Alpha-1 Antitrypsin Protects Against Cisplatin-Induced Acute Kidney Injury by Restoring Redox and Mitochondrial Homeostasis

Abstract

Cisplatin is an effective chemotherapeutic agent, yet its clinical utility is limited by dose-dependent nephrotoxicity. Alpha-1 antitrypsin (AAT) has cytoprotective, anti-inflammatory, and antiapoptotic properties, but its therapeutic potential in cisplatin-induced acute kidney injury (AKI) remains unclear. A murine cisplatin–AKI model was used to evaluate whether AAT (80 mg/kg) ameliorates renal injury. Renal function, oxidative stress, NADPH oxidase (NOX) isoforms, mitochondrial metabolism, inflammatory mediators, apoptosis, and fibrosis-related markers were assessed using biochemical, histological, immunohistochemical, and Western blot analyses. Cisplatin markedly impaired renal function and induced tubular injury; meanwhile, AAT significantly reversed these changes. Cisplatin also induced severe oxidative stress and disrupted the balance of NOX isoforms; AAT restored redox homeostasis. Cisplatin upregulated CPT1A/PDK4 and suppressed CPT2, UCP3, PGC1α, and DRP1, inducing maladaptive mitochondrial changes, indicating impaired β-oxidation and defective mitochondrial dynamics; AAT reversed these alterations, restoring normal mitochondrial metabolism. IL-1β, IL-6R, OPN, and F4/80 expression, recovery of the Bax/Bcl-2 ratio, and MAPK activation were reduced, indicating decreased inflammation and apoptosis; profibrotic markers were also reduced. AAT confers multifaceted protection against cisplatin-induced AKI by restoring redox balance, mitochondrial homeostasis, and inflammatory and apoptotic signaling. These findings support AAT as a promising therapeutic agent for preventing cisplatin nephrotoxicity.

1. Introduction

Cisplatin is a chemotherapeutic agent that is widely used to treat various solid tumors; however, the clinical utility of cisplatin is limited by dose-dependent nephrotoxicity, which frequently leads to acute kidney injury (AKI) and can progress to chronic kidney disease (CKD) [1,2]. Cisplatin-induced AKI is characterized by proximal tubular cell injury, oxidative stress, mitochondrial dysfunction, inflammation, and in more severe or prolonged settings, fibrotic remodeling of the tubulointerstitium [3]. Current preventive strategies, primarily hydration and dose adjustment, offer only partial protection. Moreover, no specific pharmacologic therapy has yet been approved that reliably prevents cisplatin-induced nephrotoxicity [1,2,3]. Thus, there remains a continuing need to identify novel renoprotective agents that can mitigate cisplatin-induced injury without compromising antitumor efficacy.

Oxidative stress and mitochondrial injury are among the key pathogenic mechanisms implicated in cisplatin-induced nephrotoxicity and are considered central drivers of tubular damage [4,5]. NADPH oxidase (NOX)-derived reactive oxygen species (ROS), particularly those generated by NOX isoforms, NOX1, NOX2, and NOX4, contribute to DNA damage, lipid peroxidation, and activation of proinflammatory and proapoptotic signaling pathways [6,7,8]. Although NOX4 promotes cisplatin-induced tubular cell death and inflammatory responses [7], experimental models of ischemic AKI have shown that NOX4 deficiency can exacerbate tubular injury, suggesting context-dependent and potentially adaptive roles for NOX isoforms [9]. NOX2-mediated ROS production further amplifies proximal tubular damage and inflammation during cisplatin nephrotoxicity [10]. Moreover, perturbations in the NAD⁺/NADH redox balance, impaired mitochondrial bioenergetics, mitochondrial-derived ROS generation, and dysregulated mitophagy have emerged as key contributors to the propagation of tubular injury [5,11]. Accordingly, small-molecule interventions targeting NOXs, Nrf2/HO-1 signaling, or mitochondrial redox homeostasis have demonstrated protective effects in cisplatin-induced AKI, underscoring the therapeutic potential of redox modulation [12,13,14].

