Gluconeogenesis in frogs during cooling and dehydration exposure: new insights into tissue plasticity of the gluconeogenic pathway dependent on abiotic factors

It is undeniable that anurans undergo physiological changes and metabolic variation influenced by different life stages, nutritional status, geographic gradients, seasons and abiotic environmental stressors, such as temperature and humidity (Pough and Kamel, 1984; Schmuck et al., 1994; Navas, 1996; Secor, 2005; Navas, 2006; Navas and Otan, 2007; Costanzo et al., 2015; Storey and Storey, 2017; Timpone et al., 2020; Moreira et al., 2021; de Amaral et al., 2022). Their complex life cycle and morphophysiological characteristics require access to water, prioritizing habitats with minimal temperature and humidity fluctuations (Duellman and Trueb, 1986; Pough et al., 2004). However, certain anuran species are able to inhabit environments deviating from these conditions, such as low humidity or cold temperatures, through physiological adaptations.

During freezing conditions or periods of low humidity, these anurans developed, for example, a hyperglycemic response (Storey et al., 1984; Churchill and Storey, 1994; Costanzo and Lee, 2013; Storey and Storey, 2013). This same response to both stressors corroborates the idea that these physiological adaptive mechanisms in anurans are derived and optimized from pre-existing resources (Storey and Storey, 2017). With this in mind, it is imperative to identify and elucidate glucose metabolism during these conditions to gain a deeper understanding of anuran metabolic strategies. Furthermore, anurans are one of the groups most threatened by climate change (Luedtke et al., 2023). Thus, understanding the adaptation and physiological mechanisms they employ to face extreme temperatures or dry conditions is a tool to assist in their conservation efforts (Madliger et al., 2018).

During prolonged fasting, hibernation or vigorous exercise, glycogen stores become depleted, necessitating the de novo synthesis of glucose to maintain blood glucose levels (Gleeson, 1985; Costanzo et al., 2015; Nakajima et al., 2023). Consequently, the participation of the gluconeogenic pathway becomes indispensable for anuran survival. Furthermore, gluconeogenesis encompasses multiple enzyme-dependent stages regulated by hormones, nutrient intake, substrate concentrations and stress conditions (Shah and Wondisford, 2020). The literature consistently indicates gluconeogenesis activity through enzyme levels or substrate correlations in anurans (see Nagano et al., 1975; Cowan and Storey, 2001; Vogiatzis and Loumbourdis, 2001; Kiss et al., 2011; Fan et al., 2022), but the conventional approach for assessing gluconeogenesis involves tracing labeled carbon in glucose (¹⁴C or ¹³C) originating from labeled substrates such as lactate, alanine or glycerol (Radziuk and Pye, 2001). Despite the pivotal role of gluconeogenesis, there remains a scarcity of studies offering insights into this pathway within the Anura group.

Additionally, studying glucose pathways could provide new discoveries in the area. Fournier and Guderley (1992), for example, indicate the possibility of gluconeogenesis from lactate in muscle because, after strenuous exercise, most of the lactate accumulated by frogs appears to be recycled into muscle glycogen. Therefore, these results open an interesting field for study, as gluconeogenesis in muscle tissue is not uncommon, as our group has already described in the crab Chasmagnathus granulata using isotope dilution techniques (Schein et al., 2004, 2005). By employing isotope dilution techniques, the ratio of radiolabeled glucose to the labeled precursor reflects the percentage contribution of the precursor to glucose production (Shah and Wondisford, 2020). To our knowledge, only one study has used this technique in anuran species to demonstrate the conversion of lactate to glucose in frog muscle tissue (Petersen and Gleeson, 2011). Consequently, our study aimed to provide novel insights into this pathway in frog tissues using the subtropical tree frog Boana pulchella as a model.

