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. Author manuscript; available in PMC: 2022 Mar 10.
Published in final edited form as: J Exp Zool B Mol Dev Evol. 2019 Oct 4;336(2):129–144. doi: 10.1002/jez.b.22902

Towards comparative analyses of salamander limb regeneration

Varun B Dwaraka 1,2, S Randal Voss 1
PMCID: PMC8908358  NIHMSID: NIHMS1783527  PMID: 31584252

Abstract

Among tetrapods, only salamanders can regenerate their limbs and tails throughout life. This amazing regenerative ability has attracted the attention of scientists for hundreds of years. Now that large, salamander genomes are beginning to be sequenced for the first time, omics tools and approaches can be used to integrate new perspectives into the study of tissue regeneration. Here we argue the need to move beyond the primary salamander models to investigate regeneration in other species. Salamanders at first glance come across as a phylogenetically conservative group that has not diverged greatly from their ancestors. While salamanders do present ancestral characteristics of basal tetrapods, including the ability to regenerate limbs, data from fossils and data from studies that have tested for species differences suggest there may be considerable variation in how salamanders develop and regenerate their limbs. We review the case for expanded studies of salamander tissue regeneration and identify questions and approaches that are most likely to reveal commonalities and differences in regeneration among species. We also address challenges that confront such an initiative, some of which are regulatory and not scientific. The time is right to gain evolutionary perspective about mechanisms of tissue regeneration from comparative studies of salamander species.

Keywords: comparative analysis, evolution, limb regeneration, salamander

1 |. INTRODUCTION

Modern amphibians – anurans, caecilians, and salamanders – are as deeply diverged from one another as they are to humans. Their evolutionary origin traces to ancestors that inhabited the Pangean landmass over 300 million years ago. Subsequently, these groups evolved independently as Laurasian and Gondwanan supercontinents drifted apart (Feller & Hedges, 1998; Hallam, 1995). Given this deep split, it is not too surprising that amphibians and amniotes show variation in a number of traits, including tissue regeneration. Similarly, it is not too surprising that variation exists in mechanisms of regeneration among species of salamander families that diverged over 100 million years ago. The reality is that we have only begun to scratch the surface in documenting variation in tissue regeneration among salamanders. The goal of our review is to motivate comparative studies of regeneration, to obtain data that are needed to understand why salamanders (and not humans) can regenerate appendages and other organs.

2 |. LIMB REGENERATION – PHYLOGENETIC CONSIDERATIONS

The origin of salamanders is hypothesized to have occurred approximately 297 MYA during the late Permian, marked by the lineage split between salamanders and anurans (Zhang & Wake, 2009). All salamanders belong to the order Caudata (Evans, 1996), which includes two taxonomic suborders representing 10 families: Cryptobranchoidea (Hynobiidae, Cryptobranchidae) and Salamandroidea (Ambystomatidae, Dicamptodontidae, Salamandridae, Protidae, Rhyacotritonidae, Amphiumidae, Plethadontidae, and Sirenidae). These 10 families comprise approximately 614 extant species, the majority of which are found in the family Plethodontidae (Baitchman & Herman, 2014; Blackburn & Wake, 2011; Pyron & Wiens, 2011). Most of what we know about limb regeneration come from studies of relatively few species, with Ambystoma mexicanum associated with the majority of citations in the literature, followed next by Notophthalamus viridescens and other salamandrids (Pleurodeles waltl, Cynops pyrrhogaster, Cynops orientalis). There have been relatively few limb regeneration studies of the speciose plethodontid group and other remaining families (Table 1). In general, comparative studies require well-resolved phylogenies and outgroups from which to identify shared derived and unique traits, and identify examples of homoplasy (a shared trait observed among a group of species but not their common ancestor), which is rampant in salamanders (Mueller, Macey, Jaekel, Wake, & Boore, 2004; Parra-Olea & Wake, 2001; Sanderson & Hufford, 1996; Wake, 1991; Wiens, Chippindale, & Hillis, 2003). Indeed, one of the foremost authorities of salamander morphological evolution remarked that it is difficult to find a single trait that can be conclusively demonstrated to have evolved as a shared-derived trait (Wake, 2009). What this means is that many traits in salamanders have evolved independently within taxa, including the number of digits on limbs, digit morphology, and webbing between digits. There is no reason to suspect that molecular and genetic traits will be any less homoplasious and thus great caution should be taken in drawing conclusions about the evolution of regeneration traits in absence of genomic data and robust phylogenetic hypotheses. While there is little debate about salamander species assignments to families, new species continue to be described and some speciose groups, like Bolitoglossa in South America, have received relatively little systematic attention. Thus, although molecular phylogenies have helped resolve family level relationships (Pyron & Wiens, 2011; Zhang & Wake, 2009), there is still need to perform phylogenetic studies of additional species in support of evolutionary developmental studies at lower taxonomic levels.

TABLE 1.