Alpha-1 antitrypsin (AAT) is a 52 kDa serine protease inhibitor, classically known for its role in neutralizing neutrophil elastase, and for its established use in AAT deficiency; however, accumulating evidence indicates that AAT exerts broad anti-inflammatory, antiapoptotic, antioxidative, and cytoprotective actions [12,15,16]. Furthermore, AAT has been shown to modulate immune responses, cytokine signaling, and cellular stress pathways in multiple tissues [14,15,16,17,18,19]. Indeed, exogenous AAT administration in renal injury models protects against ischemia–reperfusion injury and tacrolimus-induced nephrotoxicity by reducing tubular necrosis, inflammatory infiltration, and fibrotic remodeling [19,20,21,22]. AAT also inhibits epithelial-to-mesenchymal transition (EMT) and extracellular matrix accumulation [23] and suppresses apoptosis in human peritoneal mesothelial cells exposed to toxic stimuli [18]. Clinically, circulating AAT levels correlate with AKI severity and postoperative outcomes in cardiac surgery patients, suggesting a potential endogenous protective role during acute renal stress [24,25].

Despite these observations, studies on whether AAT administration can mitigate cisplatin-induced AKI remain limited. Thus, considering the key pathogenic pathways involved in cisplatin nephrotoxicity [1,3,5,12] and the broad cytoprotective actions of AAT [16,17,18,23,26], we hypothesized that AAT would mitigate cisplatin-induced renal injury. Therefore, this study aimed to investigate the effects of AAT on renal function, tubular damage, oxidative stress, NOX isoform expression, mitochondrial metabolic homeostasis, inflammation, apoptosis, and early fibrotic remodeling in a murine model of cisplatin-induced AKI.

2. Material and Methods

2.1. Animals Model

All experiments were performed on eight-week-old male C57BL/6 mice weighing 22–25 g (Samtako, Osan, Republic of Korea). The animals were housed under a 12 h light/dark cycle, with free access to standard chow and water. The animal study protocol was approved by the Institutional Animal Care and Use Committee (IACUC) of Kyungpook National University (approval no. KNU-2024-0001, approval date 4 January 2024). The mice were randomly assigned to four experimental groups (n = 6 per group): (1) control, (2) AAT (80 mg/kg), (3) cisplatin (20 mg/kg), and (4) cisplatin (20 mg/kg) + AAT (80 mg/kg). The AAT treatment was performed as previously described in our earlier study [23]. Briefly, AAT was dissolved in phosphate-buffered saline (PBS) and administered once daily by intraperitoneal injection for five consecutive days. AAT was dissolved in phosphate-buffered saline (PBS) and administered once daily by intraperitoneal injection for five consecutive days. Cisplatin was administered as a single intraperitoneal injection (20 mg/kg) on day 3, in accordance with commonly used murine models of cisplatin-induced AKI/nephrotoxicity [27,28], and was co-administered with AAT in the cisplatin + AAT group. Control mice received an equivalent volume of PBS. All mice were sacrificed on day 6 after treatment initiation, and blood and kidney tissues were collected for subsequent biochemical, histological, and molecular analyses.

2.2. Kidney Function and Histopathological Studies

The mouse serum blood urea nitrogen (BUN) and creatinine levels were evaluated by GCLabs (Yongin, Republic of Korea) using the Cobas 8000 modular analyzer system (Roche, Mannheim, Germany). Kidney tissue from each experimental group was immersion-fixed in 4% paraformaldehyde (pH 7.4) and then embedded in paraffin. Tissue samples were prepared in 2 μm sections and stained with periodic acid-Schiff (PAS) and Masson’s trichrome using standard protocols to determine the histological changes and collagen deposition, respectively. Images were acquired at 200× and 400× magnification. For quantification, 10 non-overlapping microscopic fields per mouse were randomly selected from the renal cortex and analyzed in a blinded manner. Tubular injury in PAS-stained sections was semi-quantitatively scored (0–4) based on the percentage of tubules showing cast formation, tubular dilation, brush border loss, and epithelial necrosis (0, none; 1, <25%; 2, 25–50%; 3, 50–75%; 4, >75%). Regions of collagen deposition in Masson’s trichrome-stained kidney sections were measured using the ImageJ program. A consistent threshold was applied across all groups, and collagen deposition was expressed as the percentage of trichrome-positive area.