Boana pulchella is found in southern Brazil, Uruguay, Paraguay and Argentina (Kwet et al., 2004), inhabits various natural habitats including forests, dry lowland grasslands, savannas and areas where agriculture is encroaching, transforming the habitat and drying up puddles (Brodeur et al., 2012; Larramendy and Soloneski, 2017). Despite being a subtropical species, it can endure cold temperatures and remains active during the winter season (Both et al., 2008; Canavero et al., 2008; Maneyro, 2008; Ximenez and Tozetti, 2015), displaying seasonal metabolic changes (de Amaral et al., 2022). Additionally, this species has shown metabolic adaptations during cooling exposure (de Amaral et al., 2023), demonstrating an adaptive capacity to environmental stressors. These traits make B. pulchella an intriguing model for studying physiological adjustments under abiotic stress. Thus, we assessed gluconeogenesis using ¹⁴C-lactate, ¹⁴C-alanine, ¹⁴C-glycerol and ¹⁴C-glutamine in the liver, muscle and kidney tissues of B. pulchella frogs subjected to cold and dehydration conditions. Our investigation aimed to elucidate the gluconeogenic capacity and substrate preferences within this animal group exposed to abiotic stress factors, thereby providing evidence regarding the gluconeogenic pathway in anurans and contributing data to understand metabolic adjustments in habitats subject to environmental variations.

Fifteen male Boana pulchella (Duméril & Bibron 1841) (Anura/Hylidae) frogs used in this study were captured during the autumn in Eldorado do Sul, Brazil (30°06′02.9″S, 51°40′35.0″W). In this site, the recorded temperature ranged from 8 to 21°C (Bergamaschi et al., 2013) and after the captures it ranged from 5 to 15°C (measured using a single handheld Kestrel 3500 Weather Meter^®). Wild-caught frogs (mass ∼4 g) were transported to the Federal University of Rio Grande do Sul laboratory after capture and washed in a tetracycline bath (0.5%) to prevent infections during the exposure. Subsequently, we followed the exposure protocol described by de Amaral et al. (2023). Briefly, the frogs were acclimated for 2 weeks at 5°C under a 10 h:14 h light:dark regimen in a temperature-controlled incubator (FOC 225E incubator, VELP Scientifica^®, Usmate Velate, Italy; TC-900e POWER thermostat, Full Gauge Controls^®, Niterói, Brazil). The frogs were housed in ventilated plastic containers with damp sphagnum moss and paper towels during acclimation. They had free access to dechlorinated water, which was also sprayed on them daily, and were fed twice weekly with mealworm larvae and small beetles during the acclimation period. A 48 h fast was conducted before the experimental exposure to low temperatures.

The control group of animals (n=5) was randomly sampled from this condition after 2 weeks. The remaining animals were exposed to gradual cold exposure, with a temperature decrease of 1°C per day until it reached −2.5°C. Afterwards, the incubator was set to −4°C for a 45 min cooling period, allowing frog body temperature to reach subzero values and promote ice nucleation. The incubator temperature was then raised to −2.5°C for 24 h. After 24 h, the cold-exposed group was randomly sampled (n=5). The remaining animals (n=5) were subjected to 24 h at 5°C and were subsequently sampled, forming the recovery group.

All animals used in the dehydration protocol (n=21) were collected at the same Eldorado do Sul site during winter. The exposure protocol of Churchill and Storey (1994), Wu et al. (2018) and Storey and Storey (2019) was followed. First, five frogs were used to obtain the frogs' initial body water content (BWC_i), necessary to determine the dehydration level during the experiment. As soon as they arrived at the laboratory, the five animals were weighed and euthanized with 5% lidocaine (topical anesthetic cream) applied in the inguinal region and oral cavity. They were then placed in an oven at 80°C and weighed every 12 h. When the mass did not change for 24 h, the body mass was considered entirely dehydrated, and the BWC_i was determined by subtracting the final mass from the initial mass. The BWC_i of the B. pulchella frog was estimated to be 0.797±0.005 g of water per gram of body mass.

The remaining animals (n=16) were transported to the laboratory, where they were washed in a tetracycline bath (0.5%) and acclimated under the same conditions as those in the cooling protocol, differing only in the housing arrangement. The frogs were housed in glass vacuum desiccators equipped with porcelain disks but without vacuum pressure to facilitate air exchange. Damp sphagnum moss and paper towels were placed inside the desiccators. During the acclimation period, mealworm larvae and small beetles were offered to the animals twice a week, and they were not fed 48 h before the experimental exposure. The control group of animals (n=5) was randomly selected from this condition after a 2 week acclimation period.

