Integrated Metabolomic and Bioenergetic Profiling Reveals Axl as a Key Regulator of Macrophage Homeostasis

2.1. Axl Deficiency Alters the Composition and Phenotype of Peritoneal Macrophages Under Homeostatic Conditions

To investigate PMs under homeostatic conditions, peritoneal fluid (PF) cells were analyzed by flow cytometry, and CD19⁺ B‐1 cells were excluded from the gating strategy (Figure 1A). PMs were identified based on CD11b and F4/80 expression [25], and full‐body (global) Axl^−/− mice exhibited a significant reduction in PMs compared with WT controls (Figure 1B,C). Next, based on MHCII and CD206 expression, PM subpopulations were analyzed, revealing a significant decrease in both the transitional CD206⁺MHCII⁺ PMs and the MHCII⁺ PM subset in Axl^−/− mice relative to WT animals (Figure 1D,F,G), which shows that the PM pool is partially dependent on Axl signaling that may be either cell‐intrinsic within PMs or indirect, mediated through other Axl‐expressing cell types within the peritoneal environment that can influence PM composition and phenotype. Given that MHCII⁺ macrophages are commonly associated with a more inflammatory, proinflammatory‐like profile, whereas CD206⁺ cells display features of alternatively activated, anti‐inflammatory‐like macrophages, these data suggest that Axl contributes to the homeostatic balance among phenotypically distinct PM subsets [29]. Further phenotypic characterization demonstrated a significant reduction in the mean fluorescence intensity (MFI) of the resident macrophage‐associated markers F4/80 and CD11b in PMs from Axl^−/− mice compared with WT controls (Figure 1H,I), indicating impaired macrophage maturation. In addition, PMs from Axl‐deficient mice showed a trend toward increased Ly6C expression (p = 0.0687) (Figure 1J), suggestive of an enrichment of less mature, monocyte‐like cells within the PF. Consistently, MHCII expression was also significantly reduced in PMs From Axl^−/− mice (Figure 1L), pointing to a diminished acquisition of an antigen‐presenting phenotype, whereas CD206 levels remained unaffected (Figure 1K). Collectively, these data demonstrate that global Axl deficiency disrupts the phenotypic composition of the peritoneal macrophage compartment.

2.2. Axl Deficiency Alters the Metabolic and Oxidative Profile of Peritoneal Macrophages

The phenotypic alterations observed in PMs suggest a shift toward a less mature and functionally reprogrammed PM (Figure 1). Given that macrophage maturation and functional specialization are tightly linked to distinct metabolic programs, proinflammatory‐like cells are predominantly glycolytic, and anti‐inflammatory‐like cells depend on oxidative phosphorylation and the tricarboxylic acid (TCA) cycle [30]. We next examined whether these phenotypic changes were associated with alterations in metabolic and effector functions. PMs from Axl^−/− mice exhibited significantly reduced nitrite levels after 24 and 48 h under unstimulated conditions (Figure 2A), indicating impaired basal nitric oxide production. In contrast, lactate accumulation in the culture supernatants of Axl^−/− PMs was significantly increased at both time points (Figure 2B), suggesting a shift toward enhanced glycolytic metabolism. This phenotype was further amplified following LPS stimulation, with Axl^−/− PMs producing markedly higher levels of lactate compared with WT controls (Figure 2C). Importantly, lactate dehydrogenase (LDH) measurements confirmed that the increased lactate levels were not due to cell death, indicating that these metabolic changes were intrinsic to macrophage nitrite production (Figure 2D).

Assessment of ROS production revealed no significant differences between WT and Axl^−/− PMs under basal conditions (Figure 2E). However, upon LPS stimulation, WT PMs exhibited a robust increase in ROS production over a 60 min period, whereas Axl^−/− PMs generated significantly lower levels of ROS (Figure 2F). Analysis of mitochondrial ROS using MitoSOX Red revealed that Axl^−/− PMs exhibited an increased frequency of MitoSOX‐positive cells (Figure 2G,H), consistent with elevated mitochondrial ROS levels in Axl‐deficient peritoneal macrophages. Collectively, these data demonstrate that Axl is essential for maintaining the metabolic and functional profile of peritoneal macrophages, linking alterations in macrophage phenotype to impaired effector responses.

