All experiments with animals were approved by the Animal Committee of the Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry (Moscow, Russia) and the Animals (Scientific Procedures) Act 1986, and the Declaration of Helsinki (Hollands, 1986). Amputation and injections of Xenopus tails were performed with MS222 anesthesia.

To test the effects of ag1 and agr2 downregulation, conventional morpholino oligonucleotides (MOs), vivo-morpholinos (vivo-MOs) and photo-activating morpholinos (photo-MOs)¹ were used (see Supplementary Figure 1 for MOs structure, specificity and efficiency).

In brief, 4–8 cell embryos were injected in blastomeres, mostly giving rise to the tail bud, with conventional MOs specific to ag1 and agr2 mRNAs (4–5 nl of 0.3 mM MO water solution per blastomere) and incubated at 20–22°C until stages 40–42. After tail amputation, tadpoles were incubated at 20–22°C for 1–7 days. Regenerates of 1–7 days were used for regeneration rate analysis, immunochemistry and qRT-PCR. However this approach has one significant weak point. As Agr genes are very important for early development the injections at 4–8 blastomere stages sometimes did not allow to avoid totally Agr MO influence on early development leading to high percent of abnormal and dead tadpoles.

To minimize the possible early effects of conventional MOs, we temporary inactivated them by the complementary photo-MOs, which contained photo-sensitive bonds cleavable with 365 nm light (GeneTools). As a result, ag1 and agr2 mRNA translation was not blocked until the embryos were illuminated at the desired stage with 365 nm light, which induces cleavage of photo-MOs and the release of anti-sense MOs. The 4–8 cell embryos were injected (4–5 nl) with 0.3 mM solution of the corresponding anti-sense MO mixed with the sense photo-MO in a dilution of 1:1.3 according to the manufacturer’s recommendations. Importantly, all procedures during and after injections were performed under > 560 nm light (we used a red lamp in a dark room). All embryos were then incubated until stage 40–42 in dark conditions. Before amputation, the tadpoles were exposed for 30 min to 365 nm UV light for activation (releasing) of anti-sense MO. After tail tip amputation, they were incubated at 20–22°C in daylight for 1–7 days. Regenerates of 1–7 days were used for regeneration rate analysis, immunochemistry and qRT-PCR. Photo-MOs are extremely effective, but all procedures with them must be performed under red light, which leads to increased run-off of the MO solution as it was necessary to check the flow from the capillary before each injection. As photo-MOs are used together with conventional MO, experiments with them become quite expensive. Unfortunately, it worth noting that nowadays Gene-Tools no longer produces photo-MO.

In the third approach, we injected fresh tail stumps by the anti-sense vivo-MOs, which can penetrate through plasma membrane due to a unique covalently linked delivery moiety. Thus, local injection of vivo-MO at a certain developmental stage allows one to knock-down the target gene at the desired spatio-temporal parameters. After amputation, the anesthetized tadpoles were transferred from a 0.1 MMR solution (Marc’s Modified Ringer’s solution) with MS322 anesthetic to Petri dishes with a 3% agarose layer. For better spreading of vivo-MO, we injected 0.4 mM solution in a mixture with the fluorescent tracer FLD into the notochord and both fins in the direction of tail growth near the amputated edge immediately after amputation. We repeated injections once per day during 1–4 dpa. After blastema formation, we injected the solution into the fins and notochord and the blastema. On 7–8 dpa, tadpoles with both normally and abnormally regenerated tails were counted. Additionally, at 1–4 dpa, the regenerated tails were collected for immunochemistry, and total RNA extraction was carried out for qRT-PCR.

For statistical analysis, all tails for simplicity were divided into only two categories: (1) regeneration was considered complete, when good regeneration, similar to the regeneration in most control tails, was observed; (2) regeneration was considered defective if the tail did not grow at all or some defects were observed, such as non-regenerating or partially regenerating fins and a curved thin notochord, without surrounding fin blade. In all experiments statistical significance was calculated with t-test for independent samples.

Synthetic mRNAs were obtained by in vitro transcription using mMessage Machine SP6 Kit (Ambion) and ag1/agr2-pCS2 plasmids linearized by Not1. For the over-expression experiments, injections (4–5 nl) of the following concentrations of mRNA were used: agr2 (300 ng/μl) and ag1 (300 ng/μl), ag1 + agr2 (150 ng/μl + 150 ng/μl). Experiments were performed according to the previously validated method. In brief, we injected 4–8-cell embryos using the solution of the target mRNA and RDA (Rhodamine-labeled dextran-amine) into the blastomeres of the prospective tail buds. The control tadpoles were injected with a water solution of RDA. After tail amputation in the refractory period (stages 46–47), tadpoles were incubated for 1–7 days. Then, regeneration rates were analyzed on morphological (7 dpa), cellular (2–3 dpa) and gene expression levels (0–2 dpa). In total, 400–500 tadpoles were analyzed in three independent experiments for each of control, ag1 and agr2 mRNA. To justify this method of ag1 and agr2 overexpression, we demonstrated by qRT-PCR that the injected mRNA presented in tips of tadpoles’ tails during the refractory period in amounts several times higher than the mRNA of the endogenous ag1 and agr2 (Supplementary Figure 2).