2.3. Immunohistochemistry

Kidney tissue from each experimental group was immersion-fixed in 4% paraformaldehyde (pH 7.4), and immunolabeling was performed on paraffin-embedded sections. Immunohistochemical staining was performed using anti-fibronectin (1:100; Abcam, Cambridge, UK), anti-F4/80 (1:500; Cell Signaling, Danvers, MA, USA), anti-osteopontin (OPN) (1:1000; MPIIIB10, obtained from the Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA, USA), anti-4-HNE (1:200; Bioss, Woburn, MA, USA), and anti-8-OHdG (1:100; Santa Cruz Biotechnology, Santa Cruz, CA, USA) antibodies. An EnVision-HRP kit (Dako, Carpinteria, CA, USA) was used to detect the staining. All sections were counterstained using Mayer’s hematoxylin. A Leica DM IRB inverted microscope (Leica Microsystems, Wetzlar, Germany) equipped with a CoolSNAP HQ camera (Photometrics, Tucson, AZ, USA) was used to examine immunolabeling. Images were acquired at 200× and 400× magnification. The positive area was quantified using ImageJ (version 1.54p, NIH, Bethesda, MD, USA). For quantification, 10 non-overlapping microscopic fields per mouse were randomly selected from the renal cortex and analyzed. A consistent threshold was applied across all groups, and quantification was performed in a blinded manner.

2.4. Quantitative Real-Time Polymerase Chain Reaction

Total RNA was extracted from kidney tissue using TRIzol^TM Reagent (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s instructions. RNA purity and concentration were determined using a UV-VIS Spectrophotometer (DeNovix, Wilmington, DE, USA). Total RNA (1 μg) was reverse transcribed using the Prime Script cDNA Synthesis kit (Takara Shuzo Co., Otsu, Shiga, Japan). All primers used in the quantitative real-time polymerase chain reaction (RT-qPCR) were determined using Primer (

Table 1. Oligonucleotide primer sequences.

Gene	Forward Primer (5′–3′)	Reverse Primer (5′–3′)
Mouse Nox1	CAG GCC ATG GAT GGA TCT CT	ATG TTT GGA GAC TGG ATG GGA TT
Mouse Nox2	GCT CTC TCT GAC ATC GGT GAC A	CGA GTC ACG GCC ACA TAC AG
Mouse Nox4	CAC CAA ACA CAG AAG CAC AAG AC	AAA GCA GGG TAT CAC TCC ATG AA
Mouse Sod2	GGT AGG GCC TGT CCG ATG AT	GTC CAG TTC TCC AGA GAT ATA CAA TTC A
Mouse Cat	CGA CCA GGG CAT CAA AAA CT	ATT GGC GAT GGC ATT GAA A
Mouse Gapdh	TAA AGG GCA TCC TGG GCT ACA CT	TTA CTC CTT GGA GGC CAT GTA GG

) Express V1.5 software (Applied Biosystems, Foster City, CA, USA). Real-time qPCR was performed in duplicate for each sample to evaluate the expression of Nox1, Nox2, Nox4, Sod2, Cat, and Gapdh. The PCR reaction was pre-denatured at 50 °C for 2 min, then heated to 95 °C for 10 min, followed by 40 cycles at 95 °C for 15 s and 60 °C for 1 min. The results were normalized to the corresponding ΔCt values for the control primer set, Gapdh mRNA, for which fold enrichment was calculated as 2^{−[ΔCt(assayed gene)−ΔCt(GAPDH)]}. The relative changes in the assayed gene mRNA/Gapdh mRNA ratio between the control group and sample group were determined using the 2^−ΔΔCt formula.