2.3. Selective Disruption of Glycolytic Intermediates Reveals Impaired Metabolic Adaptability in Axl^−/− Macrophages

To further characterize the metabolic reprogramming associated with global Axl deficiency in PMs, we performed an untargeted metabolomic analysis. Unsupervised hierarchical clustering of the metabolomic data revealed a clear separation between WT and Axl^−/− PMs, indicating a global remodeling of the metabolic profile in the absence of Axl signaling (Figure 3A). Consistently, principal component analysis (PCA) demonstrated distinct clustering of WT and Axl‐deficient samples along the principal components, confirming that Axl^−/− PMs occupy a metabolically distinct state (Figure 3B). Metabolites contributing most strongly to group separation were predominantly associated with glycolysis, energy metabolism, and redox balance, in line with the metabolic and functional alterations observed in previous analyses. Based on these global differences, we next examined the individual variation of key metabolites to identify specific pathways underlying the metabolic reprogramming of Axl^−/− PMs.

To further understand the metabolic dysregulation in Axl^−/− PMs, we performed a targeted metabolomic analysis focusing on glycolytic intermediates (Figure 3C). Axl deficiency resulted in a selective reduction in intracellular hexose (Hex) levels (Figure 3D), but not a significant reduction in intracellular lactate abundance (Figure 3H). In contrast, the levels of other glycolytic intermediates, including hexose phosphates (Hex‐P), glyceraldehyde‐3‐phosphate (G3P), pyruvate, and myo‐inositol remained unchanged between WT and Axl^−/− PMs (Figure 3E–G,I). These findings indicate that global Axl deficiency does not cause a uniform alteration across the glycolytic pathway but instead selectively affects specific nodes of glucose metabolism.

2.4. Axl Regulates Mitochondrial Metabolism in Peritoneal Macrophages

Given the alterations observed in glycolytic intermediates, we next investigated the TCA cycle to determine whether mitochondrial metabolism was also affected in Axl^−/− PMs (Figure 4A). Axl^−/− macrophages displayed markedly reduced acetyl‐CoA levels (Figure 4B), indicating impaired substrate availability for mitochondrial oxidation or inefficient entry of acetyl groups into the TCA cycle. Consistent with this deficit, early‐cycle intermediates, including citrate and cis‐aconitate, were significantly decreased (Figure 4C,D), demonstrating disruptions at the initial steps of carbon flux through the TCA pathway. These changes were accompanied by pronounced reductions in major redox cofactors: NAD⁺, NADP⁺, NADPH, and FAD (Figure 4K,Q–S), and in ATP (Figure 4N–P), collectively indicating compromised oxidative metabolism, diminished electron‐transfer capacity, and impaired energy production. In addition, α‐ketoglutarate (αKG) was significantly reduced (Figure 4E), further supporting defective mitochondrial carbon processing. In contrast, the abundance of downstream intermediates such as succinate, fumarate, malate, oxaloacetate (OAA), and itaconate remained unchanged (Figure 4F–J), suggesting that later stages of the TCA cycle are relatively preserved despite upstream bottlenecks. Supporting a compensatory metabolic response, several amino acids, including leucine, valine, methionine, alanine, histidine, phenylalanine, tyrosine, and tryptophan, were significantly depleted in Axl^−/− PMs (Figure 4U), consistent with increased anaplerotic amino acid catabolism to maintain mitochondrial flux under conditions of impaired acetyl‐CoA availability. Together, these findings reveal that global Axl deficiency induces profound defects in early TCA cycle entry and redox cofactor homeostasis, prompting compensatory amino acid utilization that partially sustains downstream mitochondrial reactions but fails to restore optimal oxidative metabolism.