After acclimation, the animals underwent a dehydration procedure. Initially, all animals with identified mass were recognized using the method of photo identification of the inner parts of the hind legs (in this species, this region exhibits characteristic spots used for individual identification). In the desiccators, a layer of silica gel desiccant was positioned on the bottom of the porcelain disk to prevent the animals from contacting this material. Furthermore, the interior of the desiccator was meticulously dried, and the sphagnum moss and paper towels were replaced with dried sphagnum and paper towels. Following the weighing and identification process, we emptied the urinary bladder of the toads by gently pressing the pelvic region, and the animals were returned to the desiccators at 5°C under a 10 h:14 h light:dark cycle. Every 12 h, each frog was briefly weighed to determine the progress of water loss. Frogs were dehydrated until they reached approximately 40% total body water loss. The percentage of total body water lost was calculated using the following equation: body water loss (%)=[(M_i−M_d)/(M_i×BWC_i)]×100, where M_i represents the initial mass of the animal, M_d is the mass of the dehydrated frog at each weighing and BWC_i represents the initial body water content of the frog, as above. Once 40% body water loss was reached, the animal was immediately euthanized as above, contributing to the dehydration group (n=6), or rehydrated (n=5). For rehydration, frogs were placed in a container with distilled water (approximately 0.5 cm deep) at 5°C for 24 h before being euthanized.

All animals utilized in this study, in both cold and dehydration states, had their body mass (M) measured (in g) and snout–vent length (SVL) measured (in cm). Subsequently, they were euthanized one by one through the application of 5% lidocaine in the abdomen and the oral cavity. After confirming euthanasia, the specimens were immediately dissected over ice to remove the liver, leg muscles (gastrocnemius, gracilis, sartorius and adductor) and kidney. These tissues were promptly employed in gluconeogenesis analyses.

The gluconeogenesis assay was performed according to Oliveira and Da Silva (1997) and Oliveira et al. (2004). Liver (∼10 mg), muscle (∼80 mg) and kidney (∼10 mg) fresh tissue slices were incubated in microtubes containing 0.5 ml of Krebs–Ringer Buffer [114 mmol l⁻¹ NaCl, 2.25 mmol l⁻¹ KCl, 0.44 mmol l⁻¹ KH₂PO₄, 0.33 mmol l⁻¹ Na₂HPO₄, 1 mmol l⁻¹ MgSO₄,13 mmol l⁻¹ NaHCO₃, 10 mmol l⁻¹ Hepes, 0.1% albumin, 10 µl ml⁻¹ PMSF (Sigma-Aldrich^® #P7626) and 1 µl ml⁻¹ Protease Inhibitor Cocktail (Sigma-Aldrich^® #P8340); pH 7.63] and labeled and unlabeled substrates. Liver slices were incubated in 0.10 μCi of l-[U-¹⁴C]-alanine (¹⁴C-alanine) (151 mCi mmol⁻¹; Perkin Elmer) plus 5 mmol l⁻¹ unlabeled l-alanine, or 0.10 μCi of l-[U-¹⁴C]-lactate (¹⁴C-lactate) (250 mCi mmol⁻¹; Du Pont) plus 5 mmol l⁻¹ unlabeled l-lactate, or 0.10 μCi of l-[U-¹⁴C]-glycerol (¹⁴C-glycerol) (154 mCi mmol⁻¹; Perkin Elmer) plus 5 mmol l⁻¹ unlabeled l-glycerol. Muscle slices were incubated in 0.10 μCi of l-[U-¹⁴C]-alanine (¹⁴C-alanine) (151 mCi mmol⁻¹; Perkin Elmer) plus 5 mmol l⁻¹ unlabeled l-alanine and 0.10 μCi of l-[U-¹⁴C]-lactate (¹⁴C-lactate) (154 mCi mmol⁻¹; Perkin Elmer) plus 5 mmol l⁻¹ unlabeled l-lactate. Kidney slices were incubated in 0.10 μCi of l-[U-¹⁴C]-glutamine (¹⁴C-glutamine) (266 mCi mmol⁻¹; Amersham) plus 5 mmol l⁻¹ unlabeled l-glutamine. After adding the unlabeled substrates, the pH of the incubation medium was determined. The gaseous microtube phase was saturated with a 5% CO₂ and 95% O₂ mixture for 20 s. The slices were incubated at 25°C for 60 min in a Dubnoff metabolic shaker (60 cycles min⁻¹). Previous studies have demonstrated that, under experimental conditions, 5% CO₂ does not affect gluconeogenic capacity (Oliveira et al., 2004).