2.5. Immunoblot Analysis

Kidney tissues were homogenized and centrifuged at 4000× g and 4 °C for 15 min. Protein samples (30 μg) were mixed in 5 × SDS reducing buffer (250 mM Tris–HCl, pH 6.8, 0.5 M dithiothreitol (DTT), 10% sodium dodecyl sulfate (SDS), 0.25% bromophenol blue, and 50% glycerol), boiled, resolved on 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) gels, and transferred to a polyvinylidene fluoride (PVDF) membrane by electroblotting. The following primary antibodies were used to detect protein expressions: NGAL (1:1000; ab216462, Abcam), TNF-α (1:1000; ab6671, Abcam), IL-1β (1:2000; 12242S, Cell Signaling Technology, Danvers, MA, USA), IL-6R (1:1000; ab300581, Abcam), Bax (1:1000; 2772S, Cell Signaling Technology), Bcl-2 (1:1000; ab196495, Abcam), CPT1A (1:1000; ab128568, Abcam), CPT2 (1:500; ab181114, Abcam), PDK4 (1:1000; ab214938, Abcam), UCP3 (1:1000; DA1-24895, Thermo Fisher Scientific, Waltham, MA, USA), PGC1α (1:500; ab191838, Abcam), DRP1 (1:1000; ab184247, Abcam), fibronectin (1:500; ab2413, Abcam), α-SMA (1:2000; A2547, Sigma, Burlington, MA, USA), p-JNK (1:1000; ab124956, Abcam), total JNK (1:1000; ab179461, Abcam), p-ERK (1:1000; 4370S, Cell Signaling Technology), total ERK (1:1000; 9102S, Cell Signaling Technology), p-p38 (1:1000; 9211S, Cell Signaling Technology), total p38 (1:1000; 9212S, Cell Signaling Technology), and GAPDH (1:5000; 2118S, Cell Signaling Technology). The horseradish peroxidase-conjugated secondary antibodies (Dako, Glostrup, Denmark) corresponding to each primary antibody type, and enhanced chemiluminescence (ECL) advanced detection (GE Healthcare, Little Chalfont, UK), were used. Positive immunoreactive bands were quantified by densitometry and compared to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) expression, used as an internal control. Scion Image software (version 4.0.3, Scion, Frederick, MD, USA) was used for the density quantification.

2.6. Statistical Analysis

Data are presented as the mean ± standard error of the mean (SEM). Experiments were repeated at least three times. Statistical analyses were performed using GraphPad Prism version 5.01 (GraphPad Software, La Jolla, CA, USA) and SPSS version 22 (IBM Corp., Armonk, NY, USA). For comparisons among multiple groups, one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test was applied for normally distributed data. When the assumption of normality was not met, nonparametric analysis was applied using the Kruskal–Wallis test followed by Dunn’s multiple comparison test. A p-value < 0.05 was considered statistically significant.

3. Results

3.1. AAT Treatment Attenuates Cisplatin-Induced Renal Dysfunction and Tubular Damage

The in vivo experimental design is summarized in Figure 1A. Mice were randomly assigned to four groups (control, AAT, cisplatin, and cisplatin + AAT), and blood and kidney tissue were collected for analysis. Serum biochemical analyses revealed that cisplatin treatment markedly increased BUN and serum creatinine levels compared with the control group (Figure 1B,C). AAT co-administration significantly attenuated these elevations, indicating preserved renal function. Western blot analysis further demonstrated a strong upregulation of NGAL, a sensitive marker of tubular injury, in cisplatin-treated kidneys, whereas AAT co-administration significantly reduced NGAL expression (Figure 1D). Severe tubular necrosis, epithelial desquamation, and loss of brush border integrity were observed in the histopathologic evaluation of cisplatin-treated kidneys using PAS staining. In contrast, the normal tubular architecture was largely preserved after AAT administration (Figure 1E). These results indicate that AAT mitigates both functional and structural renal damage in cisplatin-induced AKI.