2.5. Axl Deficiency Impairs Mitochondrial Maximal Respiratory Capacity in Peritoneal Macrophages

Given that Axl ^−/− PMs exhibit increased glycolytic activity and reduced nitric oxide production (Figure 2), and that the broad suppression of early TCA intermediates, redox cofactors, and high‐energy nucleotides (Figure 4) indicates a substantial limitation in mitochondrial oxidative capacity, we next asked whether these biochemical alterations translate into functional mitochondrial impairment. To address this, we evaluated the ability of Axl ^−/− PMs to adapt their metabolism when oxidative phosphorylation is challenged. We first assessed glycolytic compensation during impaired electron transport by measuring lactate production following treatment with rotenone, a Complex I inhibitor. In WT PMs, rotenone triggered a rapid and robust increase in lactate release over 60 min, consistent with a compensatory glycolytic shift that maintains ATP generation and regenerates NAD⁺ when NADH oxidation is blocked (Figure 5A). In contrast, Axl ^−/− PMs failed to proportionally elevate lactate levels under the same conditions, indicating reduced metabolic flexibility and an impaired ability to engage glycolysis in response to mitochondrial stress. To determine whether Axl deficiency alters mitochondrial respiratory function, oxygen consumption was measured in WT and Axl^−/− PMs by high‐resolution respirometry. Representative oxygen flux traces showed a markedly blunted response to FCCP in Axl^−/− PMs compared with WT controls (Figure 5B–D). In contrast, maximal respiration was significantly reduced in Axl^−/− PMs (Figure 5E), accompanied by a marked decrease in spare respiratory capacity (Figure 5F), indicating impaired mitochondrial reserve capacity and a diminished ability to augment oxidative metabolism in response to increased energetic demand. Notably, proton leak‐associated respiration was significantly increased in Axl^−/− PMs (Figure 5G), whereas residual oxygen consumption was unchanged (Figure 5H). Importantly, FCCP‐stimulated respiration reflects not only electron transport system capacity but can also depend on substrate availability and the ability to sustain metabolic flux into the TCA cycle [31]. These findings raised the possibility that the reduced maximal respiration and spare respiratory capacity observed in Axl^−/− PMs could reflect alterations in substrate availability in addition to intrinsic changes in mitochondrial respiratory function. To determine whether substrate availability contributed to the impaired respiratory phenotype of Axl^−/− PMs, they were subjected to substrate deprivation followed by metabolic rescue with glucose, pyruvate, and GlutaMAX. Substrate deprivation significantly reduced basal respiration in WT PMs, whereas basal respiration was unaffected in Axl^−/− PMs (Figure 5I). Repletion with these substrates partially restored basal respiration in both genotypes, resulting in comparable oxygen consumption rates following rescue. Notably, substrate deprivation markedly reduced maximal respiratory capacity in WT PMs (Figure 5J). Notably, the maximal respiratory rates reached by starved WT cells closely resembled those observed in untreated Axl^−/− PMs, suggesting that the respiratory phenotype associated with Axl deficiency mimics, at least in part, the metabolic consequences of substrate limitation. Following substrate repletion, maximal respiration increased in both genotypes, although WT PMs still exhibited higher respiratory rates than Axl^−/− cells. Consistent with these findings, spare respiratory capacity also tended to decrease following substrate deprivation in WT PMs and was only partially restored upon substrate repletion (Figure 5K). Together, these data suggest that the attenuated FCCP‐stimulated respiration in Axl^−/− PMs may not solely reflect an intrinsic limitation in electron transport system capacity, but may also arise from insufficient substrate availability to support oxidative metabolism under conditions of increased energetic demand, as proposed for FCCP‐driven respiratory responses [31]. The convergence of substrate‐deprived WT and Axl^−/− PMs on a similar respiratory phenotype further supports a role for Axl in maintaining the metabolic flexibility required to sustain maximal mitochondrial respiration. [31]

2.6. Axl Deficiency Impairs Glutathione Metabolism and Antioxidant Defenses in PMs

Since NADP⁺/NADPH, key cofactors in antioxidant defense [32], were reduced in Axl ^−/− PMs (Figure 5R,S), we next assessed metabolites directly involved in redox homeostasis. Reduced glutathione (GSH) was significantly decreased (Figure 6A), whereas the other components showed no significant changes (Figure 6B,C), indicating compromised glutathione metabolism. In contrast, ferulic acid and ophthalmic acid were increased (Figure 6D–E); however, these differences did not reach statistical significance. Nevertheless, these trends may be suggestive of a more oxidatively stressed state in Axl ^−/− PMs, consistent with a model in which loss of Axl could contribute to perturbations in central metabolism and redox homeostasis, increasing metabolic vulnerability under stress conditions.