At the end of the incubation, the medium was deproteinized, and ¹⁴C-glucose, formed from l-[U-¹⁴C]-alanine and l-[U-¹⁴C]-lactate, was separated by thin-layer chromatography using 95% n-butanol, 5.4% ethyl alcohol and acetic acid in water (75:47.4:27.6 v/v/v). The spot corresponding to ¹⁴C-glucose, localized by spraying with an anisaldehyde reagent (95% ethyl alcohol/concentrated sulfuric acid/p-methoxybenzaldehyde 18:1:1 v/v/v), was marked, scraped off and dissolved in scintillation liquid (SLC): toluene:Triton X-100 (2:1, v/v), PPO 0.4%, POPOP 0.01%. The radioactivity was measured using an LKB counter (LKB-Wallac). The values of gluconeogenic activity are given as mmol of l-[U-¹⁴C]-lactate, l-[U-¹⁴C]-alanine, l-[U-¹⁴C]-glycerol or l-[U-¹⁴C]-glutamine converted to ¹⁴C-glucose g⁻¹ tissue h⁻¹.

All protocols conducted in this study were approved by the Ethics Committee of the Federal University of Rio Grande do Sul (CEUA/UFRGS) under number #39416. Specimen collection was authorized by the Chico Mendes Institute for Biodiversity Conservation (ICMBio/SISBIO) permit #75475-3.

The statistical analyses were conducted using GraphPad Prism version 8.0.2 (GraphPad Software, San Diego, CA, USA). Normality and homogeneity were assessed for all data using the Kolmogorov–Smirnov test. Outliers were identified and excluded following the ROUT method, with Q set at 1% (Motulsky and Brown, 2006). To assess differences among the three treatments (control, cooling or dehydration, and recovery groups) in the two exposures (cooling and dehydration), we conducted a one-way ANOVA test followed by the Tukey post hoc test when the data exhibited a normal distribution (liver gluconeogenesis in cooling and dehydration exposure from all ¹⁴C-substrates; muscle gluconeogenesis synthesis from ¹⁴C-alanine in cooling and from ¹⁴C-lactate in dehydration; kidney gluconeogenesis synthesis from ¹⁴C-glutamine in cooling), and the results were expressed as the mean±s.e.m. For non-parametric data (muscle gluconeogenesis synthesis from ¹⁴C-lactate in cooling and from ¹⁴C-alanine in dehydration; kidney gluconeogenesis synthesis from ¹⁴C-glutamine in dehydration), we applied the Kruskal‒Wallis test, followed by Dunn's post hoc test, and the results are presented as the median and interquartile interval 25/75. P≤0.05 was considered for determining statistical significance.

During cold exposure, there was a general decrease in liver gluconeogenesis activity in both the cooling and recovery groups (Fig. 1). Specifically, the gluconeogenesis activity from ¹⁴C-glycerol showed significant differences between the control and cooling/recovery groups (P=0.0090 and P=0.0059, respectively; Fig. 1B). There was a 47% decrease in the cooling group and a 52% decrease in the recovery group compared with the control group. Similarly, gluconeogenesis activity from ¹⁴C-lactate significantly differed between the control and cooling/recovery groups (P=0.003 and P=0.005, respectively; Fig. 1C). During cooling and recovery, gluconeogenesis activity from ¹⁴C-lactate decreased by 47% and 45%, respectively, compared with the control animals. In contrast, gluconeogenesis activity from ¹⁴C-alanine did not vary significantly between the groups (P=0.0757; Fig. 1A). However, lower means were observed in the cooling group (2059±206.6 mmol g⁻¹ min⁻¹) than in the control group (3502±494.7 mmol g⁻¹ min⁻¹). In the liver, gluconeogenesis activity from ¹⁴C-lactate and ¹⁴C-alanine presented higher values than gluconeogenesis activity from ¹⁴C-glycerol (P<0.0001).