Summary of regenerative potential across salamanders, as reported in the literature

Family Species Regen. Reference Staging NGS data TALENs CRISPR
Ambystomatidae Ambystoma mexicanum Normal Tank, Carlson, & Connelly 1976; Thoms & Stocum 1984 Faber, 1960; Tank, Carlson, & Connelly, 1976 Genome: Nowoshilow et al., 2018; Smith et al., 2019; Transcriptome: Stewart et al., 2013; Wu et al., 2013; Bryant et al., 2017; Caballero-Pérez et al. 2018; Dwaraka, Smith, Woodcock, & Voss, 2018; Gerber et al., 2018; Leigh et al., 2018; Proteome: Rao et al., 2009 Khattak, Richter, & Tanaka, 2009; Khattak et al., 2013; Flowers et al., 2014; Kumar, Gates, Czarkwiani, & Brockes, 2015; Kuo et al., 2015; Khattak et al., 2013; Flowers et al., 2014; Kumar, Gates, Czarkwiani, & Brockes, 2015; Kuo et al., 2015 Fei et al., 2014; Fei et al., 2016; Fei et al., 2017; Flowers, Sanor, & Crews, 2017
Ambystoma andersoni Normal S. Randal Voss (unpublished) – Transcriptome: Dwaraka, Smith, Woodcock, & Voss, 2018 – –
Ambystoma tigrinum Normal Borgens, Vanable, & Jaffe 1979; Scadding, 1981; Young, Bailey, & Dalley, 1983a; Young, Bailey, Dalley, 1983b; Scadding, 1981; Young, Bailey, & Dalley, 1983a; Young, Bailey, Dalley, 1983b Young, Bailey, & Dalley, 1983a Transcriptome: Eo et al., 2012 – –
Ambystoma maculatum Normal Stocum, 1979; Scadding, 1981 Stocum, 1979 Transcriptome: Burns et al., 2017; Dwaraka, Smith, Woodcock, & Voss, 2018 – –
Ambystoma laterale Normal Scadding, 1977 – Transcriptome: McElroy et al., 2017 – –
Ambystoma jeffersoniaum Normal Scadding, 1977 – – – –
Ambystoma opacum Normal Towle, 1901; Scadding, 1981; Scadding, 1981 – – – –
Ambystoma texanum Normal Young, Bailey, & Dalley, 1983a; Young, Bailey, Dalley, 1983b Young, Bailey, Dalley, 1983b Transcriptome: McElroy et al., 2017 – –
Ambystoma annulatum Normal Young, 1977; Young, Bailey, & Dalley, 1983a; Young, Bailey, Dalley, 1983b Young, Bailey, Dalley, 1983b – – –
Salamandridae Cynops pyrrhogaster Normal Inoue, 1956 – Transcriptome: Nakamura et al., 2014; Casco-Robles et al., 2018; Proteome: Tang et al., 2017 Nakajima, Nakajima, & Yaoita, 2016 –
Triturus cristatus Normal Spallanzani, 1768; Towle, 1901; Smith & Wolpert 1975; Smith, Lewis, Crawley, & Wolpert, 1974 – – – –
Lissotriton helveticus Normal Koussoulakos, Kaparos, & Stathakos, 1986; Newth, 1958 – – – –
Lissotriton vulgaris Normal Schwidefsky, 1934; Lleureux, Thoms, & Carey, 1986 – – – –
Ichthyosaura alpestris Normal Schwidefsky, 1934; Niwelinski, 1958; Koussoulakos, Sharma, & Anton, 1988 – – – –
Salamandra salamandra Normal Roguski, 1961; Skowron, 1962 – – – –
Notophthalmus viridescens Normal Prichett & Dent, 1972; Iten & Bryant, 1973; Tassava, Castilla, Arsanto, & Thouveny, 1993 Iten & Bryant, 1973; Singer, 1952; Singer, 1952 Transcriptome: Abdullayev et al., 2013; Looso et al., 2013; Proteome: Abdullayev et al., 2013 – –
Pleurodeles waltl Normal Lleureux, Thoms, & Carey, 1986; Tassava, Castilla, Arsanto, & Thouveny, 1993; Zenjari et al., 2003 – Genome: Elewa et al. 2017; Transcriptome: Elewa et al., 2017 Hayashi et al., 2014 Elewa et al., 2017; Suzuki et al., 2018
Taricha torosa Normal Chan & Balek, 1973 – – – –
Plethodontidae Bolitoglossa ramosi Normal Gómez, Molina, Zapata, & Delgado, 2017 – Transcriptome: Goodwin, 1946 – –
Bolitoglossa vallecula Normal Gómez, Molina, Zapata, & Delgado, 2017 – – – –
Bolitoglossa rufescens Normal Sessions & Larson, 1987 – – – –
Plethadon cinereus Normal Towle, 1901; Dinsmore & Hanken, 1986; Scadding, 1981 – – – –
Hydromantes italicus Normal Sessions & Larson, 1987 – – – –
Hydromantes platycephalus Normal Sessions & Larson, 1987 – – – –
Pseudotriton ruber Normal Towle, 1901 – – – –
Pseudoeurycea bellii Normal Sessions & Larson, 1987 – – – –
Pseudoeurycea leprosa Normal Sessions & Larson, 1987 – – – –
Thorius spp. Normal Sessions & Larson, 1987 – – – –
Plethadon dorsalis Normal Scadding, 1981 – – – –
Plethadon glutinosis Normal Scadding, 1977; Sessions & Larson, 1987 – – – –
Plethadon cinereus Normal Towle, 1901; Scadding, 1981; Dinsmore & Hanken, 1986; Sessions & Larson, 1987 – – – –
Plethadon dunni Normal Sessions & Larson, 1987 – – – –
Plethadon elongatus Normal Sessions & Larson, 1987 – – – –
Plethadon jordani Normal Sessions & Larson, 1987 – – – –
Plethadon larselli Normal Sessions & Larson, 1987 – – – –
Plethadon richmondi Normal Sessions & Larson, 1987 – – – –
Plethadon vehiculum Normal Sessions & Larson, 1987 – – – –
Plethadon welleri Normal Sessions & Larson, 1987 – – – –
Plethadon yonahlossee Normal Sessions & Larson, 1987 – – – –
Desmognathus monticola Normal Sessions & Larson, 1987 – – – –
Desmognathus ochrophaeus Normal Towle, 1901; Scadding, 1981; Sessions & Larson, 1987 – – – –
Desmognathus quadramaculatus Normal Sessions & Larson, 1987 – – – –
Desmognathus wright Normal Sessions & Larson, 1987 – – – –
Desmognathus fuscus Normal Jeanny & Gontcharoff, 1974; Scadding, 1981 – – – –
Euryacea bislineata Normal Scadding, 1977; Scadding, 1981; Sessions & Larson, 1987; Scadding, 1981; Sessions & Larson, 1987 – – – –
Eurycea guttolineata Normal Towle, 1901 – – – –
Euryacea quadridigitata Normal Towle, 1901 – – – –
Aneides ferreus Normal Sessions & Larson, 1987 – – – –
Aneides flavipunctatus Normal Sessions & Larson, 1987 – – – –
Aneides hardii Normal Sessions & Larson, 1987 – – – –
Aneides lugubris Normal Sessions & Larson, 1987 – – – –
Esatiana eschscholtzi Normal Sessions & Larson, 1987 – – – –
Protidae Necturus maculosus Absent or Heteromorphic Towle, 1901; Scadding, 1977; Scadding, 1977 – – – –
Sirenidae Siren intermedia Absent or Heteromorphic Scadding, 1977 – – – –
Amphiumidae Amphiuma tridactylum Absent or Heteromorphic Scadding, 1981 – – – –
Amphiuma means Absent or Heteromorphic Towle, 1901; Morgan, 1903 – – – –
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Note: The third column identifies if regeneration (Regen.) was reported as Normal or inconsistent (Absent/Heteromorphic). Literature references are provided for regenerative outcome, species-specific regeneration staging schemes, NGS data, and usage of TALENs and CRISPR gene editing tools.

