The axolotl (Ambystoma mexicanum) d/d strain used in this study was maintained in fresh water, with individuals housed separately and fed daily. Prior to surgical procedures, axolotls were anaesthetized using a 0.01% solution of ethyl-p-aminobenzoate (Benzocaine; Sigma). All animal experiments were carried out according to the guidelines of the ethics committee of Guangdong Provincial People’s Hospital (approval number KY-Q-2022-395-02).

Pdx1 mutant axolotls were generated using CRISPR/Cas9 technology, following established protocols. The Pdx1 guide RNA (gRNA) sequence (5′-AAG​GAG​GAG​GAC​AAG​AAG​CG-3′) was synthesized by GenScript. A mixture containing 0.4 μg/μL Pdx1 gRNA, 0.5 μg/μL Cas9 protein, and 1× Cas9 buffer was injected into single-cell stage embryos from the d/d strain, as previously described (Fei et al., 2018). The injected embryos developed into juveniles, which were subsequently genotyped to identify the F₀ transgenic animals. Genomic DNA was extracted from axolotl tail snips using a solution containing 50 mM NaOH and 1 M Tris-HCl. The target locus of the Pdx1 gRNA was PCR amplified using primers Pdx1-KOfw1 (5′-act​tga​cga​gct​gtt​cgc​cca​ttt​c-3′) and Pdx1-KOre2 (5′-tga​gca​tgt​cct​tgc​cag​ggt​ccc-3′). The PCR products were then subjected to Sanger sequencing (Tsingke) for alignment with the original Pdx1 sequence. F₀ transgenic axolotls were further bred to generate F₁ offspring with various genotypes (Supplementary Figure S1).

Samples were fixed in 3.7% formaldehyde freshly prepared in MEM buffer (MEMFA) overnight at 4°C. After dehydration in 30% sucrose, the samples were embedded in OCT (Sakura, 4,583). Cryosections of 10 µm were then collected for immunohistochemical analysis as previously described (Fei et al., 2017). The sections were blocked with a blocking buffer (0.3% Triton X-100, 5% fetal calf serum in PBS) for 1 h at room temperature, then incubated with primary antibodies overnight at 4°C. The primary antibodies used in this study were: Rabbit anti-Somatostatin antibody (Abclonal, A9274, 1:200), Guinea pig anti-Glucagon antibody (TAKARA, M182, 1:1,000), Mouse anti-Insulin antibody (Sino Biological, 11038-MM14-4.5, 1:1,000). Following several washes in PBST (0.3% Triton X-100 in PBS), the corresponding secondary antibodies Alexa Fluor 488-Goat anti-guinea pig (Invitrogen, A11073, 1:500), Alexa Fluor 555-Donkey anti-Rabbit (Thermofisher, A-31572, 1:500), Alexa Fluor 647-Donkey anti-Mouse (Jackson, 715605150, 1:500) and DAPI (Sigma, 28718903,1:000), diluted in blocking buffer were applied to the sections and incubated for 2 h at room temperature. The slides were then washed several times in PBST and mounted with Mowiol medium. Immunofluorescence images were acquired using a confocal microscope (ZEISS, LSM980).

Pancreatic tissue from adult axolotls were collected and fixed in MEMFA overnight at 4°C. After dehydration using a graded series of ethanol (50%, 70%, 95% and 100% diluted in PBS) and clearing in xylene, the samples were embedded in paraffin, and 5 μm sections were collected. For H&E staining, the sections were dried at 65°C for 1 h, treated with xylene to remove the paraffin, rehydrated in decreasing concentrations of ethanol, and finally in distilled water. The sections were incubated in hematoxylin solution for 5 min. After several washes with distilled water, the sections were sequentially incubated in 1% hydrochloric acid in ethanol for 10 s, washed gently with running water for 15 min, and then incubated in eosin solution for 1 min. The stained sections were dehydrated with ethanol and transferred into xylene, then mounted with neutral resin. Once the resin dried, the sections were imaged using an inverted microscope (Olympus, IX83).