2.7. Redox and Metabolic Dysfunction Extend to the Liver in the Absence of Axl Signaling

Given the redox imbalance and impaired antioxidant capacity in Axl^−/− PMs, we next investigated whether these alterations extended to other tissues within the peritoneal environment. We focused on the liver, a central organ in metabolic and redox regulation [33], by assessing markers of oxidative stress and antioxidant defenses in hepatic homogenates from WT and Axl^−/− mice. Although the levels of lipid peroxidation products, including thiobarbituric acid reactive substances (TBARS) and lipid hydroperoxides, were not reduced in Axl^−/− livers (Figure 7A,B), protein carbonylation was significantly increased (Figure 7C). In addition, thiol content and superoxide dismutase (SOD) activity were diminished (Figure 7D,F), while total antioxidant capacity (TAC) and catalase were elevated (Figure 7E,G), consistent with a compensatory response to heightened protein oxidation. Indicators of hepatic dysfunction accompanied these redox changes, including increased glycogen, reduced total protein, and elevated γ‐glutamyl transferase (γ‐GT) activity (Figure 7H–J). Together, these data indicate that Axl signaling is required for systemic redox and metabolic homeostasis. Although its loss drives cell‐intrinsic metabolic dysfunction in PMs, the associated oxidative stress phenotype extends to the liver, highlighting a broader role for Axl in coordinating immune and metabolic responses under homeostatic conditions.

4.1. Mice

Male and female C57BL/6 wild‐type (WT) and Axl^−/− mice, aged 8 to 12 weeks, were generously provided by Dr. Brian Kelsall (NIAID, NIH, Bethesda, MD, USA). Animals were maintained in the animal facility of the Universidade Federal do Rio de Janeiro under specific pathogen‐free (SPF) conditions, housed in individually ventilated cages with ad libitum access to food and water, and maintained on a 12 h light/dark cycle. All experimental protocols were reviewed and approved by the Institutional Animal Care and Use Committee (CEUA) of the Universidade Federal do Rio de Janeiro (protocol number 041/17).

4.2. Isolation of Peritoneal Macrophages

Resident peritoneal macrophages (PMs) were isolated by peritoneal lavage with 5 mL of cold phosphate‐buffered saline (PBS). The recovered fluid was collected with a syringe, centrifuged (1500 rpm, 15 min, 4 °C), and cells were resuspended in RPMI 1640 medium without phenol red (Gibco, Thermo Fisher Scientific) supplemented with 2 mM L‐glutamine, 50 mM β‐mercaptoethanol (Gibco, Life Techonologies), 100 mM sodium pyruvate (Gibco), 10 µg/mL gentamicin (Sigma‐Aldrich, Cotia, SP, Brazil), amino acids (Gibco), and 5% fetal bovine serum (FBS) (Embriolife). For in vitro assays, cells were plated for 1 h to allow adherence, followed by gentle washing to remove nonadherent cells and debris.

4.3. Peritoneal Macrophages Functional Assays and Metabolic Analyses

Adherent macrophages were stimulated to assess nitrite production, lactate, and lactate dehydrogenase (LDH) levels. For nitrites, lactate, and LDH quantification, macrophages were stimulated or not with ultrapure lipopolysaccharide (E. coli K12; 2000 ng/mL; Invivogen) for 24 h. Metabolic activity (lactate and LDH production) was further evaluated after treatment with rotenone (1 µg/mL, Sigma) for up to 60 min [53]. For rotenone experiments, macrophages were first allowed to adhere for 1 h, after which the medium was replaced with fresh RPMI containing rotenone. Lactate levels measured immediately after rotenone addition were considered the 0 min time point, and additional supernatants were collected at the indicated time points for subsequent lactate quantification.

4.4. MitoSOX Red Mitochondrial Superoxide Detection

Mitochondrial superoxide production was quantified using MitoSOX Red (Thermo Fisher Scientific), a fluorogenic probe that selectively targets mitochondria and fluoresces upon oxidation by superoxide. Peritoneal macrophages were incubated with 5 µM MitoSOX Red in PBS for 10 min at 37°C, protected from light. Cells were then washed twice, resuspended in fresh buffer, and immediately analyzed by flow cytometry. Mean fluorescence intensity (MFI) was used as a measure of mitochondrial ROS levels.

4.5. Cytosolic ROS Measurement Using Dihydroethidium (DHE)

Cytosolic reactive oxygen species (ROS) were quantified in PMs using dihydroethidium (DHE; Thermo Fisher Scientific). Cells were seeded in 96‐well plates and incubated with 5 µM DHE diluted in HBSS. Parallel wells were left unstimulated or stimulated with LPS (2000 ng/mL) for 60 min. DHE fluorescence was recorded kinetically using a SpectraMax Paradigm plate reader, with excitation at 520 nm. Five consecutive measurements were acquired at 15 min intervals over the 60 min incubation period. ROS production was expressed as mean fluorescence intensity.