Abbreviations: CRISPR, clustered regularly interspaced short palindromic repeats; NGS, next generation sequencing; TALEN, transcription activator-like effector nucleases.

Fossil data for a presumptive ancestor of modern day salamanders, Micromelerpeton crederni, show limb abnormalities akin to those observed in extant salamanders that experience injuries in nature (Fröbisch, Bickelmann, & Witzmann, 2014; Fröbisch, Bickelmann, Olori, & Witzmann, 2015). This suggests that appendage regeneration may have been an ancient feature of dissorophoid tetrapods, predating the origin of modern salamanders (Fröbisch et al., 2014). Of the existing amphibian groups, salamanders more closely share the body form of their amphibian ancestors, whether they be temnospondyls or lepospondyls (Anderson & Jenkinson, 2007; Anderson, Reisz, Scott, Fröbisch & Sumida, 2008; Bolt, 1969). Most modern salamanders have four morphologically similar limbs, elongated bodies, and prominent tails, traits that define stem tetrapod anatomy. In contrast, caecilians and anurans show greater divergence in body form. For example, the tail is retained across the metamorphic boundary in salamanders but regresses in anurans. Moreover, anurans develop specialized hind limbs and a compacted vertebral column to facilitate a jumping mode of locomotion not seen in salamanders or caecilians (Calow & Alexander, 1973; Duellman & Trueb, 1994; Zug, 1972). In regard to caecilians, all species are limbless and present adaptations for a fossorial lifestyle. The fossil record further suggests that stem tetrapods and salamanders likely inhabited similar environments and presented similar life histories (Pierce, Clack, & Hutchinson, 2012). Thus, salamanders seem to have retained more ancestral tetrapod traits than anurans and caecilians, including possibly their mode of limb development and ability to regenerate appendages. For example, the salamander limb bud differs morphologically relative to tetrapods, as the apical epidermal ridge (AER) lacks a distinct thickening of the distal limb bud epithelium (Tank, Carlson, & Connelly, 1977). Following limb bud establishment, skeletogenesis commences similar to that of amniotes and anurans (Shubin, Sibirtsev, Rasskazov & Bagriantsev, 1986). De novo cartilage condensations arise in a proximodistal orientation with proximal stylopod elements (humerus and femur) forming before distal zeugopodial elements (radius and ulna). However, subsequent zeugopodial and autopodial patterning events differ profoundly between salamanders and all other tetrapods. In particular, distal, pre-axial condensations appear early in the salamander autopod and it is only after the basale commune forms (anterior metacarpal digit 2) that the radius and ulna undergo branching and segmentation (Shubin & Wake, 2003). Subsequent to this, digit formation proceeds in a pre-axial to post-axial direction. This salamander-specific mode of digit formation (II-I-MI-IV-V) is referred to as pre-axial development and contrasts with the post-axial mode of development observed in all other tetrapods (e.g. IV-(V)-III-II-I-digit formation). Fossil limbs of the branchiosaurid Apateon show stages of pre-axial development (Fröbisch, Carroll, & Schoch, 2007), thus providing further support for the idea that salamanders have conservatively retained characteristics of ancestral tetrapods.

