Molecular genetic system for regenerative studies using newts


Author to whom all correspondence should be addressed.



Urodele newts have the remarkable capability of organ regeneration, and have been used as a unique experimental model for more than a century. However, the mechanisms underlying regulation of the regeneration are not well understood, and gene functions in particular remain largely unknown. To elucidate gene function in regeneration, molecular genetic analyses are very powerful. In particular, it is important to establish transgenic or knockout (mutant) lines, and systematically cross these lines to study the functions of the genes. In fact, such systems have been developed for other vertebrate models. However, there is currently no experimental model system using molecular genetics for newt regenerative research due to difficulties with respect to breeding newts in the laboratory. Here, we show that the Iberian ribbed newt (Pleurodeles waltl) has outstanding properties as a laboratory newt. We developed conditions under which we can obtain a sufficient number and quality of eggs throughout the year, and shortened the period required for sexual maturation from 18 months to 6 months. In addition, P. waltl newts are known for their ability, like other newts, to regenerate various tissues. We revealed that their ability to regenerate various organs is equivalent to that of Japanese common newts. We also developed a method for efficient transgenesis. These studies demonstrate that P. waltl newts are a suitable model animal for analysis of regeneration using molecular genetics. Establishment of this experimental model will enable us to perform comparable studies using these newts and other vertebrate models.


Regeneration, which enables tissue reconstruction after injury, has been studied for many years. Although a lot of research has been performed, the mechanisms of regeneration at the genetic and molecular levels are mostly unclear. In order to study gene function in vivo, molecular genetic analyses are very powerful. It involves gene manipulation, such as transgenesis and knockout, in addition to the analyses of mutants. In particular, it is important to establish transgenic or knockout (mutant) lines, and systematically cross these lines to study the functions of the genes. In fact, such systems have been developed for vertebrate models of regenerative research such as Xenopus, Axolotl (Ambystoma mexicanum; neotenic salamander; belonging to family Ambystoma), and Zebrafish. However, there is currently no experimental model system using molecular genetics for newts (family Salamandridae, subfamily Pleurodelinae; not including Axolotl).

Newts have the remarkable ability to regenerate lost tissues, such as limbs, optical tissues (lens, retina and cornea), brain, spinal cord, intestine, and heart (reviewed in: Brockes & Kumar 2002; Agata & Inoue 2012). Of the vertebrate models mentioned above, only newts are able to regenerate all of these organs, and the regeneration is observed throughout their life even after metamorphosis. Thus, establishment of molecular genetic systems using newts would represent valuable experimental models for studies involving regeneration, stem cells, and reprogramming. However, newts, which are commonly used for regenerative research, are not currently suitable as the model animal in molecular genetics due to difficulties encountered with their large-scale breeding. For example, the maturation of sperm and eggs is seasonal, and each female only spawns <10 eggs per spawning in Japanese common newts (Cynops pyrrhogaster) (Makita et al. 1995; Ueda et al. 2005). In addition, more than 3 years are required for sexual maturation of Cpyrrhogaster.

Recently, procedures to produce transgenic newts have been developed using C. pyrrhogaster (Makita et al. 1995; Ueda et al. 2005; Casco-Robles et al. 2011). However, it is extremely difficult to not only obtain the progeny of transgenic newts, but also to establish transgenic lines in these newts due to difficulty with breeding. Therefore, these transgenic newts are not available for analyses at the molecular genetic level. If these disadvantages can be overcome, newts will be an excellent animal model. We looked for other newts that are easy to breed, and the Iberian ribbed newt (Pleurodeles waltl) was chosen as a potential candidate.

In the present study, we have confirmed that P. waltl spawned fertilized eggs all year around in the laboratory. Each female laid more than 150 eggs per spawning and they spawned every 2 weeks. We established methods for artificial insemination and I-SceI mediated transgenesis using these newts. In addition, we were able to shorten the period for sexual maturation from 18 months to 6 months, and obtained F1 transgenic newts. These results indicate that P. waltl newts are suitable as a model animal for studying tissue regeneration using molecular genetics.