4.6. Flow Cytometry

PMs were first incubated with anti‐CD16/CD32 (Fc block, BD Pharmingen, 1:200) for 20 min on ice to block Fc receptors. Cells were then stained for 30 min with the following fluorochrome‐conjugated antibodies: PerCP‐anti‐CD11b (clone M1/70, BioLegend, 1:750), PE‐Cy5‐anti‐CD19 (clone eBio1D3, eBioscience, 1:750), PE‐Cy7‐anti‐F4/80 (clone BM8, BioLegend, 1:250), eFluor450‐anti‐mouse CD206 (clone MR6F36‐10A1, eBioscience, 1:750), and APC‐Cy7‐anti‐I‐A/I‐E (clone M5/114.15.2, BioLegend, 1:750). After staining, cells were washed with PBS containing 2 mM EDTA and 50% FBS. The cells were fixed with 2% paraformaldehyde and acquired on a BD LSRFortessa flow cytometer. Data were analyzed using FlowJo software (Tree Star Inc.).

4.7. Nitric Oxide Assay

Nitric oxide production was quantified indirectly by measuring nitrite (NO₂ ⁻) accumulation in the cell culture supernatant using the Griess reagent, according to a previously described colorimetric assay [54].

4.8. Lactate Measurement

Lactate levels in macrophage culture supernatants were quantified using an enzymatic assay kit (Bioclin, Brazil) according to the manufacturer's instructions. The assay is based on the conversion of L‐lactate to pyruvate by LDH, coupled to NAD⁺ reduction, with absorbance measured at 340 nm. Samples (5 µL) were incubated with 195 µL of working reagent in a 96‐well plate at 37°C for 5 min. Absorbance was read at 340 nm using a Spectra Max Paradigm plate reader (Molecular Devices, USA), and lactate concentration was calculated based on a standard curve.

4.9. LDH Activity Assay

LDH activity was determined using a kinetic assay kit (Bioclin, Brazil) by monitoring NADH consumption at 340 nm at 37°C, according to the manufacturer's instructions. Absorbance was recorded over 4 min, and enzyme activity was calculated based on the rate of absorbance decrease (ΔA/min × 8016), expressed in U/L.

4.10. Polar Metabolite Extraction and LC‐MS Analysis

Polar metabolites were extracted from 2–8 × 10⁶ PMs using a chloroform:methanol:water (2:2:1.8, v/v/v) solution as previously described [55]. p‐Nitrophenyl phosphate (10 nmol) was added at the beginning of the extraction procedure as an internal standard. Polar fractions were dried under vacuum centrifugation, resuspended in 25 µL of 70% acetonitrile in water, and 10 µL of each sample was injected into a Nexera X2 UHPLC system (Shimadzu) coupled to a Maxis Impact Q‐TOF mass spectrometer (Bruker) equipped with an electrospray ionization (ESI) source. Chromatographic separation was performed on a SeQuant ZIC‐pHILIC column (150 × 2.1 mm, 5 µm particle size; Millipore‐Merck) maintained at 35°C, using a 0.175 mL/min flow rate. A discontinuous gradient of mobile phase A (20 mM ammonium carbonate, pH 9.0 in water) and mobile phase B (acetonitrile) was applied following Ahn et al. (2017): 0–5 min: 95% B; 5–35 min: 95% → 30% B; 35–45 min: 30% B (wash); 45–60 min: 95% B (re‐equilibration). Mass spectra were acquired in negative ion mode, over a 30–1200 m/z range, at 0.67 Hz, without fragmentation. ESI source parameters were set as follows: nebulizer gas (N₂), 2.0 bar; drying gas (N₂), 8.0 L/min; source temperature, 200°C; capillary voltage, 4500 V. External mass calibration was performed using a 100 µM sodium formate solution. To support metabolite identification, a pooled quality control sample was analyzed in data‐dependent acquisition (DDA) mode, targeting the top 5 precursor ions per cycle for isolation and fragmentation at 15 and 30 eV collision energies, with all other MS parameters identical to full‐scan acquisitions.