It is probably too simple to think that salamanders are capable of limb regeneration because they retained more characteristics of a regeneration competent ancestor than anurans and caecilians, or because of their pre-axial model of limb development. Salamanders as a group exhibit considerable variation in life history, development, physiology, and morphology (Bonett & Blair, 2017), perhaps more variation than is observed in other tetrapod groups (Blanco & Alberch, 1992; Franssen, Marks, Wake, & Shubin, 2005; Vorobyeva & Hinchliffe, 1996, reviewed in Fröbisch, 2008; Fröbisch & Shubin, 2011; Shubin & Wake, 2003). This includes variation in limb morphology and digit number. Relative to the majority of salamanders, species of the family Sirenidae lack hind limbs and produce less digits, and species of the family Amphiumidae exhibit reduced fore- and hindlimb length. Some aspects of limb development that are associated with larval developmental ecology vary more subtly. Larvae adapted for pond habitats (e.g. Ambystoma mexicanum) develop forelimbs many days in advance of hindlimbs, a lag which is less pronounced in stream-developing and direct developing salamander larvae (Blanco & Alberch, 1992; Shubin & Wake, 2003; Vorobyeva, Antipenkova, Kolobayeva, & Hinchliffe, 2000; Wake & Shubin, 1998). The timing of digit development also varies among direct developing salamanders (e.g. families Plethadontidae and Bolitoglossinae). Rather than producing a conical limb bud “pallete” as observed in metamorphic and paedomorphic species (e.g. N. viridescens and A. mexicanum), direct developers form a limb “paddle” which is characterized by nearly simultaneous development of digits, a pattern that is similar to amniote autopod development (Shubin & Wake, 1991; Franssen et al., 2005; Gómez, Molina, Zapata, & Delgado, 2017; Kumar, Gates, Czarkwiani, & Brockes, 2015). Additionally, limb patterning variations have been observed between Plethodon cinereus and other direct developing plethodontids: (a) Pre-axial development of digits is continuous with the post-axially developed ulna/fibula, rather than independent (e.g. Desmognathus aeneus, Franssen et al., 2005); (b) There is no observed preaxial dominance among zeugopodial columns; and (c) relative to Bolitoglossa subpalmata, digit II differentiates earlier and digit I differentiation is delayed (Kerney, Hanken, & Blackburn, 2018). Lastly, variation in limb development may vary among populations of a species, as supported by the identification of distinct forelimb skeletal pattern variants among P. cinereus collected from different geographical locations (Hanken & Dinsmore, 1986).

While limb regeneration appears to transcend different modes of salamander development and life history, and variation in limb anatomy, representatives of all species have not been evaluated for limb regenerative potential (Table 1). It would obviously be fantastic to have a non-regenerative species that could be used as a model to re-engineer a limb regeneration response. However, available comparative data mostly support the idea that all salamanders mount a regenerative response following limb amputation (Dearlove & Dresden, 1976; Goodwin, 1946; Wallace, 1981; Young, 1977; Young, Bailey & Dalley, 1983a). Older papers reported species that show either an absence of limb regeneration or significant defects during limb regeneration (Table 1, Morgan, 1903; Scadding, 1981; Towle, 1901). However, some of these species were shown in later studies to regenerate (e.g. Ambystoma maculatum, Ambystoma tigrinum; Stocum, 1979; Young et al., 1983a). For example, Young et al. (1983a) showed that ambystomatids previously reported as exhibiting no or heteromorphic limb regeneration did indeed regenerate when surveyed for a longer period of time. The time to complete regeneration varied by more than 35 days among the three metamorphic species (A. texanum, A. maculatum, A. tigrinum) that were compared. Thus, earlier claims of non-regenerating species may not have anticipated such variability. Additionally, Young et al. (1983a) speculated that earlier studies may have housed animals in groups, thus facilitating bite injuries that can block regeneration and cause heteromorphic limbs (Bryant et al., 2017; Dearlove & Dresden, 1976; Thompson, Muzinic, Muzinic, Niemiller, & Voss, 2014). Finally, the use of older animals by Scadding (1981) may have contributed to the classification of a few animals as non-regenerative, because age is negatively correlated with regenerative potential and fidelity (Monaghan et al., 2014). While it seems likely that most if not all salamanders can regenerate their limbs, studies are needed to reassess purported examples of regenerative failure and also examine species that have not been evaluated for regenerative potential. If regenerative failure were confirmed for Necturus maculosus, Siren intermedia, and either Amphiuma tridactylum or Amphiuma means, it would provide a striking example of homoplasy.

3 |. LIMB REGENERATION – WHAT ASPECTS ARE CONSERVED AMONG SPECIES?

Most of what we know about limb regeneration comes from studies of A. mexicanum and N. viridescens. Limb regeneration in these species is thought to proceed through a stereotypical set of developmental stages (Iten & Bryant, 1973; Tank, Carlson, & Connelly, 1976). Upon limb amputation, hemostatic processes stop bleeding at the area of injury within minutes (Hay & Fischman, 1961) and innate immune responses are enacted to mitigate pathogens. Re-epithelialization soon follows, in which basal cells of the epidermis migrate from the edge of the wound and cover exposed tissues of the limb stump. The wound is initially covered by a thin wound epithelium (WE; Repesh & Oberpriller, 1980) approximately one layer thick. The formation of the WE is critical for proper regeneration, as grafting skin in the place of the WE (Mescher, 1976), or repeated excision of the WE surface (Thornton, 1957) results in failure to regenerate. Following re-epithelialization, stump tissues undergo histolysis and this breaks down components of the injured tissue such as the extracellular matrix (ECM), muscle, and bone to seemingly provide space for regenerating tissue to form (Wallace, 1981). The WE continues to thicken to form the apical epithelial cap (AEC). The AEC secretes factors to the underlying blastema, a transient structure composed of progenitors of different tissue lineages (Currie et al., 2016; Kragl et al., 2009). The formation of the blastema is an important feature of regeneration as it develops and gives rise to different tissues of the regenerated limb. The blastema develops under the influence of neurotrophic factors and other signals provided by nerves at the site of injury (Brockes, 1984; Farkas, Freitas, Bryant, Whited & Monaghan, 2016; Satoh, Graham, Bryant, & Gardiner, 2008; Singer, 1978). Indeed, transection of various nerves (e.g. dorsal root ganglion, sciatic nerves) interferes with blastemal outgrowth and successful salamander limb regeneration is dependent on having a minimum number of nerve fibers to regenerate (Scadding, 1983; Singer, 1952). Once the blastema achieves critical size, the bulbous structure flattens and the blastema exhibits outgrowth. Differentiation initiates in a process not unlike that of embryogenesis and digits are redeveloped pre-axially (Grim & Carlson, 1974). Digit outgrowth initiates as the limb completes patterning and undergoes growth to attain the correct size, although as mentioned above the time to complete regeneration can vary as a function of size and age. It is important to note that the WE, AEC, and blastema have been identified in multiple salamander species, suggesting these hallmarks are conserved during limb regeneration (Gómez et al., 2017; Inoue, 1956; Kato et al., 2003; Niwelinski, 1958; Roguski, 1961; Sessions & Larson, 1987; Smith, Lewis, Crawley, & Wolpert, 1974; Stocum, 1979; Tassava, Castilla, Arsanto, & Thouveny, 1993; Young et al., 1983a; Zenjari et al., 2003).

