological, developmental, and phylogenetic studies across diverse kinetic-shelled turtles.

(b) Gradual tissue repatterning underpins the convergent evolution of shell kinesis

The iconic ‘hinge’ joint that defines the anatomy of shell kinesis develops gradually

[37-39]. Based on previous observations [23, 40, 41], we expected this extraordinary transformation to unfold via two plausible sequences of events, summarized as: (i) a

‘heterochronic’ model of delayed sutural fusion followed by elastic tissue formation in the incipient hinge region or (ii) a ‘repatterning’ model predicting normal sutural fusion followed by reorganization with elastic tissue formation. Our histological and morphometric analyses were in agreement with this ‘repatterning’ scenario. Ectodermal plates (i.e. scutes) that cover the shell must align with bone sutures to permit hinge movement. Because this key configuration was absent in hatchlings, scutes must undergo positional rearrangement as turtles grow. We showed that abdominal-pectoral scute boundaries aligned with the hypoplastral-hyoplastral suture as kinesis was activated in juvenile (~ 3 yr) box turtles (T. ornata). Though partially fused, this transient hinge suture was highly flexible. Flexibility increased as the suture was broken down and filled with dense collagen fiber (> 5 yr), corroborating the hypothesis that mechanical strain exerted by muscles during shell closure prevents sutural fusion and leads to the emergence of elastic hinge tissue

[23, 41]. Consistent with comparisons of hatchling and adult emydids [43], we demonstrated morphological variation related to hinge differentiation in T. ornata: the plastron underwent substantial shape deformation, plastron length increased disproportionately, and bony buttresses that structurally bridge the plastron and carapace were repatterned. These changes were less extensive in E. blandingii, consistent with its less derived form of kinesis. Still, buttresses were also reduced and lined with elastic tissue. Akinetic-shelled C. picta did not exhibit shell tissue repatterning.

Shell repatterning in emydid turtles may be generalized to other kinetic-shelled species, particularly geoemydids that also feature a hypoplastral-hyoplastral hinge, i.e.

Cyclemys and Cuora [26]. The hinge is situated distal to the shell buttresses of all other species. Thus, repatterning is likely confined to the developing hinge region of Kinosternon and Pelusios, similarly to lineages that lack scutes (Lissemys) or that feature carapacial kinesis

(Kinixys). Despite structural shell differences, cellular mechanisms that govern sutural fusion and elastic tissue production are probably common to all turtles, as they are shared by all vertebrates [82]. That such mechanisms are employed gradually in ontogeny and long after embryo life stages is noteworthy.

Fibrous joint development often requires mechanical strain provided by skeletal muscle contraction [82, 83]. In kinetic-shelled turtles, novel neck-shoulder-plastron muscle connections established during embryo life stages exert strain on the developing plastron [23-

25]. Hence, hinge differentiation in juveniles is a delayed response to embryonic alterations that transform the shell tissue microenvironment. Remarkably, such musculoskeletal mechanical linkages that enable the development and function of kinesis vary in a lineage- specific manner, rendering the convergent evolution of shell kinesis a fascinating example of many-to-one mapping, i.e. similar function achieved by diverse structures [84]. Similarly, cranial sutures respond to mechanical cues from diverse musculoskeletal sources [85], potentially explaining the convergence of cranial kinesis in a wide variety of vertebrates [86].

Thus, delayed trait differentiation via strain-induced repatterning is crucial to phenotypic diversification.

(d) Evo-devo implications of delayed trait ‘acquisition’ via tissue repatterning

Kinetic shell hinges are not merely ‘acquired’, at least not in a Lamarckian sense [87], by slowly-growing turtles. Instead, delayed trait differentiation is a function-induced developmental transformation (e.g. [88]), which initially requires heritable genetic change in embryonic traits. Crucially, form-to-function effects on the shell will only manifest as individuals mature. This key developmental process promotes novel phenotypic variants that may otherwise not arise in embryogenesis owing to developmental constraints. Although these plastic properties of developing skeletal tissue were first described in the 19th century

[89], how they facilitate macroevolutionary change has only been emphasized in recent decades [90-94]. Likewise, our study highlights that the genotype-to-phenotype translation is not always a one-to-one relationship, as it may often involve tinkering of intricately interrelated traits over the course of multiple life stages.

Ethics. Procedures were approved by the ISU Institutional Animal Care and Use committee

(protocol # 2-11-7091-J).

Data accessibility. Data are available as electronic supplementary material.

Competing interests. We declare none.

Authors’ contributions. G.A.C. designed the study, wrote the paper, and performed phylogenetic analyses; G.A.C. and K.Q. conducted histological assays and morphometric analyses; G.A.C. and F.J. conducted field sampling; F.J. provided logistical support and advice.

Funding. NSF Doctoral Dissertation Improvement Grant (DEB-1310874) and grants DEB-

0640932 and DEB-1242510. Additional support from: California Academy of Sciences,

Chicago Herpetological Society, Nebraska Herpetological Society, ISU-EEOB, and Society for Integrative and Comparative Biology.