Materials and methods


Iberian ribbed newts (Pleurodeles waltl) were originally purchased from Tao (Chiba, Japan). The P. waltl newts used in this study were raised in our laboratory from these newts. The developing stages (st.) were defined according to a criteria described Shi & Boucaut (1995). Adult Japanese common newts (Cynops pyrrhogaster) were purchased from Hamamatsu Seibutsu Kyozai (Shizuoka, Japan). All animals were kept in a tap-water aquarium at 25–26°C under natural light cycles. The animals were fed more than five times in a week; hatched brine shrimp for larvae and combined feed for juvenile/adult newts (Kyorin Corporation, Hyogo, Japan). All procedures were carried out in accordance with the Institutional Animal Care and Use Committee of Tottori University (Tottori, Japan) and national guidelines of the Ministry of Education, Culture, Sports, Science & Technology in Japan.

Induction of regeneration

The newts were anesthetized with 0.1% MS222 (Sigma). The lenses were removed through a horizontal incision in the cornea to induce lens regeneration. The fore limbs were amputated at the lower arm level, and the animals were kept in the aquarium to allow regeneration. The regenerating limbs were staged according to Iten & Bryant (1973).

Artificial insemination

Sperm collection

Sexually mature male animals (older than 6 months postfertilization [p.f.]) were anesthetized with 0.1% MS222. Sperm were squeezed out by pressing the abdomen of mature males and diluted with 1 × Holtfreter's solution to 2000–5000 sperm/μL. The sperm suspension could be stored up to 8 h at room temperature. Sperm was collected from the same animals every 2–3 weeks.

Egg collection

Fourteen to 18 h prior to egg collection, 100–200 IU of human chorionic gonadotropin (ASKA Pharmaceutical, Tokyo, Japan) was subcutaneously injected into the lower jaw of mature females. On the next day, the female newt was transferred into 1 × Holtfreter's solution. When the female started laying eggs, the eggs were collected in a dry Petri dish (100 mm) and the insemination was immediately started. Eggs laid in 1 × Holtfreter's solution remained fertile for up to 30 min. We collected the eggs from the same animals every 2–4 weeks.


Holtfreter's solution was carefully removed from the eggs (excess Holtfreter's solution would decrease the efficiency of fertilization). Eggs were inseminated with diluted sperm (5 μL per 20 eggs) and gently mixed using pipette tips. Inseminated eggs were incubated at room temperature for 15–30 min. Twenty five percent Holtfreter's solution was poured into the dish, and then the eggs were incubated for a further 15–30 min to allow water absorption by the jelly.


The fertilized eggs were treated with 2% cysteine (pH 7.4–7.6) or 2% sodium thioglycolate (pH 7.9) in 0.25 × Holtfreter's solution for 5 min to remove the jelly. De-jellied eggs were rinsed in 0.25 × Holtfreter's solution and transferred into injection medium. The eggshells were removed using fine forceps (this step could be skipped when the eggs were de-jellied with sodium thioglycolate) and stored at 8–10°C until the microinjection.

I-SceI vector (Fig. 4d), in which CAGGS promoter drives GFP, was used for optimization of amount of plasmid DNA in P. waltl. CAGGS promoter construct was provided by Dr J. Miyazaki (Osaka University). For inducible or tissue specific expression, the CAGGS promoter was replaced into Xenopus heat shock protein 70 (HSP 70) promoter (Wheeler et al. 2000) or Xenopus α-Myosin heavy chain (α-MHC) promoter (Garriock et al. 2005).

The plasmid solution digested with I-SceI (NEB) was filled into a micro glass needle. Microinjection was performed using Nanoject II (Drummond). The injected eggs (embryos) were incubated at 8–10°C for 4–6 h to delay first cleavage, and then kept at 16°C overnight. The next day, the temperature was slowly returned to 25°C. The embryos (morula-stage) were rinsed three times in 0.25 × Holtfreter's solution to remove Ficoll contained in the injection medium. The embryos were incubated in 0.25 × Holtfreter's solution for 5–7 days (hatching-stage) and then transferred into tap water.