Raw LC–MS data were processed using MZmine 2 [56], and multivariate statistical analyses were performed on the MetaboAnalyst online platform [57, 58]. For targeted analysis of specific metabolites, raw data files were processed in QuantAnalysis 5.0 (Bruker), and univariate statistical analyses were conducted using GraphPad Prism 8.0. Metabolite features were annotated based on accurate masses (10 ppm tolerance) and fragmentation spectra, using the HMDB database [59], mzCloud (www.mzcloud.org), and Metlin [60]. Quantitative results were expressed as the ratio of the metabolite peak area target (RAT) to the internal standard (ISTD), normalized to the total number of cells in each sample (RAT/ISTD/cell number). This normalization strategy was used to minimize technical variability and account for differences in cell input between samples.

4.11. High‐Resolution Respirometry Assay

The respirometry experiments were conducted using the Oroboros Oxygraph‐2k system, following protocols previously described [61, 62]. WT and Axl^−/− peritoneal macrophages were cultured in RPMI 1640 medium without phenol red and seeded at a density of 1 million cells per chamber in the Oroboros O2k respirometer. The following inhibitors and uncoupler were used as described: 0.5 µg/mL of oligomycin was added to the chamber to inhibit ATP synthase, followed by the uncoupling agent, carbonyl cyanide‐4‐(trifluoromethoxy) phenylhydrazone (FCCP), which was titrated up to a final concentration of 2 µM. Lastly, 4 µM of antimycin A, the inhibitor of mitochondrial complex III, was added to the chamber. The mitochondrial physiology parameters were assessed under specific experimental conditions. Residual oxygen consumption (Rox) was determined after the addition of Antimycin A to establish a baseline for respiratory activity not associated with oxidative phosphorylation (OXPHOS). Basal cellular respiration was computed by subtracting non‐mitochondrial respiration from the basal respiration recorded at the beginning of the experiment, before any of the additions. O₂ flux linked to OXPHOS was calculated as the difference between basal respiration and oligomycin‐insensitive respiration after correction for residual oxygen consumption. Maximal respiration was determined in the presence of FCCP, with non‐OXPHOS respiration subtracted to isolate the mitochondrial contribution to this maximal activity. Spare respiratory capacity was calculated as the difference between maximal and basal respiration after correction for residual oxygen consumption. Lastly, the measurement of H⁺ leak was performed by evaluating the effects induced by oligomycin on respiration, with the nonmitochondrial component subtracted for precise evaluation. Data were analyzed using Oroboros DatLab software.

4.12. Substrate Deprivation and Respiratory Rescue Assay

To evaluate whether limited substrate availability could contribute to the respiratory phenotype observed in Axl‐deficient macrophages, WT and Axl^−/− peritoneal macrophages were subjected to acute nutrient deprivation before high‐resolution respirometry analysis. Cells were incubated for 1 h in Dulbecco's modified Eagle's medium (Sigma‐Aldrich). Following nutrient deprivation, macrophages were transferred to the Oroboros O2k respirometer at a density of 1 × 10⁶ cells per chamber. Basal oxygen consumption was initially recorded under substrate‐deprived conditions. Subsequently, 1.6 mM of GlutaMAX supplement (Thermo Fisher Scientific), 25 mM of glucose, and 1 mM sodium pyruvate were added directly to the chamber at the concentrations used in the culture medium. Oxygen consumption was monitored to assess respiratory recovery following substrate replenishment. After stabilization of oxygen flux, mitochondrial respiratory parameters were determined as previously described.

4.13. Protein Quantification

Protein levels in tissue homogenates were measured by the Bradford (Thermo Fisher Scientific) assay. Absorbance was read at 595 nm (SpectraMax Paradigm), and concentrations were calculated from the bovine serum albumin (Sigma‐Aldrich) standard curve. Results were used to normalize oxidative stress assays.

4.14. Liver Preparation

Liver tissue was collected at euthanasia, aliquoted, and stored at –80°C until the day of analysis. For this, 200 mg of the tissue was homogenized in 1 mL antioxidant buffer (Tris‐HCl 10 mM, pH 8.0, saline 0.9%, aprotinin, PMSF, MilliQ water) using Ultra‐Turrax (T25, IKA, Staufen, Alemanha), kept on ice. Samples were centrifuged (1000 rpm, 15 min, 4°C), supernatants collected and stored at –80°C for oxidative stress and antioxidant enzyme assays [63].