Acknowledgements. We thank Janzen lab members who processed shell measurements and

Andy McCollum for donating box turtle specimens. We are grateful to: John Iverson and

Anthony Beringer for help in the field, Jenny Groeltz-Thrush for assistance with histology, and Dean Adams for access to 3D scanners. We also thank instructors of the UC Davis

Applied Phylogenetics and Santa Barbara Comparative Methods and Macroevolution in R workshops.

References

1. Dobzhansky T. 1959 Evolution and Environment. In The Evolution of Life: Its Origin,

History, and Future (ed. Tax S.), pp. 403-428. Chicago, The University of Chicago Press. 2. Losos J.B. 2011 Convergence, adaptation, and constraint. Evolution 65, 1827-1840.

(doi:10.1111/j.1558-5646.2011.01289.x).

3. Wake D.B., Wake M.H., Specht C.D. 2011 Homoplasy: from detecting pattern to determining process and mechanism of evolution. Science 331, 1032-1035.

(doi:10.1126/science.1188545).

4. McGhee G. 2011 Convergent Evolution: Limited Forms Most Beautiful. Cambridge,

Massachusetts The MIT Press.

5. Agrawal A.A. 2017 Toward a Predictive Framework for Convergent Evolution:

Integrating Natural History, Genetic Mechanisms, and Consequences for the Diversity of

Life. The American Naturalist 190, S1-S12. (doi:10.1086/692111).

6. Mahler D.L., Weber M.G., Wagner C.E., Ingram T. 2017 Pattern and process in the comparative study of convergent evolution. The American Naturalist 190, S13-S28.

(doi:10.1086/692648).

7. Sanger T.J., Revell L.J., Gibson-Brown J.J., Losos J.B. 2012 Repeated modification of early limb morphogenesis programmes underlies the convergence of relative limb length in

Anolis lizards. Proc Biol Sci 279, 739-748. (doi:10.1098/rspb.2011.0840).

8. Sanger T.J., Sherratt E., McGlothlin J.W., Brodie E.D., 3rd, Losos J.B., Abzhanov A.

2013 Convergent evolution of sexual dimorphism in skull shape using distinct developmental strategies. Evolution 67, 2180-2193. (doi:10.1111/evo.12100).

9. Mallarino R., Campàs O., Fritz J.A., Burns K.J., Weeks O.G., Brenner M.P.,

Abzhanova A. 2012 Closely related bird species demonstrate flexibility between beak morphology and underlying developmental programs. Proc Natl Acad Sci USA 109, 16222-

16227. 10. Cooper K.L., Sears K.E., Uygur A., Maier J., Baczkowski K.S., Brosnahan M.,

Antczak D., Skidmore J.A., Tabin C.J. 2014 Patterning and post-patterning modes of evolutionary digit loss in mammals. Nature 511, 41-45. (doi:10.1038/nature13496).

11. Donoghue M.J., Ree R.H. 2000 Homoplasy and developmental constraint: a model and an example from plants. American Zoologist 40, 759-769.

12. Bock W.J. 1963 Evolution and phylogeny in morphologically uniform groups. The

American Naturalist 97, 265-285.

13. Cordero G.A. 2014 Re-emergence of the painted turtle (Chrysemys picta) as a reference species for evo-devo. Evol Dev 16, 184-188. (doi:10.1111/ede.12082).

14. Cordero G.A. 2017 The turtle's shell. Current Biology 27, R163-R171.

15. Pritchard P.C.H. 2008 Evolution and Structure of the Turtle Shell. In Biology of

Turtles (eds. Wyneken J., Godfrey M.H., Bels V.), pp. 45-84. Boca Raton, CRC Press.

16. Cordero G.A., Quinteros K. 2015 Skeletal remodelling suggests the turtle's shell is not an evolutionary straitjacket. Biology letters 11, 20150022. (doi:10.1098/rsbl.2015.0022).

17. Sarnat B.G., McNabb E.G. 1981 Sutural bone growth of the turtle Chrysemys scripta plastron: A serial radiographic study by means of radiopaque implants. Growth 45, 123-123.

18. Cherepanov G.O. 2005 The Turtle Shell: Morphogenesis and Evolution. St.

Petersburg, St. Petersburg State University.

19. Zangerl R. 1969 The Turtle Shell. In Biology of the Reptilia (eds. Gans C., Bellairs

A.d.A., Parsons T.S.), pp. 311-339. London, Academic Press.

20. Scheyer T.M. 2007 Comparative bone histology of the turtle shell (carapace and plastron): implications for turtle systematics, functional morphology and turtle origins. Bonn,

Rheinischen Friedrich-Wilhelms-Universitä t zu Bonn. 21. Sukhanov V.B. 2000 Mesozoic turtles of middle and central Asia. In The Age of

Dinosaurs in Russia and Mongolia (eds. Benton M.J., Shishkin M.A., Unwin D.M.,

Kurochkin E.N.). Cambridge, Cambridge University Press.