10 × Holtfreter's solution stock: 600 mmol/L NaCl, 7 mmol/L KCl, 24 mmol/L NaHCO3. Autoclave.

1 × Holtfreter's solution: Dilute 10 × Holtfreter's solution stock to 1 × with distilled water. After autoclaving, add 1.7 mmol/L CaCl2, 1.7 mmol/L MgSO4 and 5 mmol/L HEPES (pH 7.4).

0.25 × Holtfreter's solution: Dilute 10 × Holtfreter's solution stock to 0.25 × with distilled water. After autoclaving, add 1.7 mmol/L CaCl2, 1.7 mmol/L MgSO4 and 5 mmol/L HEPES (pH 7.4). This solution was supplied with 50 μg/mL gentamicin.

Injection medium

0.25 × Holtfreter's solution with 6% Ficoll 400 and 50 μg/mL gentamicin. Sterilized by filtration.

Plasmid solution

One-hundred to 400 ng of plasmids were digested in 20 μL of I-SceI solution (1 × I-SceI buffer, 23 units I-SceI, do not use bovine serum albumin (BSA)) at 37°C for 40 min (critical). The digested plasmids should be stored on ice, and used as soon as possible.


Growth rate of Pleurodeles waltl

Pleurodeles waltl have been bred at 20°C, and they required 18 months for sexual maturation (Chardard et al. 1995; Flament et al. 2009; Dumond et al. 2011). To shorten the period for sexual maturation, it is important to accelerate their growth. We raised the temperature to 26°C and observed the newts actively ate the feed, and the growth rate was increased. However, when the temperature exceeded 28°C, they showed decreased appetite and ulceration of the skin. Therefore, we kept the temperature at 25–26°C. The fertilized eggs hatched after 6–8 days incubation, and the larvae started feeding 12–14 days p.f. At 2 months p.f., the mean body size was 39.3 ± 6.8 mm (mean ± standard deviation [SD], n = 57) and 13.8% of larvae had metamorphosed (Fig. 1). Almost all animals (94.5%, n = 52/55) had metamorphosed within 3 months p.f. Body size increased rapidly up to 8 months p.f., following which the growth rate decreased after 9 months p.f. The survival rate of the P. waltl newts through the experimental period was 77.2% (44/57), and the 43.2% of these animals were the males (19/44). The adult males characteristically have a prominent cloaca compared to the female newts (Fig. 2). In contrast, the mean size of Cpyrrhogaster at 8 months p.f. was 33.4 ± 6.3 mm (mean ± SD, n = 34) (Fig. 1). Under the conditions defined in this study, we obtained sexually mature animals after 6 (male) or 9 (female) months p.f. Mature animals (Fig. 2) showed mating behavior, and the female of P. waltl spawned 150–600 eggs (Fig. 1 and Table 1). Although spontaneous spawning was rarely observed during summer (July–September), fertilized eggs were obtained after a single injection of gonadotropin throughout the year.

Table 1. Breeding properties comparable to Xenopus tropicalis and Axolotl were obtained in Pleurodeles waltl
   P. waltl X. tropicalis Axolotl
  1. Xenopus data is modified from Kashiwagi et al. 2010. Axolotl data is modified from Armstrong & Malacinski 1989.

Reproductive maturation6 months∼5 months∼12 months∼
9 months∼6 months∼12 months∼
Adult size12 cm∼4–4.5 cm15 cm∼
14 cm∼5–6 cm15 cm∼
Egg size 1.2–1.5 mm0.7–0.8 mm1.85–2.0 mm
No. eggs per spawning 150–600 (3000∼/year)1000–9000600–1600 (∼3500/year)
Figure 1.