For in situ hybridization, we used the protocol described by Harland (1991). To obtain antisense dig-RNA probe for cyclin D1, we cloned its cDNA into a pAL2-T vector (Evrogen) and conducted in vitro transcription from the PCR-product with dig-NTPs (Roche) and SP6-RNA-polymerase (Thermo Fisher Scientific).

For immunohistochemistry staining, we used the same protocol as previously described in detail (Ivanova et al., 2018). The following antibodies were used: primary rabbit anti-phosphohistone H3 (Millipore, cat. #DAM1545035) (1:100), secondary anti-rabbit CF568 (Sigma, cat. #SAB4600400 and #SAB4600425) (1:500) and anti-rabbit-FITC (Sigma, Cat. # F9887) (1:100). The results were processed by ImageJ software.²

The DeadEnd Fluorometric TUNEL System (Promega, Cat. #G3250) was used to reveal the apoptotic cells. The detailed protocol was described previously (Ivanova et al., 2015).

During these experiments, we determined the border between the old and regenerating part of the tail taking advantage of the fact that even after 5–7 dpa, the regenerating part has more transparent notochord and less structured muscles.

qRT-PCR was performed and evaluated as described previously in Ivanova et al. (2013). Briefly, for total RNA extraction from the regenerating tail tips (1–4 dpa) and isolation, we used, respectively, an RNA extract reagent (Evrogen) and RNA isolation KIT (Evrogen). About 20–30 tails were used for each sample for total RNA extraction. For both MO injections and controls, we took tail’s’ tip tissues, cutting off a piece of the stump proximally to the amputation level extending at a distance of 1/4–1/5 of the tail width from the amputation level. To equalize the amount of tissue, we usually used 5–10 more tails in experimental samples than in the control ones. The RNA quality and concentration were measured by NanoPhotometer N60 (Implen). The reverse transcription (RT) of purified RNA samples was carried out using the M-MLV reverse transcriptase kit (Evrogen) according to the manufacturer’s guidelines. The qPCR with marker primers (see Supplementary Material) and the qPCR-mix HS SYBR (Evrogen) were conducted on the DTprime 4 qPCR amplifiers (DNA-Technology) with a standard 40-cycle hot start program. The obtained PCR data were calculated using the ΔΔCt method. The geometric mean of expression of ODC and EF-1alpha (housekeeping genes) was used for the normalization of gene expression levels. The normalized PCR signal of the 0 dpa sample was taken as an arbitrary unit (a.u.) in each series. The data for each gene expression were calculated in 3–7 independent experiments.

After amputation of the tips of the tails, the tadpoles were incubated in 50 ml Petri dishes with 0.1 MMR, to which recombinant Ag1 and Agr2 were added once to a final concentration of 3 μg/ml (see Supplementary Materials and Supplementary Figure 3 for the procedure of the purification of the recombinant proteins and testing their integrity at successive days after addition to the medium with tadpoles). The same final concentration of BSA was used in the control experiments.

Previously, we demonstrated that both in the amputated tails and hind limb buds of Xenopus laevis tadpoles, the expression levels of ag1 and agr2 had strongly increased on the first day postamputation (1 dpa), reaching a maximum on the 2 dpa, and gradually decreased afterward (Ivanova et al., 2013). Using the regeneration of the amputated tails as a model, we then decided to verify if such activation of the expression of these two genes is necessary for successful tail regeneration.

To this end, we arranged a series of experiments in which we investigated the effects on tail regeneration of the downregulation of ag1 and agr2, alone or together, provoked by the antisense morpholino oligonucleotides (MO) to their mRNA. To achieve the greatest reliability, we used three different types of MOs: conventional MO, photo-MO and vivo-MO.

We injected the conventional MO into the blastomeres of 4–8 cell-stage embryos, which in most cases gives rise to tadpoles’ tails. To minimize the likelihood of possible early effects of the conventional MO, which could have long-term consequences, thus affecting tail regeneration, we downregulated ag1 and agr2 by an photo-inducible morpholinos using mixture of anti-sense conventional MO with sense photo-MOs (further named photo-MO). Although photo MOs were injected into the early embryos in the same way as conventional MOs, they were inactive almost until the tail amputation, when we activated them by 365 nm blue light (see “Materials and Methods” section for details). Finally, to avoid any manipulation of the embryos until the tadpole stage, we used ag1 and agr2 vivo-MOs, which can penetrate through plasma membrane due to a unique covalently linked delivery moiety. In these experiments, we injected vivo-MOs directly into the tail tips immediately after amputation.