22. Hasan S.I. 1941 The shell and the mechanisms of its closure in the indian pond terrapin, Lissemys punctata punctata (Bonnaterre). Proceedings of Indian Academy of

Sciences, Section B 14, 235-249.

23. Bramble D.M. 1974 Emydid shell kinesis: biomechanics and evolution. Copeia 1974,

707-727.

24. Bramble D.M., Hutchison J.H. 1981 A reevaluation of plastral kinesis in African turtles of the genus Pelusios. Herpetologica 37, 205-212.

25. Bramble D.M., Hutchison J.H., Legler J.M. 1984 Kinosternid shell kinesis: structure, function and evolution. Copeia 1984, 456-475.

26. McLaughlin C.J., Stayton C.T. 2016 Convergent evolution provides evidence of similar radiations in shell shape in the turtle families Emydidae and Geoemydidae.

Herpetologica 72, 120-129. (doi:10.1655/herpetologica-d-15-00037).

27. Shah R.V. 1960 The mechanisms of carapacial and plastral hinges in chelonians.

Brevoria 130, 1-9.

28. Hutchison J.H. 2013 New turtles from the Paleogene of North America. In

Morphology and Evolution Turtles (eds. Brinkman D.B., Holroyd P.A., Gardner J.D.), pp.

477-497. New York, Springer.

29. Claude J. 2006 Convergence induced by plastral kinesis and phylogenetic constraints in Testudinoidea: A geometric morphometric assessment. Fossil Turtle Res 1, 34-45.

30. Green H.W. 1988 Antipredator mechanisms in reptiles. In Biology of the Reptilia (eds.

Gans C., Huey R.B.), pp. 1-152. New York, Alan R. Liss, Inc.

31. Minckley W.L. 1966 Coyote predation on aquatic turtles. J Mammal 47, 137. 32. Germano D.J. 1999 Terrapene ornata luteola. Attempted predation. Herpetological

Review 30, 40-41.

33. Mitchell J.C. 2006 Kinosternon subrubrum subrubrum (eastern mud turtle). Predator escape. Herpetological Review 37, 216-217.

34. Angielczyk K.D., Feldman C.R., Miller G.R. 2011 Adaptive evolution of plastron shape in emydine turtles. Evolution 65, 377-394. (doi:10.1111/j.1558-5646.2010.01118.x).

35. Wygoda M.L., Chmura C.M. 1990 Effects of shell closure on water loss in the sonoran mud turtle, Kinosternon sonoriense. Southwestern Nat 35, 228-229.

36. Agassiz L. 1857 Contributions to the Natural History of the United States of America: pt. I. Essay on classification. pt. II. North American Testudinata. 1857, Little, Brown and

Company.

37. Legler J.M. 1960 Natural history of the ornate box turtle, Terrapene ornata ornata

Agassiz. University of Kansas Publications Museum of Natural History 11, 527-669.

38. Richmond N.D. 1964 The mechanical functions of the testudinate plastron. American

Midland Naturalist 72, 50-56.

39. Fritz U., Guicking D., Auer M., Sommer R.S., Wink M., Hundsdrfer A.K. 2008

Diversity of the Southeast Asian leaf turtle genus Cyclemys: how many leaves on its tree of life? Zoologica Scripta 37, 367-390. (doi:10.1111/j.1463-6409.2008.00332.x).

40. Ernst C.H., Lovich J.E., Laemmerzahl A.F., Sekscienski S. 1997 A comparison of plastral scute lengths among members of the box turtle genera Cuora and Terrapene.

Chelonian Conservation and Biology 2, 603-607.

41. Pritchard P.C.H. 2003 Akinesis and plastral scute homologies in Sternotherus

(Testudines: Kinosternidae). Chelonian Conservation and Biology 4, 671-674. 42. Martinez-Maza C., Rosas A., Nieto-Diaz M. 2013 Postnatal changes in the growth dynamics of the human face revealed from bone modelling patterns. Journal of anatomy 223,

228-241. (doi:10.1111/joa.12075).

43. Angielczyk K.D., Feldman C.R. 2013 Are diminurtive turtles miniaturized? The ontogeny of plastron shape in emydine turtles. Biological Journal of the Linnean Society 108,

727-755.

44. Claude J., Paradis E., Tong H., Auffray J.-C. 2003 A geometric morphometric assessment of the effects of environment and cladogenesis on the evolution of the turtle shell.

Biological Journal of the Linnean Society 79, 485-501.

45. Revell L.J. 2012 phytools: an R package for phylogenetic comparative biology (and other things). Methods in Ecology and Evolution 3, 217-223. (doi:10.1111/j.2041-

210X.2011.00169.x).

46. Huelsenbeck J.P., Nielsen R., Bollback J.P. 2003 Stochastic mapping of morphological characters. Systematic Biology 52, 131-158.

(doi:10.1080/10635150390192780).

47. Pereira A.G., Sterli J., Moreira F.R.R., Schrago C.G. 2017 Multilocus phylogeny and statistical biogeography clarify the evolutionary history of major lineages of turtles.