Pleurodeles waltl newts exhibited greater growth speed compared to Cynops pyrrhogaster. Individual total body length of P. waltl and Cpyrrhogaster are shown as the plots. Data for Cpyrrhogaster was only corrected on 8 months postfertilization (p.f.) Horizontal bars indicate the mean. Dashed line indicates the mean body length of adult Cpyrrhogaster used in our laboratory (105.1 ± 7.6 mm, mean ± standard deviation (SD), n = 134).

Figure 2.

Matured Pleurodeles waltl newts. (a) Dorsal view of matured female (top) and male (bottom) P. waltl newts. (b) Ventral view of their cloacas (female, top; male, bottom). The adult male newt exhibits well-developed cloaca (arrow) compared to the female.

These results demonstrated that breeding properties comparable to those of Xenopus tropicalis and Axolotl were developed in the present study.

Pleurodeles waltl newts have comparable regenerative ability to Japanese common newts

We compared the process of the regeneration of limbs and lenses to that of Cpyrrhogaster.


The process of regeneration was compared between adult Cpyrrhogaster, adult P. waltl, and juvenile P. waltl (Fig. 3a). Four weeks after amputation, regenerating limbs were staged (Iten & Bryant 1973). The regenerating forelimbs of adult P. waltl initiated digit differentiation (early digit stage, n = 5/6). In contrast, the forelimbs of Cpyrrhogaster did not show any sign of digit differentiation (n = 6). Remarkably, the limbs of juvenile P. waltl showed digit differentiation 3 weeks after amputation, and the patterning of the forelimb was almost completed (late digit differentiation stage) in all regenerating limbs (n = 10) after 4 weeks.

Figure 3.

Pleurodeles waltl newts had similar ability to regenerate their forelimb and lens. Regenerating forelimb (a) and lens (b). (a) Black/white arrows indicate regenerating limbs 4 weeks after amputation. The adult Cynops pyrrhogaster. and P. waltl newts were sexually mature. The juvenile P. waltl were immediately postmetamorphosis (3–4 months postfertilization [p.f.]). (b) HE staining image of regenerating lens of adult P. waltl 2 weeks after lens removal. Arrow indicates the dorsal iris.


The lenses were regenerated from the dorsal iris in the same manner as in other species of newts (Fig. 3b). The differentiation of lens-fiber cells occurred within 2 weeks after lens removal in P. waltl. However, Cpyrrhogaster usually required 3 weeks for lens-fiber cell differentiation (Hayashi et al. 2004). P. waltl newts showed an enhanced pace in lens and limb regeneration compared to that of C. pyrrhogaster newts.

Artificial insemination and I-SceI mediated transgenesis

Female of P. waltl lays fertilized eggs like other urodeles (internal fertilization). Since the female newts keep the sperm for several months after mating, we could obtain fertilized eggs without males (Dournon et al. 2001). Eggs obtained just after fertilization were required for efficient transgenesis; however, it is difficult to determine the exact timing of fertilization when it occurs internally. Therefore, we developed a procedure for artificial insemination. For artificial insemination, it is not necessary to kill both females and males (see 'Materials and methods'). The male newts have prominent cloacas (Fig. 2). The males and females were distinguishable when we compared their cloacas. The females were kept separately from the males to obtain unfertilized eggs. The unfertilized eggs were laid in 1 × Holtfreter's solution (Fig. 4a). The sperm were collected by squeezing their abdomen (Fig. 4b). The sperm did not show motility in 1 × Holtfreter's solution until they were stimulated with egg jelly (Mizuno et al. 1999; Watanabe et al. 2011). After mixing the sperm and eggs (insemination), the first cleavage was observed in 5–6 h at 26°C. The fertilized eggs were kept at the single-cell stage by cooling (8–10°C) until microinjection.

Figure 4.

I-SceI mediated microinjection efficiently produced transgenic Pleurodeles waltl newts. (a) A sufficient number of eggs for injection were obtained by administration of gonadotropin. (b) Sperm collected from a matured male P. waltl. (c) Fertilized eggs of P. waltl (arrows) and Cynops pyrrhogaster (arrowhead). (d) Plasmid construct for transgene expression which has two I-SceI recognition sites (light-blue boxes). (e) F0 transgenic larva expressing CAGGS promoter driven green fluorescent protein (GFP) in the whole body. (f) F1 siblings derived from mating with F0 transgenic male transmitting (CAGGS promoter driven GFP) plasmids, and wildtype female (non-transgenic). Arrows indicate GFP-positive st. 30 embryo. Bright field views (e, f top) and dark field views (e, f bottom) of the same larvae.