3.2. AAT Reduces Oxidative Stress in Cisplatin-Induced Acute Kidney Injury

Immunohistochemical staining revealed strong expression of oxidative stress markers 8-hydroxy-2′-deoxyguanosine (8-OHdG) and 4-hydroxynonenal (4-HNE) in the renal cortex of cisplatin-treated mice, which was markedly reduced by AAT co-administration (Figure 2A–C). Interestingly, differential regulation of NADPH oxidase isoforms was observed in the RT-qPCR analysis: Nox1 expression was significantly increased by cisplatin and normalized by AAT. In contrast, Nox4 expression was reduced following cisplatin exposure and partially restored by AAT, whereas Nox2 showed a similar trend but did not reach statistical significance (Figure 2D).

Moreover, cisplatin significantly decreased the expression of the antioxidant enzymes Sod2 and Cat, both of which were restored by AAT treatment (Figure 2E). Collectively, these findings demonstrate that AAT attenuates oxidative stress by modulating the expression of NADPH oxidase isoforms and restoring antioxidant defenses.

3.3. AAT Suppresses Inflammatory Cell Infiltration and Apoptosis

Western blot analysis showed that cisplatin slightly increased TNF-α expression, although not to levels of statistical significance. In contrast, the IL-1β and IL-6R levels were significantly elevated and clearly suppressed by AAT co-treatment (Figure 3A). Meanwhile, apoptotic signaling was assessed by measuring Bax and Bcl-2 expression. Cisplatin increased the expression of the proapoptotic Bax protein and decreased the expression of the antiapoptotic Bcl-2 protein, resulting in an increased Bax/Bcl-2 ratio, which was restored by AAT treatment (Figure 3B). Renal inflammation was further evaluated by immunostaining for the macrophage markers F4/80 and osteopontin (OPN). Both markers were markedly upregulated in cisplatin-treated kidneys but were significantly reduced following AAT administration (Figure 3C–E). Collectively, these findings indicate that AAT attenuates cisplatin-induced inflammatory responses and apoptotic signaling, accompanied by reduced IL-1β expression and macrophage infiltration.

3.4. AAT Preserves Mitochondrial Metabolic Homeostasis and Dynamics

Western blot analysis of mitochondrial metabolic and dynamics-related proteins showed that cisplatin significantly reduced the expression of CPT2, UCP3, PGC1α, and DRP1, while significantly increasing the expression of CPT1A and PDK4 compared with control kidneys (Figure 4A,B). AAT co-administration effectively reversed these cisplatin-induced alterations. These findings indicate that cisplatin induces a coordinated but maladaptive mitochondrial metabolic profile characterized by enhanced upstream fatty acid entry signaling (CPT1A, PDK4), coupled with suppression of downstream β-oxidation capacity (CPT2, UCP3), mitochondrial biogenesis (PGC1α), and fission-related regulation (DRP1). AAT treatment effectively rebalanced this mitochondrial program, restoring key metabolic and regulatory components toward a physiological range.

3.5. AAT Attenuates Renal Fibrosis and Extracellular Matrix Deposition

Western blot analysis demonstrated that cisplatin significantly increased the expression of fibronectin and α-smooth muscle actin (SMA), indicating activation of profibrotic and EMT-associated pathways (Figure 5A). AAT co-administration significantly attenuated the cisplatin-induced upregulation of both fibronectin and α-SMA. Consistent with these molecular changes, Masson’s trichrome staining revealed marked collagen deposition and interstitial fibrosis in cisplatin-treated kidneys, which were significantly reduced by AAT treatment (Figure 5B,C). In parallel, fibronectin immunohistochemistry showed prominent extracellular matrix accumulation in cisplatin-treated kidneys, whereas AAT co-treatment significantly reduced the area of fibronectin positivity (Figure 5B,D). These findings demonstrate that AAT significantly attenuates cisplatin-induced renal fibrotic remodeling, accompanied by the suppression of EMT-associated marker expression and extracellular matrix deposition.

3.6. AAT Modulates MAPK Signaling Pathways

To further examine the molecular mechanisms of AAT protection, the phosphorylation levels of ERK, JNK, and p38 were assessed. Cisplatin markedly increased the levels of phosphorylated ERK, JNK, and p38 relative to total protein, whereas AAT co-treatment significantly reduced these phosphorylation events (Figure 6A,B). These results indicate that AAT attenuates cisplatin-induced activation of the MAPK pathway, which is commonly associated with inflammatory and apoptotic signaling.