Molecular phylogenetics and evolution 113, 59-66. (doi:10.1016/j.ympev.2017.05.008).

48. Garland T., Dickerman A.W., Janis C.M., Jones J.A. 1993 Phylogenetic analysis of covariance by computer simulation. Systematic Biology 42, 265-292.

49. Pagel M. 1994 Detecting correlated evolution on phylogenies: A general method for the comparative analysis of discrete characters. Proceedings: Biological Sciences 255, 37-45.

50. Cordero G.A., Quinlan M., Blackstone S., Swarth C.W. 2012 Kinosternon subrubrum

(eastern mud turtle). Overwintering. Herpetological Review 43, 327. 51. Cordero G.A., Reeves R., Swarth C.W. 2012 Long distance aquatic movement and home-range size of an eastern mud turtle, Kinosternon subrubrum, population in the Mid-

Atlantic region of the United States. Chelonian Conservation and Biology 11, 121-124.

52. Cordero G.A., Swarth C.W. 2010 Notes on the movement and aquatic behavior of some kinosternid turtles. Acta Zoologica Mexicana 26, 233-235.

53. An Y.H., Martin K.L. 2003 Handbook of Histology Methods for Bone and Cartilage.

Totowa, New Jersey, Humana Press.

54. Warner D.A., Miller D.A., Bronikowski A.M., Janzen F.J. 2016 Decades of field data reveal that turtles senesce in the wild. Proceedings of the National Academy of Sciences of the

United States of America 113, 6502-6507. (doi:10.1073/pnas.1600035113).

55. Cordero G.A., Janzen F.J. 2014 An enhanced developmental staging table for the painted turtle, Chrysemys picta (Testudines: Emydidae). J Morphol 275, 442-455.

(doi:10.1002/jmor.20226).

56. Mossiman J.E. 1958 An analysis of shell alometry in turtles. Revue Canadienne de

Biologie, 137-228.

57. Anderson D.E., Lydic R. 1977 On the effect of using ratios in the analysis of variance.

Behavioral Reviews 1, 225-229.

58. Myers E.M., Janzen F.J., Adams D.C., Tucker J.K. 2006 Quantitative genetics of plastron shape in slider turtles (Trachemys scripta). Evolution 60, 563-572.

59. Rohlf F.J. 2010 tpsDig, version 2.16. (Department of Ecology and Evolution, Stony

Brook University, Stony Brook, NY.

60. Rohlf F.J. 2010 tpsRelW: thin plate spline relative warp analysis, version 1.49.

(Department of Ecology and Evolution, Stony Brook University, Stony Brook, NY.

61. Zelditch M.L., Swiderski D.L., Sheets H.D. 2012 Geometric Morphometrics for

Biologists: A Primer. Second Edition. Amsterdam, Elselvier Academic Press; i-iii p. 62. Rohlf F.J. 2004 Thin-plate spline, version 1.20. (Department of Ecology and

Evolution, Stony Brook University, Stony Brook, NY.

63. R Development Core Team. 2017 R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. Version 3.3.3 available at http://www.R-project.org. (

64. Moustakas-Verho J.E., Zimm R., Cebra-Thomas J., Lempiainen N.K., Kallonen A.,

Mitchell K.L., Hamalainen K., Salazar-Ciudad I., Jernvall J., Gilbert S.F. 2014 The origin and loss of periodic patterning in the turtle shell. Development 141(15), 3033-3039.

(doi:10.1242/dev.109041).

65. Rice R., Kallonen A., Cebra-Thomas J., Gilbert S.F. 2016 Development of the turtle plastron, the order-defining skeletal structure. Proceedings of the National Academy of

Sciences of the United States of America 113, 5317-5322. (doi:10.1073/pnas.1600958113).

66. Kordikova E. 2000 Paedomorphosis in the shell of fossil and living turtles. Neues

Jahrbuch für Geologie und Paläontologie Abhandlungen 218(3), 399-446.

67. Kordikova E.G. 2002 Heterochrony in the evolution of the shell of Chelonia. Part 1: terminology, Cheloniidae, Dermochelyidae, Trionychidae, Cyclanorbidae and Carettochelyidae. Neues Jahrbuch für Geologie und Paläontologie Abhandlungen 226,

343-417.

68. Suzuki H.K. 1963 Studies on the osseous system of the slider turtle. Annals of the New

York Academy of Sciences 109, 351-410.

69. Gilbert S.F., Loredo G.A., Brukman A., Burke A.C. 2001 Morphogenesis of the turtle shell: the development of a novel structure in tetrapod evolution. Evolution & development 3,

47-58. 70. Cherepanov G.O. 2016 Scute’s polymorphism as a source of evolutionary development of the turtle shell. Paleontological Journal 49, 1635-1644.

(doi:10.1134/s003103011514004x).

71. Joyce W.G., Bell C.J. 2004 A review of the comparative morphology of extant testudinoid turtles (Reptilia: Testudines). Asiatic Herpetological Research 10, 53-109.

72. Seidel M.E., Ernst C.H. 2017 A systematic review of the turtle family Emydidae.

Vertebrate Zoology 67, 1-122.