In contrast, during dehydration exposure, liver gluconeogenesis activity from ¹⁴C-alanine and ¹⁴C-glycerol was not significantly different between groups (P=0.3004 and P=0.2146, respectively; Fig. 2A,B). However, a significant difference in gluconeogenesis activity from ¹⁴C-lactate was observed between the control and recovery groups and the dehydration and recovery groups (P=0.0069; Fig. 2C). The recovery group exhibited an 85% decrease in gluconeogenesis activity compared with the control and dehydration groups.

Gluconeogenesis activity from ¹⁴C-alanine and ¹⁴C-lactate in frog muscle was evaluated for the first time in the literature in Hylidae frogs. During cooling exposure, we found no change in gluconeogenesis activity between the control, cooling and recovery groups (Fig. 3A,B). During dehydration exposure, muscle gluconeogenesis activity from ¹⁴C-alanine significantly differed between the dehydration and recovery groups (P=0.0065; Fig. 3C). The recovery group exhibited higher activity levels than the dehydrated animals. However, gluconeogenesis activity from ¹⁴C-lactate did not differ significantly between the groups (P=0.0808; Fig. 3D), although it is worth noting that the dehydration group displayed high activity of lactate conversion to glucose. Furthermore, alanine is a preferred substrate for cooling over lactate (Fig. 3A,B). Conversely, the opposite is the case during dehydration, with lactate becoming the preferred substrate over alanine (Fig. 3C,D).

The kidney gluconeogenesis activity from ¹⁴C-glutamine significantly differed between the control and cooling groups (P=0.0190; Fig. 3E). There was an 83% decrease in gluconeogenesis activity in the cooling group compared with the control group. The gluconeogenesis activity from ¹⁴C-glutamine in the kidney did not show differences between the groups during dehydration exposure (Fig. 3F).

Anurans exhibit various metabolic adaptations, enabling them to inhabit diverse environments (Weber et al., 2002; Navas et al., 2004; Costanzo and Lee, 2013; Larson et al., 2014; Muir et al., 2014; Niu et al., 2021). The study of the intermediary metabolism of anurans provides valuable information about which substrates are most important. The choice of a preferential energy substrate depends, for example, on factors such as season (King et al., 1995; Kiss et al., 2009; de Amaral et al., 2022) and environmental conditions, including temperature and humidity (de Amaral et al., 2023; Hawkins et al., 2019; Park and Do, 2020; Yoldas and Erismis, 2021). The gluconeogenic pathway appears crucial in these adaptations; however, few studies have elucidated the activity of this pathway in frog tissues. This study represents the first measurement of gluconeogenic activity using radiolabeled substrates in adult frog liver, muscles and kidneys during exposure to cold and dehydration. Gluconeogenic activity was detected in the liver, kidney and muscle tissues of the subtropical frog B. pulchella, where this pathway had not previously been demonstrated. The findings of this study suggest that environmental factors influence gluconeogenic pathways, and these pathways differ between tissues, indicating a tissue-specific profile. Additionally, this tissue plasticity for the gluconeogenic pathway may signify metabolic adaptations to various stressors and the utilization of preferential substrates depending on the environmental conditions in anurans.