To synthesize DIG-labelled or fluoresce-labelled antisense RNA probes, corresponding DNA templates containing the T7 promoter were amplified with two rounds of PCR, using axolotl pancreatic cDNA synthesized from total RNA of the pancreas as the template. The antisense RNA probes were transcribed in vitro using a T7 RNA polymerase transcription kit (NEB, M0251L). In situ hybridizations and fluorescence in situ hybridization were performed on 10 μm cryosections from axolotl pancreas or embryos, as previously described (Fei et al., 2017; He et al., 2020). Briefly, for in situ hybridizations, 500 ng/mL of DIG-labeled probes were hybridized at 60°C on 10 μm cryosections overnight. After hybridization, the cryosections were washed with a stringent buffer and incubated overnight at 4°C with anti-DIG-AP antibody (1:5,000, Roche, 11093274910). The signal was detected using BM Purple solution (Roche, 11442074001). For fluorescence in situ hybridization, a mixture of 300 ng/mL DIG-labeled and fluorescein-labeled RNA probes was hybridized at 60°C overnight on 10 μm cryosections. After washing with stringent buffer, the cryosections were incubated with anti-fluorescein-POD (1:500, Roche, 11426346910), and then the signal was detected using TSA fluorescein (1:50, PerkinElmer, NEL701A001KT) at room temperature. Following signal development, 3% hydrogen peroxide was used to deactivate any residual POD, and the cryosections were incubated with anti-DIG-POD (1:1,000, Roche, 11633716001) after washing. TSA Cy5 (1:50, PerkinElmer, NEL705A001KT) was used to generate a detectable signal. List of primers for RNA probe synthesis: Amy-forward: 5′-ATG​AGG​CTC​CTT​CTG​CTG​CTT​GC-3′, Amy-reverse: 5′-CTT​GTA​TTG​TTT​ATA​ACT​TG-3′, T7-Amy-reverse: 5′-TTG​AAA​TTA​ATA​CGA​CTC​ACT​ATA​GGG​CTT​GTA​TTG​TTT​ATA​ACT​TG-3′, Ins-forward: 5′-ATG​GCT​CTT​TGG​GTG​CGT​G-3′, Ins-reverse: 5′ACT​CCT​TCT​ACC​CCC​TCT​GC-3′, T7-Ins-reverse: 5′-TTG​AAA​TTA​ATA​CGA​CTC​ACT​ATA​GGG​ACT​CCT​TCT​ACC​CCC​TCT​GC-3′, Krt19-forward: 5′-ATG​TCG​TCG​AGT​ACA​GCT​TCA​GG-3′, Krt19-reverse: 5′-TTT​TCC​CCC​GCT​TTT​GGA​C-3′, T7-Krt19-reverse: 5′-ATT​ATG​CTG​AGT​GAT​ATC​CCT​CTT​TTT​CCC​CCG​CTT​TTG​GAC-3′, Ptf1a-forward: 5′-TGC​AGT​CCA​TCA​ACG​ACG​CCT​TC-3′, Ptf1a-reverse: 5′-GCG​TCA​CTG​TAA​TGA​AAG​C-3′, T7-Ptf1a-reverse: 5′-TTG​AAA​TTA​ATA​CGA​CTC​ACT​ATA​GGG​GCG​TCA​CTG​TAA​TGA​AAG​C-3′, Mln-forward: 5′-CCC​ACA​TCC​ACA​CTC​TAC​AGT​AG-3′, Mln-reverse: 5′-TCA​GTC​CAC​ATT​TTG​GTG​TTC​TG-3′, T7-Mln-reverse: 5′-TTG​AAA​TTA​ATA​CGA​CTC​ACT​ATA​GGG​TCA​GTC​CAC​ATT​TTG​GTG​TTC​TG-3′.