73. Feldman C.R., Parham J.F. 2002 Molecular phylogenetics of emydine turtles: taxonomic revision and the evolution of shell kinesis. Mol Phylogenet Evol 22(3), 388-398.

(doi:10.1006/mpev.2001.1070).

74. Tucker J.K., Filoramo N.I., Janzen F.J. 1999 Size-biased mortality due to predation in a nesting freshwater turtle, Trachemys scripta. The American Midland Naturalist 141(1), 198-

203. (doi:10.1674/0003-0031(1999)141[0198:Sbmdtp]2.0.Co;2).

75. Salmon M., Scholl J. 2014 Allometric growth in juvenile marine turtles: possible role as an antipredator adaptation. Zoology 117, 131-138. (doi:10.1016/j.zool.2013.11.004).

76. Bonin F., Devaux B., Dupre ́ A. 2006 Turtles of the World. London, A and C

Publishers.

77. Pritchard P.C.H. 1979 Encyclopedia of Turtles. New Jersey, T.F.H. Publications, Inc.

78. Benson R.B.J., Domokos G., Va ́rkonyi P.L., Reisz R.R. 2011 Shell geometry and habitat determination in extinct and extant turtles (Reptilia: Testudinata). Paleobiology 37,

547-562. (doi:10.5061/dryad.8705).

79. Scheyer T.M., Danilov I.G., Sukhanov V.B., Syromyatnikova E.V. 2014 The shell bone histology of fossil and extant marine turtles revisited. Biological Journal of the Linnean

Society 112, 701-718. 80. Stephens P.R., Wiens J.J. 2009 Bridging the gap between community ecology and historical biogeography: niche conservatism and community structure in emydid turtles.

Molecular ecology 18(22), 4664-4679. (doi:10.1111/j.1365-294X.2009.04378.x).

81. Iverson J.B., Le M., Ingram C. 2013 Molecular phylogenetics of the mud and musk turtle family Kinosternidae. Molecular phylogenetics and evolution 69, 929-939.

(doi:10.1016/j.ympev.2013.06.011).

82. Hall B.K. 2015 Bones and Cartilage, Developmental and Evolutionary Skeletal

Biology. London, Academic Press.

83. Salva J.E., Merrill A.E. 2017 Signaling networks in joint development. Dev Dyn 246,

262-274. (doi:10.1002/dvdy.24472).

84. Wainwright P.C. 2007 Functional versus morphological diversity in macroevolution.

Annual Review of Ecology, Evolution, and Systematics 38(1), 381-401.

(doi:10.1146/annurev.ecolsys.38.091206.095706).

85. Khonsari R.H., Olivier J., Vigneaux P., Sanchez S., Tafforeau P., Ahlberg P.E., Di

Rocco F., Bresch D., Corre P., Ohazama A., et al. 2013 A mathematical model for mechanotransduction at the early steps of suture formation. Proceedings Biological sciences /

The Royal Society 280(1759), 20122670. (doi:10.1098/rspb.2012.2670).

86. Mezzasalma M., Maio N., Guarino F.M. 2014 To move or not to move: cranial joints in European gekkotans and lacertids, an osteological and histological perspective. Anat Rec

297, 463-472. (doi:10.1002/ar.22827).

87. Gissis S.B., Jablonka E. 2011 Transformations of Lamarckism: From Subtle Fluids to

Molecular Biology. The MIT Press, Cambridge, Massachusetts.

88. Zelditch M.L., Wood A.R., Swiderski D.L. 2008 Building Developmental Integration into Functional Systems: Function-Induced Integration of Mandibular Shape. Evolutionary

Biology 36, 71-87. (doi:10.1007/s11692-008-9034-7). 89. Wolff J. 1892 Das Gesetz der Transformation der Knochen. Berlin, Verlag von

August Hirschwald.

90. Frazzetta T.H. 1976 Complex Adaptations in Evolving Populations. Sunderland,

Massachusetts, Sinauer Associates, Inc.

91. Milinkovitch M.C., Manukyan L., Debry A., Di-Poi N., Martin S., Singh D., Lambert

D., Zwicker M. 2013 Crocodile head scales are not developmental units but emerge from physical cracking. Science 339, 78-81. (doi:10.1126/science.1226265).

92. Newman S.A. 2003 From physics to development: the evolution of morphogenetic mechanisms. In Origination of Organismal Form (eds. Mü ller G.B., Newman S.A.), pp. 221-

240. Cambridge, Massachusetts, The MIT Press.

93. Cordero G.A., Berns C.M. 2016 A test of Darwin's 'lop-eared' rabbit hypothesis. J

Evol Biol 29(11), 2102-2110. (doi:10.1111/jeb.12938).

94. Griffin C.T., Nesbitt S.J. 2016 Anomalously high variation in postnatal development is ancestral for dinosaurs but lost in birds. Proceedings of the National Academy of Sciences of the United States of America 113, 14757-14762. (doi:10.1073/pnas.1613813113).