In vertebrates, gluconeogenic activity occurs in the liver and kidney, where glucose is produced from lactate, amino acids or glycerol (Moon, 1988; Berg et al., 2011). Particularly in fasted states, the liver is a major metabolic organ that helps maintain plasma glucose levels through gluconeogenesis (Veyrenc et al., 2022). However, during cold exposure and up to 24 h afterwards, the gluconeogenesis pathway exhibits decreased activity in the B. pulchella liver. The literature has already suggested a decrease in gluconeogenesis during cold exposure in other anurans. In Rana sylvatica, fructose-1,6-bisphosphatase (FBPase), a crucial enzyme regulating gluconeogenesis, significantly declined, suggesting its suppression and the inhibition of liver gluconeogenesis during freezing (Varma and Storey, 2022). Seasonally, in Rana catesbeiana, cytosolic phosphoenolpyruvate carboxykinase (PEP carboxykinase), a key enzyme in gluconeogenesis, exhibits maximal activity during the summer for adult frogs, compared with the coldest months (Rexer-Huber et al., 2011). These results may indicate that during cold states, the liver experiences gluconeogenesis suppression from glycerol and lactate substrates, while alanine becomes the preferential substrate used during these situations. Moreover, at least 24 h after exposure to cold, gluconeogenesis activity from lactate and glycerol continues to be reduced, suggesting that it takes time to re-establish the activity of the pathway.

During cold exposure, glycerol may serve other purposes than glucose production. It is well known that glycerol acts as a cryoprotectant during freezing or increases in concentration during cold acclimation (Rexer-Huber et al., 2011; Zimmerman et al., 2007). As previously demonstrated, B. pulchella exhibits higher glycerol concentrations during the winter season or when exposed to cold conditions compared with other freeze-tolerant frogs (de Amaral et al., 2022, 2023). Additionally, the expression of genes related to fatty acid oxidation was downregulated in the fish Oreochromis niloticus when exposed to 10°C, suggesting reduced lipolysis and increased lipid storage to provide energy under cold stress (Huang et al., 2022). These results may indicate that B. pulchella maintains higher glycerol storage during cold conditions to cope with freezing temperatures and preserve lipid stores. This is particularly important for this species, as it inhabits areas with harsh winters and yet concentrates its activity during these colder months (Both et al., 2008; Maneyro, 2008; Antoniazzi et al., 2019). As a result, glucose may not be primarily used as a cryoprotective agent. Thus gluconeogenic flux from glycerol during cold exposure may decrease, so retaining this substrate for potential cold challenges.

However, lactate typically accumulates in the liver of frozen frogs, indicating that anaerobic glycolysis occurs during freezing or overwintering (Storey et al., 1984; Storey and Storey, 1984, 1986; Niu et al., 2023). Therefore, during cooling and recovery, the reduced gluconeogenesis from lactate in the liver may suggest that lactate is utilized through anaerobic glycolysis to provide energy to this tissue. Similarly, during dehydration exposure, the decreased gluconeogenesis from lactate in the liver during recovery could be associated with a strategy to provide energy for re-establishing metabolism. In the recovery phase from dehydration, the activity of lactate dehydrogenase (LDH), the enzyme responsible for the last step in anaerobic glycolysis that converts pyruvate to lactate, was less active in the dehydrated liver of Xenopus laevis (Childers and Storey, 2019). This suggests that a significant portion of pyruvate formed via glycolysis may be directed toward the tricarboxylic acid (TCA) cycle, favoring a return to oxidative phosphorylation (Childers and Storey, 2019) and perhaps less lactate is being directed towards the gluconeogenesis pathway. Furthermore, while the liver is a primary site for lactate recycling from the muscle, lactate may no longer be able to reach the liver during exposure to cold and recovery. Lactate may be recycled within the muscle, as previously reported in post-exercise situations (Fournier and Guderley, 1992; Gleeson and Dalessio, 1990). This reduced transfer of lactate from the liver to muscle could decrease the activity of gluconeogenesis from this precursor in the liver. Recycling lactate within the muscle might be a strategic adaptation to conserve energy, avoiding the expenditure associated with the Cori cycle or potential loss of lactate during liver–muscle transfer. This approach enables rapid energy generation directly within the muscle during stress exposure, which is particularly crucial for B. pulchella. This frog remains active during winter (Maneyro and Carreira, 2012; de Amaral et al., 2022), and optimizing energy resources is essential to sustain activities such as vocalization. Additionally, this animal inhabits areas of extensive agriculture, where puddles may disappear as a result of planting activities (Brodeur et al., 2012). Therefore, adaptation to dry periods and optimization of energy resources during these periods may ensure the survival of individuals.