On 4–5 dpa, we scored the regeneration effectiveness in each experimental group, comparing it with that in the groups of the control sibling embryos injected with the control MOs (Figures 1A–E). As a result, we established that whereas in all control (control and control MO) groups there were about 90–95% of normally regenerating tails, the percentage of such tails was dramatically lower in the groups of tadpoles in which ag1 and agr2 were downregulated alone or together. In these groups, the percentage of normally regenerating tails varied from 20 to 38% depending on the type of MOs (Figure 1E). We also confirmed these effects of ag1 and agr2 downregulation by revealing at 2 dpa in the amputated tail tip tissues a strong decrease in the expression of three essential regulators of tail regeneration, fgf20, msx1b and wnt5a (Beck et al., 2003; Lin and Slack, 2008; Figure 1F).

All these results confirm that the activity of ag1 and agr2 during the first days after amputation is essential for tail regeneration.

As was discovered, downregulation of ag1 and agr2 resulted in a significant shortening of tail regenerates or the absence of growth or elongation. To verify if these effects were the result of cell proliferation inhibition, apoptosis activation, or both, we compared cell proliferation and apoptosis in the regenerating tail tips of the control tadpoles and those in which ag1 and agr2 were downregulated by photo- and/or vivo-MOs.

When we had analyzed the mitotic activity using monoclonal antibodies to the specific marker of the S-phase, phosphor-histone 3B, a decrease in the number of mitotic cells in the 1–2 dpa regenerating tails with downregulated ag1, agr2, or both, as compared to the control tails, was detected. Notably, the effect was more pronounced in tails injected with ag1 MOs (Figures 2A–E). In some of the tails with downregulated ag1 or agr2, we observed that the number of dividing cells in the tail area near the amputation plane was much less than in the control tails (compare Figures 2A,A’ with Figures 2B–D’). During early period of regeneration (1–4 dpa), an intensive epithelial cell proliferation covering the injury followed by dedifferentiation and proliferation of blastema cells took place (Tseng and Levin, 2008). In support of the critical roles of ag1 and agr2 for cell proliferation, we also revealed by qRT-PCR a statistically significant decrease in the expression levels of several cell cycle regulatory genes, cyclin D1, cdk4 (cyclin-dependent kinase 4) and cdca9 (cell division cycle-associated 9) (Sampath et al., 2004; Musgrove et al., 2011), in the regenerating tail tissues of tadpoles at 1–4 dpa with downregulated ag1, agr2, or both, as compared to the control vivo-MO tails (Figure 2F).

In addition, we examined changes in the spatial pattern of cyclin D1 expression by in situ hybridization on regenerates with normal and downregulated expression of ag1/agr2. During normal regeneration, cyclin D1 mRNA was clearly detected in the formation of blastemas at 2 and 3 dpa (Figures 3A–B’). This spatiotemporal pattern of cyclin D1 expression correlates well with the data of intensive cell proliferation of dedifferentiated cells in the forming blastema. However, in the tails injected with ag1/agr2 MO, cyclin D1 expression was significantly reduced (Figures 3C–D’).

Thus, we concluded that the activities of ag1 and agr2 are necessary for active cell proliferation in the regenerating tails.

Then, to test if the increased cell death could also give rise to the suppression of tail regeneration in tadpoles with downregulated ag1 and agr2, we investigated patterns of the apoptotic cells in regenerating tadpole tails injected with ag1, agr2, or control vivo-MO, using the TUNEL assay. However, we could not find statistically significant differences in the mean density of apoptotic cells after injections of ag1/agr2 vivo-MO between normally regenerating tails and tails with suppressed regeneration, neither in the regenerating tips themselves nor the regions proximal to the level of amputation (Figures 3E–H and Supplementary Figure 4). These results indicate that the suppression of tail regeneration caused by downregulation of ag1 and agr2 was not the result of changes in the normal intensity of apoptosis.

Despite Xenopus laevis tadpoles, in general, being able to regenerate amputated tails, there is a special refractory period between stages 45 and 47, during which the regeneration ability is temporarily blocked (Slack et al., 2004). While initial causes of this blockage are not completely known, it was shown that critical processes for the earliest steps of regeneration processes such as reactive oxygen species production, activation of the HIF-1α pathway and recruitment of innate immune cells to the injury site (such as macrophages) are downregulated in this period (Fukazawa et al., 2009; Love et al., 2013; Ferreira et al., 2018). Activation of these processes by various experimental cues was shown to be sufficient for the activation of tail regeneration in the refractory period. Moreover, downregulation in this period of several other late regulators of tail regeneration was reported, and the activation of regeneration in case of their overexpression was also demonstrated (Tseng et al., 2010; Kakebeen and Wills, 2019). Notably, we previously demonstrated the same for two proteins, whose genes were lost during evolution in poorly regenerating higher vertebrates: for small GTPase Ras-dva1 and the transmembrane modulator of FGF and purinergic signaling, c-Answer (Ivanova et al., 2018; Korotkova et al., 2019).