The efficiency of RNA-Seq makes it an especially powerful tool to identify shared and divergent gene expression signatures in comparative studies. Indeed, a recent RNA-Seq analysis (Dwaraka, Smith, Woodcock, & Voss, 2018) identified 405 genes that were commonly differentially expressed in larvae of three salamander species (A. mexicanum, A. andersoni, and A. maculatum) at 24 hr post-limb amputation (Dwaraka et al., 2018). Many of the genes identified in this study were identified previously in earlier studies of A. mexicanum limb regeneration (e.g. Voss et al., 2015), and thus this group of genes represents an especially robust set for comparative studies. Moreover, some of the genes identified as differentially expressed during limb regeneration in B. ramosi exhibited similar temporal profiles to those in A. mexicanum (Gómez, Woodcock, Smith, Voss, & Delgado, 2018), further suggesting conservation of gene expression across salamander families. The genes that were identified indicate a diverse, transcriptional response to injury, involving genes that function in both wound healing and tissue development. Upregulated genes were associated with tissue remodeling (mmp3, mmp28, timp1), keratinocyte differentiation (krt5, krt12, col4a1, thbs1), cell cycle regulation (ckdn1a, tp53inp1), and transcriptional regulation (cyr61, areg, marcks, spi1, klf2, klf3), and downregulated genes were associated with muscle tissue (myh3, myh6, myhl3, tpm1) and cellular metabolism (cox4i2, dld uqcrc2, uqcrfs1). Decreased, muscle gene expression has been observed in several studies and it may be associated with the histolysis and dedifferentiation of muscle tissue in the area of the limb stump where the blastema forms. Indeed, muscle tissue transcripts decreased to a greater degree in metamorphic A. maculatum relative to the other two paedomorphic species, suggesting a more robust tissue histolysis response, which in turn, correlates with a shorter time to complete limb regeneration (Figure 1). It is interesting to consider that intrinsic and extrinsic variables may regulate the rate of limb regeneration within the constraint of completing metamorphosis, a property that is predicted to show considerable variation within and among species with biphasic life histories. At any rate, muscle genes can be used as transcriptional biomarkers for evaluating species differences in tissue histolysis.

FIGURE 1.

FIGURE 1

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Relative timing and duration of limb regeneration stages among eight salamander species from three different salamander families – Salamandridae (N. viridescens), Ambystomatidae (A. maculatum, A. tigrinum, A. mexicanum), and Plethodontidae (P. dorsalis, D. ochrophaeus, E. bislineata, B. ramosi), Stages from Tank et al. (1976) are represented by color and proceed from time of amputation (0 days post amputation – dpa) to 95 dpa. Species and family level relationships are from Timetree.org. Data were obtained from Iten and Bryant (1973), Tank et al. (1976), Stocum (1979), Scadding (1981), and Gómez et al. (2017)

4 |. LIMB REGENERATION – WHAT ASPECTS VARY AMONG SPECIES?

As we noted at the beginning of this review, homoplasy is rampant in salamanders and thus we should expect to find differences in regeneration mechanisms. However surprisingly, relatively few differences have been discovered to date. In part, this reflects the difficulty in establishing species differences with incomplete molecular toolkits. For example, without complete knowledge of a gene sequence it is difficult to know the specificity of antibodies for homologous proteins and hybridization probes for mRNA. And without genome references, it is difficult to establish in comparative studies that the same genes are being compared, that genes are differentially represented among genomes, or if different gene paralogs are performing analogous functions. However, even at this early stage of genome sequencing, the future for finding species differences seems bright, as evidenced by recent comparative studies of muscle regeneration among N. vridescens, P. waltl, and A. mexicanum. Regenerated muscle tissue in N. viridescens forms from progenitors that arise from dedifferentiated muscle fibers with little contribution from Pax7 + satellite cells. In contrast, Pax7 + satellite cells in the axolotl make the dominant contribution to muscle tissue within the regenerated limb (Sandoval-Guzmán et al., 2014). Interestingly, Tanaka et al. (2016) showed that pre-metamorphic N. viridescens regenerate muscle tissue predominantly from satellite cells, but post-metamorphic individuals regenerate muscle from dedifferentiated muscle fiber cells, as previously reported by Sandoval-Guzman et al. (2014). To determine if cellular dedifferentiation is unique to metamorphs, Tanaka et al. (2016) induced metamorphosis in A. mexicanum by thyroid hormone (TH) injection. They found that muscle tissue regenerated from Pax7 + satellite cells, as was observed for paedomorphic individuals. Although it is not clear why N. viridescens has two different mechanisms for muscle regeneration, the cellular de-differentiation mechanism requires Pax3 in another salamandrid (P. waltl; Elewa et al., 2017), a gene not present in the A. mexicanum genome (Nowoshilow et al., 2018). Clustered regularly interspaced short palindromic repeats (CRISPR)-mediated inhibition of Pax3 during regeneration inhibited formation of muscle tissue in P. waltl during limb regeneration, whereas Pax7-inhibition did not affect muscle composition of the developed limb (Elewa et al., 2017). This seminal body of work clearly established that molecular-level regeneration mechanisms can vary among salamanders. Moreover, it established the need to pursue comparative studies of muscle tissue regeneration beyond Ambystomatidae and Salamandridae.