The stability of gluconeogenic activity via alanine in the liver during exposure may be attributed to the dietary habits of these carnivorous animals, which provide a rich protein source, thereby promoting a pool of this type of substrate (Schermerhorn, 2013). Studies involving crabs fed high-protein diets have shown that hemolymph glucose levels are primarily maintained through effective gluconeogenesis from ¹⁴C-alanine (Dias dos Santos et al., 2021). Furthermore, alanine could originate from muscle tissue and reach the liver, contributing to hepatic gluconeogenesis (Solé and Pelz, 2007). A reduction of total muscle proteins in B. pulchella subjected to cooling has previously been shown (de Amaral et al., 2023). Considering the substantial protein content in the diet (Withers, 1978; Da Rosa et al., 2011), it is conceivable that the reduction in gluconeogenesis stemming from alanine may exhibit a time-delayed response during cold exposure. Consequently, prolonged cold exposure could potentially intensify the decrease in this activity. In summary, the liver of B. pulchella demonstrates varying gluconeogenic activity during episodes of cold and dehydration exposure, thereby revealing distinct substrate preferences in response to differing environmental conditions. Furthermore, it exhibits an augmented gluconeogenic activity from ¹⁴C-alanine and ¹⁴C-lactate compared with ¹⁴C-glycerol.

As mentioned earlier, the kidney also exhibits a high level of gluconeogenic activity. In frogs, cold temperatures appear to have a suppressive effect on kidney gluconeogenesis. This phenomenon has been observed in Lithobates sylvaticus, where freezing reduces the activity of the regulatory enzyme of gluconeogenesis, FBPase, in the kidney, and subsequently, during the thawing phase, the activity is re-established (Cowan and Storey, 2001). Similarly, in B. pulchella, gluconeogenesis using ¹⁴C-glutamine as a substrate decreased during cold exposure. It is plausible that exposure to cold promotes the suppression of kidney gluconeogenesis, which may contribute to stabilizing the cryoprotectant pool by reducing the potential for glucose phosphorylation and catabolism. During cooling in ectotherms, it is known that P_CO2 decreases, leading to an increase in pH (Sastrasinh and Sastrasinh, 1990). In the kidney, lowering extracellular pH stimulates the inward transport of glutamine (Nissim, 1999) and alkaline intracellular pH, reducing the transport of glutamine into the mitochondrial matrix and consequently reducing renal ammonia (NH₃) production (Goldman and Witkovsky, 1987). Thus, lower gluconeogenic activity from renal glutamine might be due to lower glutamine levels in renal cells in response to a more alkaline environment induced by lower temperatures. Additionally, lower glutamine concentrations could prevent urea accumulation, given the previous understanding that cooling reduces the concentration of this osmolyte in the tissues of B. pulchella (de Amaral et al., 2023). In contrast, it is noteworthy that stress does not appear to decrease gluconeogenic pathway activity during dehydration. Under these conditions, glucose production may offer advantages in overcoming osmotic stress. This observation underscores the distinct responses to different stressors and how they may modulate the gluconeogenic pathway in various ways.

In anurans, gluconeogenesis is not limited to the liver and kidney; studies have also revealed its presence in other tissues. For instance, in R. catesbeiana, the existence of a gluconeogenic pathway in the retina has been demonstrated (Goldman, 1988). Additionally, Bufo marinus toads exhibit gluconeogenic activity in the urinary bladder (Morrison et al., 1972) and gastric mucosa (Finol and Chacin, 1980). Fournier and Guderley (1992) suggested that Rana pipiens could directly recycle lactate to glycogen in muscle after vigorous activity, and in R. catesbeiana, muscular gluconeogenesis from lactate is implied (Petersen and Gleeson, 2011). Our study extends this understanding by demonstrating that the gluconeogenesis pathway occurs not only from lactate but also from alanine in the muscle of B. pulchella. This finding aligns with previous research on the role of this pathway in frog muscle energy dynamics. While R. catesbeiana has shown measurable gluconeogenic capacity from lactate in muscle, this capacity did not vary between cold and warm conditions. Furthermore, in cold temperatures, incubation with insulin did not impact muscle gluconeogenesis activity (Petersen and Gleeson, 2011). In our previous study, B. pulchella exhibited higher lactate concentrations in muscle, while concentrations in other tissues remained constant (de Amaral et al., 2022). Notably, these concentrations did not change during cooling exposure. Considering these findings, we can hypothesize that muscle lactate is an essential substrate recycled within the muscle tissue during stress exposure, possibly through gluconeogenesis. This lactate may be directed toward energy production as glucose through oxidative phosphorylation or stored as glycogen (Fig. 4), as suggested in reptiles (Gleeson, 1985; Gleeson and Dalessio, 1990).