Axolotl embryos were fixed overnight in MEMFA at 4°C. Following fixation, the embryos were washed with PTW (DEPC-treated 0.1× PBS with 0.2% Tween) through a gradient of decreasing methanol concentrations (100%, 90%, 75%, 50%, and 25%). The samples were then treated with proteinase K (5 μg/mL) at room temperature for 5 min, followed by incubation at 65°C in hybridization buffer with 500 ng/mL antisense RNA probes. To remove unbound RNA probes, the samples were washed multiple times with post-hybridization buffer and MAB buffer. Next, samples were incubated with anti-DIG-AP antibody (1:5,000, Roche, 11093274910) in 1× blocking buffer (Roche, 11096176001). The signal was detected using BM Purple substrate (Roche, 11442074001). Finally, the samples were stored in 50% glycerol diluted in PBS at 4°C and imaged using an Olympus SZX10 microscope.

TEM was conducted following standard procedures. Briefly, the isolated axolotl pancreas was cut into approximately 1 mm³ tissue blocks, which were then transferred to an EP tube containing fresh fixative for TEM (Servicebio; G1102) and fixed overnight at 4°C. After fixation, the samples were rinsed three times with 0.1M phosphate buffer (pH7.4) and post-fixed in 1.0% osmium tetroxide in phosphate buffer at room temperature for 2 h. The tissue was then dehydrated through a graded series of ethanol, embedded in Epon resin, and polymerized at 60°C overnight. Ultrathin sections were cut, mounted on copper grids, and stained with uranyl acetate and lead citrate. All grids were observed and imaged using a transmission electron microscope (Hitachi; HT7800/HT7700).

Total RNA was extracted from the pancreas of larvae (about 3–4 cm, 2 months old) and adult axolotls (about 25 cm, 2 years old) using TRIzol reagent (Invitrogen) and purified via LiCl precipitation. RNA sequencing was conducted on six biological samples, comprising three replicates from both the larval and adult groups. Specifically, three adults and twenty-four larvae were used for sequencing sample preparation. Three micrograms of total RNA were utilized to construct cDNA libraries according to the standard Illumina RNA-seq protocol. The generated cDNA library was sequenced using the Illumina NovaSeq6000 via a 2 × 150 bp paired-end protocol. Quality filtering and removal of residual adaptor sequences was conducted on read pairs using Trimmomatic v0.33. Salmon v1.10.0 was employed to build the axolotl genome index and guide transcript quantification. Axolotl genome assembly v6.0-DD from the axolotl-omics database (https://www.axolotl-omics.org/assemblies) (Schloissnig et al., 2021) served as the reference genome along with annotation files.

Transcript expression levels were imported into R v4.3 and summarized at the gene level using the R/tximport v1.30.0. Gene count data were analyzed for differential expression using the Bioconductor software package DESeq2 v1.42.1. Gene expression was quantified using the transcript per million (TPM) values. Metascape was utilized for gene functional annotation. Mfuzz was applied to analyze the clustering of time series gene expression in larval and adult axolotl pancreas. For conservation analysis, RNA sequence data from untreated human samples (GSE205853) (Benaglio et al., 2022), Wt_2w and Wt_12w group of mice (GSE171251) (de Jesus et al., 2021) were compared to our adult axolotl RNA sequence data. For the identification of homologous genes, we obtained gene annotations from two sources: the human and mouse genome annotations from Ensembl, and the axolotl genome annotation (version 6.0-DD) from the axolotl-omics database. Genes that share annotations with mouse and human were considered homologous genes. Gene length and sequencing depth were standardized by using TPM values to compare homologous gene expression across the three species. |log2Fold Change| ≥ 1 with a False Discovery Rate (FDR) < 0.05 was utilized to capture meaningful changes in gene expression during development for the axolotl and mouse. To identify higher and lower expressed genes in axolotls compared to mice and humans, |log2Fold Change| ≥ 2 and FDR <0.01 were applied. Principal component analysis (PCA) was performed using the prcomp function on TPM-transformed raw counts of the most highly expressed genes across all pancreas samples. Pearson correlation coefficients were calculated to assess the correlations between the three species.