Analyses of limb regeneration in direct-developing salamanders have further identified limb regeneration differences within and between species. Dinsmore and Hanken (1986) performed amputations of P. cinereus limbs to determine if natural variation in skeletal limb patterning could be attributed to abnormal limb regeneration. They discovered a different patterning mechanism during limb regeneration, with fewer skeletal elements and increased fusions of carpal/tarsal structure (e.g d3-d4 digit fusions) observed in regenerated limbs. Thus, in P. cinereus, patterning observed during limb regeneration is different from patterning during limb development. This suggests either a failure to faithfully recapitulate the limb patterning program or fundamental differences in the regulation of limb development and regeneration. Regardless, such variation is not observed in primary salamander models (e.g. A. mexicanum, N. viridescens) wherein limb regeneration and development are thought to be regulated by similar morphogenic processes (Gerber et al., 2018; Nacu & Tanaka, 2011). Gómez et al. (2017) characterized developmental stages of B. ramosi and B. vallecula limb regeneration and described additional variations observed in relation to staging systems of A. mexicanum and N. viridescens. The rate of regeneration is largely delayed (Figure 1), as the late bud (LB) blastema is formed much later in the Bolitoglossa species (60 dpa) compared to A. mexicanum and N. viridescens (24 dpa; Iten & Bryant, 1973; Tank et al., 1976). In addition, a “pigmented blastema” formed at 40 dpa, a feature that has not been observed during limb regeneration in other salamanders. Moreover, greater collagen production was observed under the wound epidermis in B. ramosi and B. vallecula in comparison to A. mexicanum and N. viridescens. RNA sequencing analysis further supported this morphological difference as genes associated with collagen and ECM formation were upregulated to a higher degree relative to A. mexicanum (Gómez et al., 2018). As bolitoglosines are obligate terrestrial salamanders, increased collagen production may be needed to support body weight movement on land and reduce the negative effects of mechanical abrasion of the blastema (and patterning) during limb regeneration, a hypothesis also proposed by Dinsmore and Hanken (1986) to explain variation in mesopodial structures in regenerating P. cinereus. This hypothesis awaits further studies of terrestrial salamanders.

In the last decade, several regeneration-associated genes have been identified which appear to be unique to the species that they were identified or salamanders more generally. For example, the scaffolding protein Prod1 was first identified in N. viridescens as a GPI-linked protein that affects cell-cell adhesion and migratory properties of progenitor cells to facilitate proximo-distal patterning during limb regeneration; later prod1 was also shown to be required for skeletal patterning during limb development (da Silva, Gates, & Brockes, 2002; Garza-Garcia, Driscoll, & Brockes, 2010; Kumar et al., 2015; Kumar, Godwin, Gates, Garza-Garcia, & Brockes, 2007). prod1 orthologs have been isolated from several salamander species, including A. mexicanum, but not from frogs, fish, or amniotes (Garza-Garcia et al., 2010; Geng et al., 2015). In a study that compared Prod1 biochemical properties, N. viridescens and A. mexicanum orthologs were both shown to activate mmp9 transcription and ERK1/2 Map kinase signaling, presumably via interactions with the EGFR receptor, even though Am-prod1 did not contain a GPI-anchor attachment sequence originally identified in Nv-prod1 (Blassberg, Garza-Garcia, Janmohamed, Gates & Brockes, 2011). Functional analyses of protein domains showed that the GPI-anchor was not essential for Nv-prod1 signaling activity. It was hypothesized that GPI-anchored and GPI-anchorless prod1 orthologs engage the EGFR receptor via different mechanisms to achieve similar cell-signaling and limb patterning outcomes. Studies of additional species are needed to determine if GPI-anchored prod1 mediated signaling is a shared-derived trait of salamandrids or another example of salamander homoplasy, in this case molecular-level functional convergence of a regeneration mechanism (Geng et al., 2015). In general, genomes are known to contain lineage-specific genes that arise via duplication and de novo processes from noncoding DNA. Given the large sizes of salamander genomes and the deep phylogenetic history of salamander families, it stands to reason that many additional lineage-specific genes will be reported in the near future as more salamander genomes are sequenced. It is exciting to think that some of these will encode proteins that function in limb regeneration.

Given the possibility that limb development and regeneration mechanisms may differ within species (Hanken & Dinsmore, 1986), it is interesting to consider the possibility that genetic variation for regeneration traits may exist within amphibian populations. One trait that may show such variation is regeneration rate (Figure 1), which Young, Bailey, and Dalley (1983b) found to vary considerably among different metamorphic ambystomatid species. While the pattern observed in Young’s study is most parsimoniously explained as species-level variation associated with differences in body size, it does suggest a mechanism that may operate among populations of a species that are distributed across different thermal environments. When populations from low and high elevation sites are brought into the lab and reared under common garden conditions, including a common temperature, adaptive differences in developmental rates may be revealed. In other words, high elevation populations that typically develop under low temperature conditions show faster development at a common temperature. This has been termed counter-gradient variation (Levins, 1969) because mechanisms that act to counter the effect of slower development under low temperature conditions are favored. This pattern of phenotypic variation has been documented for organisms (plants and animals) that develop at the mercy of prevailing environmental temperature (Conover & Schultz, 1995). This mode of selection may operate among salamander populations that are distributed across elevational gradients to moderate overall development and perhaps regeneration rate. Whether or not regeneration rate can be decoupled from overall developmental rate is unknown; this could be a focus of comparative studies.