4.15. Lipid Peroxidation Assay

Tissue homogenates were mixed with phosphate buffer and 10% tricarboxylic acid, cooled, and centrifuged. Supernatants reacted with thiobarbituric acid substrates (TBAS) and were heated at 100°C for 20 min. After cooling, absorbance was measured at 532 nm (Spectra Max Paradigm). Concentrations were calculated using a TMP (tetramethyl‐p‐phenylenediamine) standard curve and expressed as μ[TMP]/mg protein [64].

4.16. Total Hydroperoxide Measurement

Tissue homogenates were mixed with 40 mM butylated hydroxytoluene (BHT) in methanol, vortexed, and centrifuged (6000 rpm, 5 min, 4 °C). Supernatants (20 µL) were incubated with 360 µL of FOX reagent composed of 100 µM xylenol orange, 250 µM ammonium ferrous sulfate, 25 mM sulfuric acid, and 90% ethanol (v/v). After 2 h of incubation in the dark at room temperature, absorbance was measured at 540 and 560 nm using a Spectra Max Paradigm plate reader. Hydroperoxide levels were determined from the absorbance difference (ΔA) and expressed as µM/mg protein [65].

4.17. Protein Carbonyl Content Assay

Tissue homogenates were mixed with 2,4‐dinitrophenylhydrazine (DNPH) or phosphate buffer (blank), incubated for 10 min at room temperature in the dark, followed by addition of NaOH and further 10 min incubation. Absorbance was measured at 450 nm (SpectraMax Paradigm). Carbonyl concentration was calculated using the molar extinction coefficient 22,308 M^{− 1}cm^{− 1} and expressed as µmol/mg protein [66].

4.18. Thiol Group Assay

Tissue homogenates were prepared as blanks (sample + thiol buffer + methanol) and test samples (sample + thiol buffer + methanol + DTNB). After 15 min incubation at room temperature and centrifugation (5000 rpm, 15 min, 25°C), absorbance of the NTB₂ ⁻ ion was measured at 412 nm (Spectra Max Paradigm). Concentrations were calculated using the molar extinction coefficient 13,600 M^{− 1}cm^{− 1} and expressed as mM NADPH/mg protein [67].

4.19. Total Antioxidant Capacity Assay

Samples (0.5 mL) were mixed with 3 mL absolute ethanol and 0.3 mL of 0.5 mM DPPH (2,2‐difenil‐1‐picrilhidrazila) solution in ethanol. After 100 min incubation, absorbance was measured at 517 nm (Spectra Max Paradigm). Antioxidant activity was calculated from a DPPH standard curve and expressed as mM DPPH [68].

4.20. Catalase Activity Assay

Liver homogenates (10 µL) were added to 990 µL phosphate buffer and incubated at 37°C. The reaction was started by adding 5 µL of 30% hydrogen peroxide, and absorbance at 240 nm was measured every 15 s for 90 s (Spectra Max Paradigm). Catalase activity was calculated from the linear decrease in absorbance and expressed as k·mg^{− 1} protein [69].

4.21. Superoxide Dismutase Activity Assay

Enzyme activity was measured by the inhibition of adrenaline auto‐oxidation to adrenochrome at 480 nm. Samples (10 µL) were mixed with glycine buffer (50 mM, pH 10.2) and catalase (0.0024 g/mL), incubated at 32°C for 1 min, then adrenaline (60 mM) was added to start the reaction. Absorbance was recorded every 10 s for 180 s (SpectraMax Paradigm). Activity was calculated by subtracting blank readings (auto‐oxidation without sample) and expressed as U/mg protein [70].

4.22. Hepatic Glycogen Quantification

Liver homogenates were assayed for glycogen using a Bioclin enzymatic kit, following manufacturer instructions. Absorbance was measured in a microplate reader and glycogen content calculated from the standard curve.

4.23. γ‐Glutamyl Transferase Activity Assay

γ‐glutamyl transferase (γ‐GT) activity was measured in tissue homogenates using a commercial enzymatic kit (Bioclin), following the manufacturer's protocol. Samples were incubated with reagents that generate a colorimetric reaction proportional to γ‐GT activity. Absorbance was read using a microplate reader, and enzyme activity was calculated according to the standard curve.

4.24. Statistical Analyses

All statistical analyses were performed using GraphPad Prism software (version 9.0). Comparisons between two groups were conducted using Student's t‐test. For analyses involving more than two groups, one‐way analysis of variance (ANOVA) followed by Tukey's multiple comparisons post hoc test was applied. Detailed information regarding statistical significance is provided within the figures and corresponding legends.