Animals measuring about 15 cm in length were used for GTT and ITT. For the GTT experiment, axolotls were fasted for 12–16 h before being intraperitoneally injected with D-Glucose (2 mg/g) after anesthesia. Blood glucose levels were monitored at specific intervals using a glucose meter (Roche, Accu-Chek), sampling from the punctured dorsal body surface vessel. 48 h after the GTT experiment, the animals were returned to clear water for normal feeding. For the ITT experiment, axolotls were fasted for 6 h before being intraperitoneally injected with recombinant human insulin (insulin glargine; 5 μL/g) after anesthesia. Blood glucose levels were monitored at specific intervals as outlined in the GTT procedure. Similarly, following the ITT experiment, the animals were injected with 2 mg/g glucose solution and returned to clear water for normal feeding; the same concentration of glucose solution was injected for the next 2 days.

All axolotls were randomly assigned to different groups, and all experiments were repeated at least three times independently. For experimental data analysis, calculations were performed using GraphPad Prism 8.0.2. Unpaired, two-tailed Student’s t-tests were performed to analyze the P values for single comparisons between two groups, while one-way ANOVA was conducted to analyze P values for multiple comparisons. All data were presented as the mean ± s. e.m. Each experiment was performed blind, and no predetermination was made for sample size. P value < 0.05 was considered significant. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001; ns, P > 0.05.

To systematically characterize the axolotl pancreas, we isolated pancreas from adult axolotls and examined its morphological features, composition and distribution of pancreatic cell types. Consistent with other vertebrates, the axolotl pancreas is situated beneath the stomach. The pancreas head and tail connect to the duodenum and spleen, respectively, while the body of the pancreas is attached to the jejunum via the mesentery (Figures 1A, B). The uncinate process of the pancreas wraps around the duodenum and connects to the liver and gall bladder (Figure 1C). H&E staining of paraffin sections of the pancreas showed that islets are distributed throughout the pancreatic tissue, surrounded by exocrine tissues, including acinar cells, ducts and blood vessels (Supplementary Figure S2B; Figure 1D). In situ hybridization using classical pancreatic cell markers (Muraro et al., 2016) demonstrated the presence of Ins, a marker for endocrine β cells in islets, and Amylase (Amy) in acinar cells along with Keratin 19 (Krt19) in ductal cells (Figures 1E, F) in the axolotl pancreas.

We next focused on characterizing the endocrine cells. It is well-known that pancreas contains five types of endocrine cells in mammals (Baron et al., 2016). To this end, we used specific endocrine cell markers INS, GCG, and SST to identify β, α, and δ cells respectively. Immunofluorescence on pancreatic sections showed that the main body of the pancreas (head, body, and tail) consists predominantly of INS⁺ β cells-clustered islets, surrounded by sparsely distributed GCG⁺ α cells and closely associated SST⁺ δ cells (Figures 1G, H). In contrast, the uncinate process of the pancreas exclusively contained GCG⁺ α and SST⁺ δ cells, with no INS⁺ β cells (Supplementary Figure S2A; Figure 1G). Additionally, double fluorescence in situ hybridization revealed that pancreatic islets composed of Ins ⁺ β cells were tightly surrounded by a limited number of Motilin (Mln)⁺ ε cells (Figure 1I). Interestingly, immunofluorescence analysis using an antibody against PPY revealed that PPY is expressed exclusively in GCG⁺ α cells (Figure 1J), suggesting the identities of α and PP cells are perhaps intermingled in axolotls. Electron microscopy showed acinar cells containing large electron-dense zymogen granules likely enriched with digestive proteins, while β cells were filled with numerous insulin granules that lacked halos, in contrast to their mammalian counterparts (Roep, 2020; Susan, 2004) (Supplementary Figures S2C, G). In addition, we also found the presence of α and δ cells in the stomach and small intestine of embryonic and juvenile axolotls (Supplementary Figure S3). In summary, axolotl pancreas, although exhibiting some unique feature, demonstrates significant structural, compositional, and subcellular similarities to other species.