95. Carr A.F. 1952 Handbook of Turtles: The Turtles of the United States, Canada, and

Baja California. Ithaca, Comstock Pub. Associates.

96. Pough F.H., Janis C.M., Heiser J.B. 2001 Vertebrate Life 6th edition. Upper Daddle

River, New Jersey, Prentice Hall. Figure Legends

Figure 1. Shell kinesis via hinge rotation in box turtles (illustration credit: Jessica Gassman; a). Pectoral (Pec) and abdominal (Ab) ectodermal scutes are aligned with the suture of hyoplastron (Hyo) hypoplastron (Hyp) bones (b; modified from [95]), unlike in akinetic- shelled species (b; modified from [96]). Kinesis enhances predator defense on land: an eastern box turtle (Terrapene carolina) survives an attack by activating its plastral hinge (c; from Life in Cold Blood with permission from the British Broadcasting Company). Shell kinesis (absent

= blue; present = red) has evolved at least 8 times, probably in conjunction with evolutionary transitions from aquatic (cyan) to terrestrial (yellow) habitats in small-bodied turtles (CL = carapace length in mm) (d-e); time-calibrated phylogeny from [47].

Figure 2. The immature junction (inlet: H & E sagittal section) of the pectoral (Pec) and abdominal (Ab) scutes in hatchling Terrapene ornata does not feature a plastral hinge (a).

Ossification centers of the hyoplastral (Hyo) and hypoplastral (Hyp) bones (in alizarin red) expand and begin to fuse near the Pec-Ab scute junction a year after hatching in T. ornata (b).

These bones are sutured three years post-hatching (c). By year five, the suture is re-patterned and replaced by fibrous connective tissue, particularly on the external surface of the plastron

(d). Fibrous connective tissue proliferates externally and the serrated edges on the interior plastral surface are further reduced in fully-grown adults (e). Silhouettes are based on images from [77].

Figure 3. Plastral buttresses (axillary = Ax; inguinal = In) are fully developed in Chrysemys picta (a), partially reduced in Emys blandingii (b), and fully reduced in Terrapene ornata (c).

This key difference is pre-patterned in hatchlings (left panel) and corresponds to akinesis in

C. picta, as well as varying degrees of kinesis (arrows) in adult E. blandingii versus T. ornata (c; middle and right panels). Progressive breakdown of bone and scute tissue and replacement with fibrous connective tissue near the axillary plastral buttress in growing T. ornata (d). The hyoplastral-hypoplastral suture (VVG sagittal section) displays an interdigitating pattern in akinetic turtles (left: Glyptemys insculpta; scale bar = 10 µm), in contrast to the cornified

(yellow) and collagen-rich (red) fibrous connective tissue comprising the hinge of kinetic- shelled species (right: T. ornata; scale bar = 50 µm) (e).

Figure 4. Scaling of the plastron during embryonic and post-hatching stages varied among species (a), as T. ornata (TO) displayed a higher growth rate and thus attained a larger adult plastron-to-carapace length ratio compared to C. picta (CP) and E. blandingii (EB) (b).

Plastron shape deformation during post-hatching growth was also greater in T. ornata (c). A principal component (PC) plot supports that T. ornata undergoes substantial shape deformation (d), as the anterior plastral lobe broadens during post-hatching growth (e). Figure 1 Figure 2 Figure 3 Figure 4 ELECTRONIC SUPPLEMENTARY MATERIAL

Convergent phenotypes develop slowly in turtles with shell kinesis G.A. Cordero, K. Quinteros, F.J. Janzen

1. Supplementary materials and methods

(a) Character state data

Character data (Fig. S1; Table S1) were obtained from anatomical appraisals of shell kinesis [1-4]. Putative modes of shell kinesis not associated with musculoskeletal mechanical linkages that enable shell closure were excluded, for example shell kinesis related to egg-laying in females [5]. Habitat type data for 262 species were obtained from the comprehensive review of Mota-Rodrigues and Felizola Diniz-Filho [6], which followed a binary (aquatic versus terrestrial) classification scheme [7]. Habitat type data for an additional 30 species was obtained from other sources [8-19]. Maximum carapace length (CL) was obtained for 277 species from the most recent global analysis of turtle body size evolution [20]. Additional data for 11 species were obtained from other reviews [21, 22], as well as the primary literature for three species [23-25].

(b) Museum specimens used in this study

Hatchlings of Pelusios castaneus were purchased from a private breeder, otherwise all other museum- preserved hatchlings (see Fig. S2) originated from eggs collected in the wild and incubated in moist vermiculite (-150 kPa water potential) in an environmental chamber set to a constant 27 ºC or 30 ºC [see ref. 26]. We measured embryos of Chrysemys picta (N = 74), Emys blandingii (N = 43), and Terrapene ornata (N = 38) also previously sampled by Cordero and Quinteros [26]. In addition, hatchlings from their study were measured: Chrysemys picta (N = 33), Emys blandingii (N = 16), and Terrapene ornata (N = 6).