5 |. APPROACHES AND RESOURCES FOR COMPARATIVE STUDIES OF LIMB REGENERATION

New insights about variation in regeneration mechanisms are likely to come from comparative analyses that use omics approaches and highly repeatable treatments to perturb shared pathways. Transcriptomics can reveal temporal changes in the regulation of hundreds-to-thousands of genes from tissues and more recently single cells (Bryant et al., 2017; Caballero-Pérez et al., 2018; Dwaraka et al., 2018; Gerber et al., 2018; Gómez et al., 2018; Leigh et al., 2018; Stewart et al., 2013; Wu, Tsai, Ho, Chen, & Lee, 2013). The temporal dynamics of gene expression change, when layered on top of developmental stages of regeneration, can then be used to generate hypothesis for mechanisms that may differ among species. For example, microarray analysis of the first 28 days of A. mexicanum limb regeneration identified a significant transcriptional burst of genes at 22–24 days post amputation that mapped to the transition between late bud and palette stages (Figure 2). Gene ontology analysis of these differentially expressed genes showed significant enrichment for steroid and cholesterol metabolism, which are essential for limb and skeletal development in mice (Schmidt et al., 2009). This example suggests the requirement of a highly conserved steroidogenic transcriptional regulatory network in skeletal patterning during limb development and regeneration. Similar methods can be used across species to enable salamander-wide comparisons, thereby providing deeper comparative insight about mechanisms of limb regeneration.

FIGURE 2.

FIGURE 2

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A developmental timeline of limb regeneration that integrates morphological and histological data from Tank et al. (1976) and biological processes inferred from a transcriptional analysis of limb regeneration (Voss et al., 2015). Red asterisks indicate intervals of time where punctuations in gene expression were observed during limb regeneration. The timeline spans amputation (A) to complete regrowth of a forelimb, and includes pre-bud blastema (PB), early-bud blastema (EB), mid-bud blastema (MB), late-bud blastema (MB), and palette (P). Gene expression profiles at the bottom of the figure show significantly upregulated genes associated with steroid and cholesterol metabolism at 22–24 dpa

However, it is also important to test single gene candidates like Prod1 and others, such as Marcks-like protein (MLP), a protein that may regulate cell-cycle re-entry (Sugiura, Wang, Barsachhi, Simon, & Tanaka, 2016) and IL8, which may provide immunomodulatory signals that are essential for blastema formation (Tsai, Baselga-Garriga, & Melton, 2019). Genetic and pharmacological methods can be used to perturb homologous genes and mechanisms among species. Already, CRISPR and transcription activator-like effector nuclease (TALEN)-mediated genome editing of tyrosinase has been used to generate albino A. mexicanum, P. waltl, and Cynops pyrrhogaster (Fei et al., 2014; Hayashi et al., 2014; Nakajima, Nakajima, & Yaoita, 2016; Suzuki et al., 2018; Woodcock et al., 2017). These approaches have been extended to test the requirement and necessity of candidate genes for limb development (Elewa et al., 2017; Flowers, Timberlake, McLean, Monaghan, & Crews, 2014; Kumar et al., 2015; Kuo et al., 2015; Nowoshilow et al., 2018) and regeneration (Elewa et al., 2017). Additionally, transgenic approaches have been optimized to genetically label and document progenitor cell contributions to regenerating tissues within limbs (Flowers, Sanor, & Crews, 2017; Kumar et al., 2015; Kuo et al., 2015). Morpholinos and retroviral vectors have also been used to inhibit and overexpress, respectively, candidate genes during limb regeneration (Sugiura, Wang, Barsacchi, Simon, & Tanaka, 2016; Whited et al., 2013). These tools have established, for example, the critical role of kazald1 during limb regeneration; inhibition of kazald1 consistently produced smaller blastemas and overexpression yielded deformed limbs (Bryant et al., 2017). Several drugs have been shown to inhibit signaling pathways and cell populations that are required for limb regeneration in A. mexicanum (Denis et al., 2016; Godwin, Pinto, & Rosenthal, 2013; Lévesque et al., 2007; Sader, Denis, Laref, & Roy, 2019; Yun, Davaapil, & Brockes, 2015). Additionally, many commercially available compounds were recently shown to inhibit A. mexicanum tail regeneration and thus represent potentially useful tools for studies of limb regeneration (Baddar, Chithrala & Voss, 2019; Ponomareva, Athippozhy, Thorson, & Voss, 2015; Voss et al., 2019).