This hypothesis challenges the conventional understanding of gluconeogenesis from lactate, which typically involves the liver in mammals, where lactate is exported and gluconeogenesis occurs in the liver via the Cori cycle (Cori and Cori, 1946; Cori, 1981). In contrast, our findings suggest that in anurans, particularly in B. pulchella, gluconeogenesis from lactate may occur within the muscle tissue. Furthermore, the observation that B. pulchella remains active during cold exposure (de Amaral et al., 2023) and throughout the winter in its natural habitat (Maneyro and Carreira, 2012; Ximenez and Tozetti, 2015) supports the notion that cold temperatures do not significantly impact gluconeogenesis in these frogs. This resilience to cold conditions may represent a metabolic adaptation that evolved in frogs that inhabit colder environments.

Furthermore, it is worth noting that B. pulchella primarily inhabits wetlands in southern Brazil, and these environments typically do not undergo natural dry spells (Ximenez and Tozetti, 2015; Santos et al., 2016). However, as agricultural practices advance in the habitats of this species (Brodeur et al., 2012), the puddles are drying up, necessitating their movement to find new moist areas to expand their suitable habitat (Alves-Ferreira et al., 2022), exposing them to dehydration periods. Consequently, the observed decrease in gluconeogenesis activity from ¹⁴C-alanine in muscle during experimental dehydration, compared with the recovery phase, may suggest that when fully hydrated after a period of water restriction, gluconeogenesis from protein sources could play a crucial role in supplying glucose to re-establish muscle activity. Studies with C. granulata crabs showed that 24 h of hyperosmotic shock increased ¹⁴C-alanine incorporation into glucose, PEPCK gene expression and mitochondrial enzyme activity in the muscle of winter-collected animals (Schein et al., 2005), and the concentration of amino acids during hyperosmotic states in crustacean tissues increases (Gilles, 1997). Dehydration could lead to protein catabolism in the muscle, providing a higher alanine concentration as a substrate for conversion to glucose when the animal returns to a hydrated state. Thus, our study highlights that the activity of this pathway in muscle can vary during different stress exposures, remaining relatively constant during cold stress but exhibiting variability during dehydration exposure.

In conclusion, it is clear that the metabolic changes in anuran frogs during stress exposure are intricate, with considerable intra- and inter-specific variability. Nevertheless, our study has yielded valuable insights into how frog tissue gluconeogenesis responds to cold and dehydration stress, likely through finely tuned adjustments in enzyme activities depending on substrate availability. This adaptation appears to facilitate organ-specific responses in B. pulchella to cope with stress exposure during seasons, such as lower temperatures, or caused by anthropogenic factors such as the drying of ponds as a result of agricultural practices. Additionally, we cannot forget that with climate change, the environments of anuran populations may be altered (Luedtke et al., 2023). Drying of ponds and temperature changes are present factors that can interfere with species dynamics (Lertzman-Lepofsky et al., 2020); therefore, having physiological mechanisms to deal with these challenges may be crucial for the survival of the species. Also, these findings pave the way for exploring the dynamics of gluconeogenesis metabolism in ectothermic species, figuring out which substrates are important, especially in unconventional tissues such as muscle. Moreover, further research into gluconeogenic pathways in frogs, including the evaluation of key enzyme activities and their expression under diverse conditions, could contribute to a more profound understanding of the role of this pathway within the amphibian group.

We thank the staff of the Agronomic Experimental Station of UFRGS (EEA-UFRGS) for their support during the animal collections, the Graduate Program in Biological Sciences: Physiology for providing space for conducting the experiments, and the funding agencies for financial support.