We examined adults specimens of E. blandingii, Glyptemys insculpta, and T. carolina donated to the Iowa State University herpetological collection by the Iowa Department of Natural Resources. Fluid- preserved specimens of T. ornata and C. picta collected in Thomson, Illinois and deposited in this collection were also examined. In addition, we sampled road-killed T. ornata collected in Grant and Hooker counties, Nebraska. Specimens of juvenile T. ornata from eastern Iowa were obtained from the teaching collection of Cornell College (Mount Vernon, Iowa). We estimated the age of juvenile T. ornata by counting primary growth rings, following the technique of Legler [27].

Sampling was conducted with permits from: Arkansas Game & Fish Commission #020520132; Illinois Dept. of Natural Resources #NH13.0073; Iowa Dept. of Natural Resources #14; Nebraska Game & Parks Commission #310. Representative specimens examined in this study were transferred to the herpetological collections of the University of Kansas and Carnegie Museum of Natural History.

2. Supplementary results

Figure S1. Shell kinesis has evolved at least eight times in extant turtle lineages, as supported by stochastic character mapping on the time-calibrated phylogeny of Pereira et al. [28]. Shell kinesis: blue = absent; red = present. Posterior probabilities (colour gradients) were estimated on 1,000 stochastic character maps. Ancestral state reconstructions followed an equal rates model (Markov k-state 1) of state transitions (rate = 0.001).

Figure S2. Plastron development is incomplete in hatchling turtles, such that some akinetic (a = Chrysemys picta; b = Chelydra serpentina) and kinetic-shelled species feature similar ossification patterns (alizarin red stain; scale bars = 2 mm). Locations of incipient plastral hinges (arrows) may vary according to lineage: Hyoplastron-hypoplastron (Hyo-Hyp) suture in emydids Emys blandingii (c) and Terrapene ornata (d); two hinges in Kinosternidae (e; Kinosternon subrubrum) will differentiate at the Epi (epiplastron)-Hyo and Hyp-Xip (xiphiplastron) sutures; in Pelomedusidae (f; Pelusios castaneus) a hinge eventually differentiates at the Hyo-mesoplastron (Me) suture. 3. Supplementary references

1. Bramble D.M. 1974 Emydid shell kinesis: biomechanics and evolution. Copeia 1974, 707-727.

2. Bramble D.M., Hutchison J.H. 1981 A reevaluation of plastral kinesis in African turtles of the genus Pelusios. Herpetologica 37, 205-212.

3. Bramble D.M., Hutchison J.H., Legler J.M. 1984 Kinosternid shell kinesis: structure, function and evolution. Copeia 1984, 456-475.

4. Hasan S.I. 1941 The shell and the mechanisms of its closure in the indian pond terrapin, Lissemys punctata punctata (Bonnaterre). Proceedings of Indian Academy of Sciences, Section B 14, 235-249.

5. Pritchard P.C.H. 2008 Evolution and Structure of the Turtle Shell. In Biology of Turtles (eds. Wyneken J., Godfrey M.H., Bels V.), pp. 45-84. Boca Raton, CRC Press.

6. Mota-Rodrigues J.F., Felizola Diniz-Filho J.A. 2016 Ecological opportunities, habitat, and past climatic fluctuations influenced the diversification of modern turtles. Molecular phylogenetics and evolution 101, 352-358. (doi:10.1016/j.ympev.2016.05.025).

7. Claude J., Paradis E., Tong H., Auffray J.-C. 2003 A geometric morphometric assessment of the effects of environment and cladogenesis on the evolution of the turtle shell. Biological Journal of the Linnean Society 79, 485-501.

8. Das I. 2009 Melanochelys tricarinata (Blyth 1856) – tricari- nate hill turtle, three-keeled land turtle. In: Rhodin, A.G.J., Pritchard, P.C.H., van Dijk, P.P., Saumure, R.A., Buhlmann, K.A., Iverson, J.B., and Mittermeier, R.A. (Eds.). Conserva- tion Biology of Freshwater Turtles and Tortoises: A Compila- tion Project of the IUCN/SSC Tortoise and Freshwater Turtle Specialist Group. . Chelonian Research Monographs 5, 025.021-025.025. (doi:10.3854/crm.5.025.tricarinata.v1.2009).

9. Iverson J.B., Le M., Ingram C. 2013 Molecular phylogenetics of the mud and musk turtle family Kinosternidae. Molecular phylogenetics and evolution 69, 929-939. (doi:10.1016/j.ympev.2013.06.011).

10. Pritchard P.C.H. 1996 The Galápagos tortoises: nomenclatural and survival status, Chelonian Research Foundation Lunenberg, MA.

11. Yang P., Tang Y., Ding L., Guo X., Wang Y. 2011 Validity of Pelodiscus parviformis (Testudines: Trionychidae) inferred from molecular and morphological analyses. Asian Herpetological Research 2, 21-29. (doi:10.3724/sp.j.1245.2011.00021).

12. Bertl J., Killebrew F.C. 1983 An osteological comparison of Graptemys caglei Haynes and McKown and Graptemys versa Stejneger (Testudines: Emydidae). Herpetologica 39, 375-382.