To arrange similar testing for ag1 and agr2, we first compared their normal expression dynamics after tail amputation was performed before the refractory period, at stages 40–42, with the expression dynamics when the amputation was performed directly during this period, at stages 45–47. Consistent with our previous results (Ivanova et al., 2013), we found that before the refractory period, the expression of ag1 and agr2 strictly increased by six to nine times on the first and second days after amputation (Figure 4A). At the same time, in the tails amputated during the refractory period, the expression of both these genes remained at a low level during at least 5 dpa (Figure 4A).

Then, we tested whether tail regeneration in the refractory period could be initiated by overexpression of the ag1 and/or agr2. Indeed, when we overexpressed any of these genes in tadpoles by a previously validated method of injecting synthetic mRNA in the tailbud precursor blastomeres of embryos in stage 4–8 blastomeres (Ivanova et al., 2018), we obtained results clearly confirming the ability of the overexpressed ag1 and agr2 to rescue tail regeneration in refractory period (Figures 4B–E). Thus, if in the control groups 85–88% of the amputated tails did not regenerate at all or were with various defects, and only 12–15% normally regenerated, in the groups of tadpoles injected with ag1 mRNA, only 35–45% of tails regenerate with defects or not regenerate and 55–65% regenerated normally. Similar results were obtained for the amputated tails overexpressing agr2 mRNA or a mixture of ag1 and agr2 mRNAs: the corresponding values were 32–54% and 46–58% in the first case and 35–40% and 60–65% in the second (Figures 4C,D).

Importantly, after analyzing tails overexpressing ag1 and agr2, which were amputated in the refractory period, the expression of two regulators of tail regeneration, msx1b and wnt5a, was increased compared to the control amputated tails (Figure 4E). These results indicate that overexpression of ag1 and agr2 is sufficient to induce regeneration in the refractory period leading to the activation of at least some key signaling pathways that normally regulate regeneration.

The analysis of the proliferative status also showed similar results to the normal regeneration increase in the expression of cell cycle markers, cyclin D1, cdk4 and cdca9 and the mitotic index in the amputated refractory tails of tadpoles overexpressing ag1 and agr2 compared with the control ones (Figure 4E and Supplementary Figure 5).

The data obtained suggest that Agr proteins restore regeneration ability during the refractory period by activating, directly or indirectly, mitotic activity and signaling pathways essential for regeneration.

Since in the experiments described above we activated regeneration in refractory tails by injecting ag1 and agr2 mRNAs into early embryos, it remained unclear whether such activation was actually caused by the direct influence of these proteins on the stump cells or whether it was a result of some of their actions in earlier stages. In addition, in these experiments, it was impossible to distinguish which of the two possible modes of action of Ag1 and Agr2 was decisive for the activation of regeneration: from the outside or from the inside of cells. As is known, Agrs can operate either in the endoplasmic reticulum or the Golgi apparatus, where they perform the function of chaperones, changing the conformation of proteins, including, possibly, some signaling factors essential for regeneration, or, they can be secreted from the cell, executing functions of such signaling factors themselves (Maurel et al., 2019; Delom et al., 2020).

To determine whether Ag1 and Agr2 could act directly from the outside of cells of the refractory stumps, we arranged experiments in which we treated the stumps with the recombinant Ag1 and Agr2 proteins. In these experiments, we added purified recombinant Ag1 and Agr2, or BSA as a control, in the final concentration of 3 μg per ml (see Supplementary Materials for details of how they were obtained and purified) to the refractory tadpoles (stage 46) kept in 0.1 × MMR, immediately after tail tip amputation. At 2 dpa, the proteins containing the mediums were changed for 0.1 × MMR, and tadpoles were incubated at room temperature until 5 dpa when the regeneration efficiency was scored as compared to control (Figure 4F).

As a result, we detected an evident increase in the tail regeneration frequency in the groups of tadpoles treated by the recombinant Agr proteins. Thus, if there were only 10–15% of regenerating tails in the control group, in the groups of tadpoles treated by Ag1 or Agr2, we revealed 55 and 50% of complete regenerates, respectively (Figures 4G,H). These results indicate that Ag1and Agr2 can activate tail regeneration in the refractory period by influencing the stump cells from the outside.