Comparative data need to be organized into accessible databases to facilitate comparative analyses. Additionally, there is need to develop regeneration specific Gene Ontologies that can facilitate systems level analyses of genes across species and regeneration paradigms. To this end, efforts have been made to construct regeneration specific databases that sample within and across regeneration models (King et al., 2018; Nieto-Arellano & Iranzo, 2018; Warner et al., 2018; Zhao, Rotgans, Wang, & Cummins, 2016). For example, the RegenDBase integrates noncoding and protein coding RNA expression data to facilitate the identification of microRNA-mRNA regulatory circuits conserved across species (e.g. axolotl, zebrafish, and mouse) and tissue types (e.g. limb, spinal cord, fin, and heart; King et al., 2018). While resources have been developed for A. mexicanum (e.g. Sal-Site – Smith et al., 2005; Axolotl-omics.com, Nowoshilow et al., 2018; cruzomics.net, Caballero-Pérez et al., 2018; Axolotl Transcriptomics, Bryant et al., 2017), N. viridescens (Newtomics.com, Looso et al., 2013), and P. waltl (iNewt, Matsunami et al., 2019), a centralized salamander database would better facilitate comparative genomic, transcriptomic, and proteomic analyses of salamander limb regeneration, as well as other regeneration paradigms. Additionally, as salamander genome assemblies and gene models improve (Elewa et al., 2017; Nowoshilow et al., 2018; Smith et al., 2019), conservation tracks can be developed for transcriptional and epigenetic data to facilitate discovery of transcription factor binding sites, enhancers, and DNA regulatory elements. Coordination in the development of essential bioinformatic resources will be key to enabling comparative studies of salamander regeneration.

6 |. METHODOLOGICAL CONSIDERATIONS FOR COMPARATIVE STUDIES

While the rationale for studying additional salamanders is clear, each species presents unique challenges to accomplish this goal. To study regeneration in a comparative framework requires laboratory experiments and thus the need to develop species-specific husbandry procedures that ensure animal health and welfare. It is also important to control extrinsic environmental variables and collect data within frameworks that can ensure reproducibility of experimental procedures and results across species. As was mentioned above, one of the most important environmental variables to report is temperature. Salamanders are adapted to different thermal regimes and temperature affects the rate of all biological processes in ectotherms, including regeneration. Another factor that affects regeneration rate is age, which typically co-varies with body size. Older individuals typically regenerate more slowly than younger animals, and regenerative potential and mechanisms can vary across ontogeny as was described above. What this means is that post-injury time, a metric often used in regeneration studies, has limited meaning when comparing individuals within species that differ in size and age, and even less meaning when comparing species that present different rates of development. To study regeneration across species there is need to develop regeneration staging systems that have common developmental points of reference. Tank et al. (1976) developed a morphological system for axolotl forelimb with stages that can be adapted for other species, including the time of initial outgrowth of the limb bud, distinct changes in limb shape, and digit formation. Many of these stages were retained in a staging system proposed for larvae of A. maculatum (Stocum, 1979). However, morphological changes during limb outgrowth are subtle and thus likely to introduce error when comparing limb regeneration among species. Histology provides greater resolution of tissue and cellular events, and the staging systems for A. mexicanum and A. maculatum both include histological figures to resolve the timing of various events, including for example when osteoclasts are present and absent during regeneration. Even more resolution is provided by the overlay of transcriptional, proteomic, and additional omics datasets (Bryant et al., 2017; Gerber et al., 2018; Leigh et al., 2018; Rao et al., 2009; Stewart et al., 2013; Voss et al., 2015) to build integrative models of limb regeneration.

Although biological variation presents a number of hurdles for comparative studies of salamander species, policies that have been enacted to prevent the spread of infectious diseases and protect endangered species present additional hurdles that can make it difficult to do comparative studies. Due to the risk of salamander chytrid fungus (Batrachochytrium salamandrivorans) spreading into ecosystems in the United States, on January 28, 2016, the US Fish and Wildlife Service identified all species known to carry B. salamanderivorans as injurious wildlife under the Lacy Act. The listing includes 201 species of 20 genera: Chioglossa, Cynops, Euproctus, Hydromantes, Hynobius, Ichthyosaura, Lissotriton, Neurergus, Notophthalmus, Onychodactylus, Paramesotriton, Plethodon, Pleurodeles, Salamandra, Salamandrella, Salamandrina, Siren, Taricha, Triturus, and Tylototriton. This policy bans the import of these species into the US and interstate transportation between States, the District of Columbia, the Commonwealth of Puerto Rico, or any territory or possession of the United States of any live or dead specimen, including parts. This proactive policy imposes a necessary layer of regulation to ensure that permits are obtained when using injurious salamander species for zoological, educational, medical, or scientific purposes. Although there is a clearly stated permitting process in this case, the amount of red-tape and time to obtain a permit for the collection and use of animals in research can be considerable. For example, researchers in Colombia face considerable red tape in obtaining permits to work with native salamanders, a process that can take several years. As a result, population genetic and phylogenetic data are lacking to delineate cryptic species that might prove valuable in the study of limb regeneration.

7 |. CONCLUSION

Given the specter of developmental homoplasy in salamander evolution and paucity of comparative data for regeneration, we should expect to find complex patterns of evolution for regeneration mechanisms. Limb regeneration likely occurs in all salamanders but it is not orchestrated in exactly the same way. The few salamanders that have been studied within a comparative context have revealed mechanistic differences in tissue regeneration. With unprecedented advances in the development of genomic, transcriptomic, genetic, and bioinformatic resources for primary salamander models, the time is right to gain new insights about limb regeneration from a broader sampling of salamander species.

ACKNOWLEDGMENTS

RV was supported by the National Institutes of Health (P40OD019794, R24OD010435).

Funding information

National Institutes of Health, Grant/Award Numbers: P40OD019794, R24OD010435

Footnotes

The peer review history for this article is available at https://publons.com/publon/10.1002/jez.b.22902

CONFLICT OF INTERESTS

The submitted work was not carried out in the presence of any personal, professional or financial relationships that could potentially be construed as a conflict of interest.

DATA AVAILABILITY STATEMENT

Data sharing is not applicable to this article as no new data were created or analyzed in this study.

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