13. Rhodin A.G.J., Mittermeier R.A. 1983 Description of Phrynops williamsi, a new species of chelid turtle of the South American P. geoffroanus complex. In Advances in Herpetology and Evolutionary Biology Essays in Honor of Ernest E Williams (eds. Rhodin A.G.J., Miyata K.), pp. 58- 73. Cambridge, Massachusetts, Museum of Comparative Zoology. 14. Souza F.L., Abe A.S. 1998 Resource partitioning by the neotropical freshwater turtle, Hydromedusa maximiliani. Journal of Herpetology 32, 106-112.

15. Georges A., Adams M., McCord W. 2002 Electrophoretic delineation of species boundaries within the genus Chelodina (Testudines: Chelidae) of Australia, New Guinea and Indonesia. Zoological Journal of the Linnean Society 134, 401-421.

16. Whitaker N., Vijaya J. 2009 Biology of the forest cane turtle, Vijachelys silvatica, in south India. Chelonian Conservation and Biology 8, 109-115.

17. Molina F.B., Machado F.A., Zaher H. 2012 Taxonomic validity of Mesoclemmys heliostemma (McCord, Joseph-Ouni & Lamar, 2001) (Testudines, Chelidae) inferred from morphological analysis. Zootaxa 3575, 63-77.

18. Santana D.O., Marques T.S., Vieira G.H.C., Moura G.J.B., Faria R.G., Mesquita D.O. 2016 Mesoclemmys tuberculata (Luederwaldt 1926) – Tuberculate Toad-headed Turtle. In: Rhodin, A.G.J., Iverson, J.B., van Dijk, P.P., Saumure, R.A., Buhlmann, K.A., Pritchard, P.C.H., and Mittermeier, R.A. (Eds.). Conservation Biology of Freshwater Turtles and Tortoises: A Compilation Project of the IUCN/SSC Tortoise and Freshwater Turtle Specialist Group. Chelonian Research Monographs 5, 097.091-097.098. (doi:10.3854/crm.5.097.tuberculata.v1.2016).

19. Magnusson W.E., Vogt R.C. 2014 Rhinemys rufipes (Spix 1824) – Red Side-necked Turtle, Red-footed Sideneck Turtle, Perema. In: Rhodin, A.G.J., Pritchard, P.C.H., van Dijk, P.P., Saumure, R.A., Buhlmann, K.A., Iverson, J.B., and Mittermeier, R.A. (Eds.). Conservation Biology of Freshwater Turtles and Tortoises: A Compilation Project of the IUCN/SSC Tortoise and Freshwater Turtle Specialist Group. Chelonian Research Monographs 5, 079.071-079.077. (doi:10.3854/crm.5.079.ru).

20. Itescu Y., Karraker N.E., Raia P., Pritchard P.C.H., Meiri S. 2014 Is the island rule general? Turtles disagree. Global Ecology and Biogeography 23, 689-700. (doi:10.1111/geb.12149).

21. Jaffe A.L., Slater G.J., Alfaro M.E. 2011 The evolution of island gigantism and body size variation in tortoises and turtles. Biology letters 7, 558-561. (doi:10.1098/rsbl.2010.1084).

22. Halamkova L., Schulte II J.A., Langen T.A. 2013 Patterns of sexual size dimosphism in chelonia. Biological Journal of the Linnean Society 108, 396-413.

23. Pritchard P. 2013 Madagascar: island continent of tortoises great and small. In Turtles on the Brink in Madagascar: Proceedings of Two Workshops on the Status, Conservation, and Biology of Malagasy Tortoises and Freshwater Turtles (eds. Castellano C.M., Rhodin A.G.J., Ogle M., Mittermeier R.A., Randriamahazo H., Hudson R., Lewis R.E.), pp. 17-24. Luneburg, Chelonian Research Foundation.

24. Hansen D.M., Donlan C.J., Griffiths C.J., Campbell K.J. 2010 Ecological history and latent conservation potential: large and giant tortoises as a model for taxon substitutions. Ecography 372, 272-284. (doi:10.1111/j.1600-0587.2010.06305.x).

25. Barreto L., Lima L.C., Barbosa S. 2009 Observations on the ecology of Trachemys adiutrix and Kinosternon scorpioides on Curupu Island, Brazil. Herpetological Review 40, 283-286. 26. Cordero G.A., Quinteros K. 2015 Skeletal remodelling suggests the turtle's shell is not an evolutionary straitjacket. Biology letters 11, 20150022. (doi:10.1098/rsbl.2015.0022).

27. Legler J.M. 1960 Natural history of the ornate box turtle, Terrapene ornata ornata Agassiz. University of Kansas Publications Museum of Natural History 11, 527-669.

28. Pereira A.G., Sterli J., Moreira F.R.R., Schrago C.G. 2017 Multilocus phylogeny and statistical biogeography clarify the evolutionary history of major lineages of turtles. Molecular phylogenetics and evolution 113, 59-66. (doi:10.1016/j.ympev.2017.05.008).