We next investigated the developmental origin of the adult pancreas. The differences in endocrine cell types between the uncinate process and the main portion of the axolotl pancreas suggest distinct embryonic developmental mechanism. During mammalian pancreas development, it has been reported that the ventral and dorsal pancreatic anlage give rise to the uncinate process and the main pancreas, respectively (Longnecker, 2014). To determine whether the axolotl pancreas develops from ventral and dorsal buds during the embryonic stage, we performed whole-mount and cryosection in situ hybridization on axolotl embryos. We detected the pancreatic ventral and dorsal buds as early as stage 36, as indicated by the expression of Pancreas associated transcription factor 1a (Ptf1a) and Amy (Figures 2A, D, E). By stage 43, the dorsal bud expanded and elongated in conjunction with the growth of the intestinal tube (Figures 2B, F, G). Eventually, the dorsal and ventral buds fused to form early pancreas a few days following stage 43 (Figures 2C, H, I). Consistent with the cell types identified in adult pancreas, immunohistochemistry analysis revealed that the β cells, δ cells and α cells in the dorsal bud started to develop at stages 40, 42 and 43, respectively (Figure 2J), while only α and δ cells were observed in the ventral bud (Figure 2K). These results indicate that the major body of the pancreas originates from the dorsal bud, whereas the uncinate process arises from the ventral bud, aligning with mammalian pancreas development.

To explore the molecular characteristics of both the developing and adult pancreas, we sequenced and analyzed the gene expression profiles of total RNA extracted from 2-month-old larvae and 2-year-old adult axolotls. Our analysis identified a total of 23,931 genes expressed in the developing pancreas and 19,592 genes in the adult pancreas. Of these, 18,492 genes were commonly expressed at both stages (Supplementary Figure S4A). Gene Ontology analysis of the genes uniquely expressed in the developing pancreas (5,439 genes, with 2,634 annotated genes) revealed their primarily involvement in tissue morphogenesis and digestive system development. In contrast, the genes expressed in the adult pancreas (1,100 genes, with 565 annotated genes) were mainly associated with the positive regulation of the phosphorus metabolic process (Supplementary Figure S4B).

Further analysis on the 18,492 (16,614 annotated genes) commonly expressed genes identified 1,888 upregulated genes (1,238 annotated genes) and 2,016 downregulated genes (1,210 annotated genes) in the adult pancreas compared to the developing pancreas (Supplementary Figure S4C). Gene Ontology analysis indicated that the upregulated genes were primarily linked to hormone responses, secretion regulation, and hormone level control, while the downregulated genes were typically associated with the mitotic cell cycle, DNA metabolic processes, and cell cycle regulation (Figure 3A). Moreover, clustering of time series gene expression shows that genes involved in ribonucleoprotein complex biogenesis and RNA splicing are also highly active, in addition to those related to mitotic cell cycle phase transitions in larvae. Genes associated with signal release, peptide transport, and insulin secretion are even more active in adults (Supplementary Figure S5). These findings suggest a transition of the axolotl pancreas from an actively developing state in early stages to a homeostatic state in adulthood, indicating functional maturation during development, similar to observations in mice (de Jesus et al., 2021) (Supplementary Figures S4D, E).

The development and functional maintenance of the pancreas require coordinated gene expression regulation. We focused on well-known genes that play crucial roles during the development of pancreatic lineages (Aigha and Abdelalim, 2020; Burlison et al., 2008; Yu et al., 2021). The expression levels of markers for most pancreatic multipotent progenitor cells (Pdx1, Ptf1a, Nkx6.1, Nkx2.2 and Foxa2), exocrine progenitor cells (Ptf1a and Sox9), and endocrine progenitor cells (Neurog3, Isl1, Neurod1 and Pax6) were significantly higher in larvae compared to adult axolotls (Figure 3B). Notably, α cell fate determiners Arx, MafB, and Peg10 exhibited comparable expression levels between larvae and adults. In contrast, β cell fate-related genes Gng12 and MafA were expressed at higher levels in larvae. Functional hormone-coding genes in the pancreas, including Ins and Amy, were expressed significantly higher in adults than in larvae, while Gcg, Sst, Ppy and Mln showed lower expression levels. This regulation pattern correlates with increased demands for food digestion and metabolic homeostasis during growth. Overall, the expression patterns and dynamics of key pancreatic genes during development in juvenile and adult axolotls are conserved with those observed in mammals (Yu and Xu, 2020).