In the artificial insemination, 73.0% (281/385) of eggs developed to the blastula stage (st. 5–7), and then 71.2% (200/281) of these blastula embryos hatched normally. On the other hand, 88.0% (365/415) of the eggs developed to the blastula stage, and 85.2% (311/365) of these embryos hatched normally during natural spawning. There was no difference in terms of the pace and morphology in their development between the artificially inseminated embryos and the naturally spawned embryos.

The properties of the eggs (size, jelly and egg shell) were similar to the eggs of Xenopus rather than Cpyrrhogaster (Fig. 4c). Basic methods developed for Xenopus transgenesis (Ogino et al. 2006; Pan et al. 2006) were used with some modification for P. waltl. The conditions of microinjection were optimized using a reporter construct of CAGGS promoter driven GFP shown in Figure 4d. The efficiency of transgenesis was estimated by GFP expression in swimming larvae 2 weeks after injection (Fig. 4e and Table 2). The injection volume did not affect the results when using 9.2–18.4 nL (data not shown). The amount of plasmid injected into each egg was tested at three levels (110, 220 and 440 pg/egg), and we found that 110–220 pg/egg was optimal (Table 2).

Table 2. Optimization of amount of plasmid DNA (I-SceI vector, Fig. 3b) in Pleurodeles waltl
Plasmid (pg/egg)Survival rates (larvae/eggs)GFP expression patternsa
  1. a

    Green fluorescent protein (GFP) expression patterns are shown as numbers of GFP positive animals/survived larvae 14 days after fertilization.

044.7% (17/38)
11026.7% (20/75)55% (11/20)30% (6/20)10% (2/20)
22036.3% (26/66)58% (14/24)4% (1/24)21% (5/24)
4400% (0/120)

These transgenic newts started sexual maturation 6 months after fertilization, and we obtained F1 transgenic newts by crossing founder (F0) transgenic and wild type (non-transgenic) newts (Fig. 4f and Table 3). Eighty-eight percent of GFP positive F1 animals shown in Table 3 developed normally. We also revealed that tissue-specific promoters (Xenopus α-MHC promoter, Fig. 5a; Xenopus cardiac actin promoter, data not shown) and inducible promoter (Xenopus HSP 70 promoter, Fig. 5b) successfully worked in newts.

Table 3. Summary of creation of F1 transgenic animals
  1. These 2 (F0) newts designated as Tg1 or Tg2, respectively. Green fluorescent protein (GFP) positive embryos/developing embryos.

Number of F0 transgenic animals examined9
Number of F0 animals transmitting GFP to F1 progenies2 (Tg1 and Tg2)
 Parent; F0 Progeny; F1
Percentages of GFP expressing F1 animalsTg110.5% (6/57)
Tg218.9% (10/53)
Figure 5.

Tissue-specific and inducible expressions of green fluorescent protein (GFP) in Pleurodeles waltl larvae. (a) Cardiac muscle specific GFP expression, which is driven with α-MHC promoter. Bright (top) and dark (bottom) field views of the same larvae (st. 40). (b) Induction of HSP 70 promoter driven GFP expression. The transgenic larva was heat-shocked at 34°C for 8 h. Top and bottom panels show dark field views of the same larva (st. 40) before (top) and after (bottom) heat shock. Inset shows the bright field view.

These data showed that simple artificial insemination and efficient transgenic techniques were developed using P. waltl, and that the F0 transgenic newts transmitted the transgene to their progenies.


An experimental model system for molecular genetics in newts has been needed in order to understand the mechanisms of regeneration from the perspective of gene functions. In this study, we have successfully revealed three critical points as the model animal in molecular genetics using P. waltl newts. (i) We developed breeding properties comparable to those of Xenopus tropicalis. (ii) We have revealed that their ability to regenerate various organs is equivalent to that of Japanese common newts. (iii) An efficient transgenic technique using P. waltl has been developed. These results demonstrated that Pwaltl newts, similar to Xenopus and Axolotl, are suitable as a laboratory animal for the regenerative studies.

Short generation time and convenience in breeding are important factors for suitability as model animals. Under the conditions established in this study, we have been able to increase their growth rate and shorten the period required for sexual maturation from 18 months to 6 months (Chardard et al. 1995; Flament et al. 2009; Dumond et al. 2011). The breeding properties shown in this study are comparable to those of Xenopus tropicalis (Kashiwagi et al. 2010), which is another excellent model for amphibian molecular genetics (Table 1). The number of eggs obtained from each spawning was fewer than that in Xenopus, but P. waltl spawns every 2 weeks. We confirmed that a sufficient number of eggs (embryos) for the experiments or transgenesis were available in all seasons. In addition, we were able to raise adult newts for use in regenerative studies in 6 months. Sex-reversal (female to male) and limb development abnormalities have been reported in P. waltl newts reared at high temperature, for example 30–32°C (Dournon et al. 1990, 1998); however no abnormality was observed in the sex ratio or limb regeneration under our conditions (25–26°C).

Pleurodeles waltl are known for their regenerative ability, like other newts. They regenerate various tissues, for example, lens, retina, tail, limbs, and spinal cord (Lheureux et al. 1986; Arsanto et al. 1992; Becker et al. 1993; Caubit et al. 1993; Mitashov et al. 1995). In this study, we confirmed that lens and limb regeneration in P. waltl progressed at an enhanced pace compared to that of C. pyrrhogaster newts.

In addition, using P. waltl, we can compare the speed of regeneration at different ages in sibling animals from the larva to adult stage, since their date of birth is known and a large number of newts can be obtained as littermates. One definite advantage of this newt is that we can investigate the influence of age factors on regeneration (Eguchi et al. 2011). Because the most newts used for regenerative studies were collected from the field, it was really difficult to know their age. These results indicate that P. waltl newts are very useful for regenerative studies.

Artificial insemination and transgenesis are key techniques in molecular genetics. The procedures developed in this study were simple and sufficient for high-throughput transgenesis in P. waltl are useful for transgenic newts.

In addition, we successfully obtained F1 transgenic animals, the first result in newts, by shortening the period required for sexual maturation.

Establishment of transgenic lines makes it possible to obtain homogenous animals in larger numbers compared to using founder (F0) animals. Moreover, the crossing of transgenic lines enables new experimental designs, for example, genetic label-chase experiments using a tissue-specific CRE-loxP system. It is confirmed that some inducible gene expression protocols including Cre-loxP work in Axolotl (Whited et al. 2012).

Sperm-freezing procedures have been developed for Xenopus and Zebrafish (Sargent & Mohun 2005; Carmichael et al. 2009; Draper & Moens 2009; Mansour et al. 2009). Such a procedure would be helpful to maintain the transgenic lines of P. waltl.

We have demonstrated that P. waltl newts are suitable as laboratory newts for developing a molecular genetic model system for regenerative studies. Establishment of such an experimental model enables comparable studies between these newts and other vertebrate models such as mouse, Axolotl, Xenopus, and Zebrafish.


Kyorin Corporation (Hyogo, Japan) kindly provided the feeds for the newts. We thank Drs K. Agata (Kyoto University), H. Yokoyama (Tohoku University), K. Tamura (Tohoku University), P. Krieg (University of Arizona), S. Hoppler (University of Aberdeen), J. Miyazaki (Osaka University) and H. Ogino (NAIST) for their kind advice and guidance concerning the microinjections and for providing the plasmid vectors. This research was partially supported by a Japanese Ministry of Education, Science, Sports and Culture, Grant-in-Aid for Scientific Research on Priority Areas.