To explore the evolutionarily distinctive features of the axolotl pancreas, we compared the transcriptome of adult axolotls with those of humans and mice (Benaglio et al., 2022; de Jesus et al., 2021). Among the 19,592 genes in the adult pancreas, 11,860 were found to have homologous counterparts in both humans and mice. Among these, 6,457 genes were commonly expressed across all three species (Supplementary Figure S4F; Figure 3C). Key pancreatic genes were present in all three species, except for Neurog3, which is absent in both humans and axolotls (Figure 3E). We identified and analyzed differentially expressed genes exhibiting at least four-fold difference in expression levels between axolotls and mice or humans. Our results revealed that 131 genes were expressed at higher levels and 126 genes at lower levels in adult axolotls compared to the other two species (Supplementary Figure S4G). The highly expressed genes were primarily involved in cytoplasmic translation, while the lowly expressed genes were related to hormone transport and negative regulation of cell population proliferation (Figure 3D). Among the 126 downregulated genes, such as Pdx1, Neurod1, Ptprn, Slc30a8, Rbp4, and Sox4, many were associated with insulin synthesis, insulin secretion, and glucose response, indicating that axolotl pancreas may display a delayed insulin secretion response to glucose compared to mammals (Figure 3F). Therefore, we examined the expression of metabolic genes in the axolotl and found that glucose sensor genes Slc2a1 and Slc2a2, insulin secretion genes Stx1a and Vamp8, and glycolytic-related kinases Pfkl, Pfkm, Pkm, Gck, Pgk1, and Ldhb are lowly expressed in the axolotl (Supplementary Figure S4H). Additionally, the genes Slc7a5, Pdx1, Cdkn1a, and Cebpb are known to be downregulated in association with positive effects on organ regeneration (Fitzner et al., 2013; Holland et al., 2005; Juran et al., 2021; Wang et al., 2022). We observed that these genes were expressed at lower levels in axolotls (Figure 3F). In conclusion, the transcriptomic characteristics of the axolotl pancreas indicate an evolutionary conservation of gene expression, suggesting that glucose regulation within pancreatic endocrine cells is less sensitive compared to mammals. Moreover, this may point to a significant regenerative potential within the axolotl pancreas.

Evaluating gene function is crucial for understanding biological processes. Therefore, we targeted Pdx1 to assess the functional conservation and diversity of pancreatic-specific genes. Pdx1 is a transcription factor expressed in multipotent progenitor cells; it is essential for pancreatic development and the maintenance of β cell function (Zhu et al., 2021a; Hui and Perfetti, 2002; Stoffers et al., 1997). In mammals, Pdx1 knockout results in severe, life-threatening pancreatic developmental defects (Fujitani et al., 2006; Jonsson et al., 1994; Schwitzgebel et al., 2003). To investigate this further, we used CRISPR/Cas9 technology to generate Pdx1 mutants. The gRNA was designed to target the region immediately following the nuclear localization signal within the homeobox domain (Figure 4A). F₀ founders were bred with d/d axolotls to produce F₁ heterozygous embryos, in which we identified various insertions or deletions (indels) at the targeted site in germline-transmitted F₁ animals (Figure 4A). Additionally, we performed intercrosses of individual F₀ founders to obtain “homozygous” mutants, considering both alleles with frameshift mutations as “homozygous”.

The “homozygous” mutants survived but predominantly exhibited stunted growth and dwarfism (Figure 4K). Following a 24-h fasting period, we assessed the blood glucose levels of axolotls measuring 7 cm in length, subsequently conducting genotyping. The results indicated that the Pdx1 double allelic mutants, as opposed to heterozygous individuals, exhibited hyperglycaemia compared to the d/d controls (Figure 4B). We then isolated the pancreas from 8-month-old “homozygous” mutants and sibling controls to investigate pancreatic developmental phenotypes. Based on pancreatic morphology, we classified the bi-allelic “homozygous” mutants into two categories: mild (Pdx1 ^m/m ) and severe (Pdx1 ^s/s ) phenotypes. The pancreas of Pdx1 ^m/m mutants showed relatively normal in size and morphology, while that of Pdx1 ^s/s mutants was significantly reduced, consisting only of a dense remnant of the pancreatic head (Figures 4C–E).

mRNA in situ hybridization using an antisense Amy probe revealed a notable reduction in exocrine tissue in “homozygous” mutants, particularly pronounced in Pdx1 ^s/s mutants, alongside an increased proportion of ductal structures (Figure 4F). Further immunofluorescence analysis using antibodies against INS, GCG and SST demonstrated a decrease in endocrine cells in Pdx1 ^m/m mutants and a near-total depletion in Pdx1 ^s/s mutants (Figures 4G–I). Intriguingly, the presence of INS⁺/GCG⁺ dihormonal cells and a small number of GCG⁺/SST⁺ dihormonal cells, which were nearly absent in 8-month-old d/d individuals, was noted in Pdx1 ^m/m mutants (Figure 4J; Supplementary Figure S6). These findings underscore the critical role of Pdx1 in pancreas development and blood glucose regulation in axolotls, consistent with observations in mammals (Offield et al., 1996; Oliver-Krasinski et al., 2009). In summary, Pdx1 mutant axolotls exhibit pancreatic developmental defects and a reduction in both endocrine and exocrine cell populations, characteristics similar to those observed in mammals. However, unlike mammals (Jonsson et al., 1994), these mutant axolotls can survive for long period, despite exhibiting severe pancreatic defects.

GTT and ITT approach are standard methods for assessing glucose metabolism and insulin response. We employed intraperitoneal GTT and ITT to characterize these processes in axolotls. Assessing these processes in smaller models like zebrafish is challenging, so it is particularly relevant to determine their feasibility in axolotls. We initially followed established procedures used in mice (Cassano et al., 2020; Qu et al., 2019; Zhu et al., 2021b), but noted unexpected fluctuations in glucose and insulin levels during the measurement periods. Considering the evidence of the slower glucose metabolism in axolotls suggested by sequencing data, we optimized our experimental protocols for intraperitoneal GTT and ITT. Blood glucose levels were measured at various time points: 0, 1, 2, 6, 12, 24, 36 and 48 h following a 2 mg/g glucose injection during GTT (Figure 5A). In comparison to blood glucose measurement sampling in mice (Small et al., 2022), the intervals for measurements in axolotls are longer. Our findings revealed that the average fasting blood glucose level in adult axolotls was 3.317 ± 0.419 mmol/L 12 h after an intraperitoneal injection of glucose (2 mg/g), blood glucose levels rose to 7.55 ± 0.369 mmol/L. By 36 h post-injection, glucose levels returned to basal level (2.867 ± 0.486 mmol/L). ITT was performed on 6 h fasted axolotls. 5 μL/g insulin glargine was injected into the axolotl, and blood glucose levels were measured at 0, 0.5, 1, 2, 4, 6 and 12 h after insulin injection (Figure 5B). The results showed that blood glucose levels decreased from 3.962 ± 0.134 mmol/L at baseline to 1.525 ± 0.207 mmol/L at the 12-h mark, where levels stabilized. In summary, axolotls exhibit lower blood glucose homeostasis compared to mammals, aligning with the notion that blood glucose levels correlate positively with metabolic rates (Simon et al., 2019; DeFronzo et al., 1979). This aligns with gene expression analysis and further indicates slower glucose metabolism in axolotls.