4. Discussion

Cisplatin-induced AKI arises from closely interconnected processes, including oxidative stress, mitochondrial dysfunction, inflammation, apoptosis, and fibrotic remodeling [1,2,5]. Our study demonstrates that AAT exerts multifaceted renoprotective effects by modulating these critical pathogenic pathways in a coordinated manner. Thus, AAT interrupts early ROS-driven signals that initiate tubular injury by reducing oxidative stress and normalizing NOX activity. In parallel, AAT also preserves mitochondrial metabolic integrity and biogenesis, thereby preventing the energetic collapse that amplifies cisplatin-mediated cellular dysfunction. The suppression of IL-1β-driven inflammatory activation and the restoration of antiapoptotic signaling further underscore the ability of AAT to counteract downstream inflammatory and cell death-related pathways. Moreover, by limiting EMT-associated profibrotic signaling, AAT mitigates maladaptive tissue remodeling that contributes to long-term renal impairment.

Overall, these findings support an integrated mechanistic model in which cisplatin induces NOX1-dominant oxidative stress, mitochondrial metabolic dysregulation, inflammatory and apoptotic signaling, and subsequent EMT-associated fibrotic remodeling in the kidney. As summarized in Figure 7, AAT counteracts these pathogenic processes by rebalancing NOX isoform activity and antioxidant defenses, preserving mitochondrial metabolic homeostasis and dynamics, suppressing inflammatory and apoptotic pathways, and limiting extracellular matrix accumulation and fibrosis.

Oxidative stress represents a principal trigger of cisplatin-induced tubular injury [1,2,5]. Excess ROS production originates from both mitochondrial electron leakage and dysregulated NOX activity, particularly NOX1, NOX2, and NOX4, leading to lipid peroxidation, DNA damage, and redox imbalance [6,8,9,10]. Consistent with this pathogenic sequence, AAT attenuated oxidative injury and restored endogenous antioxidant defenses. These findings align with prior reports showing that AAT reduces oxidative tissue damage in various organ systems by modulating redox signaling and suppressing NOX-mediated ROS generation [19,21,22]. Hence, AAT may prevent the downstream amplification of inflammatory and apoptotic pathways through early interruption of oxidative stress.

Importantly, cisplatin induced distinct isoform-specific patterns of NOX dysregulation, with Nox1 markedly upregulated, and Nox2 and Nox4 strongly suppressed. This divergence is consistent with prior evidence indicating that NOX1 primarily drives pathological epithelial ROS generation, whereas NOX2 and NOX4 can participate in more homeostatic or adaptive redox signaling, depending on the injury context [6,7,9,10,29]. AAT administration normalized this imbalanced isoform profile by preventing aberrant Nox1 induction and restoring Nox4 expression, with Nox2 showing a trend toward recovery. Interestingly, AAT appears to selectively suppress injurious NOX1-derived ROS while re-establishing physiological NOX2/NOX4 activity rather than globally reducing NOX isoforms. This interpretation is supported by the overall reduction in oxidative damage markers, such as 8-OHdG and 4-HNE, along with the restoration of antioxidant enzymes, including SOD2 and catalase, indicating a net decrease in oxidative stress accompanied by rebalancing of NOX isoform expression. Thus, AAT effectively shifts redox signaling from pathological NOX1-driven ROS generation toward a more physiological redox state.

Mitochondrial dysfunction is another hallmark of cisplatin nephrotoxicity, characterized by impaired β-oxidation, altered NAD⁺/NADH balance, dysregulated mitophagy, and disturbed mitochondrial dynamics [3,4,11,30,31]. In our study, cisplatin induced a distinct pattern of metabolic and structural mitochondrial dysregulation, with CPT1A and PDK4 upregulated and CPT2, UCP3, PGC1α, and DRP1 markedly reduced. This profile suggests an incomplete and maladaptive metabolic response. Meanwhile, upregulation of CPT1A and PDK4 is typically observed under metabolic stress and reflects a shift toward greater reliance on fatty acid substrates and inhibition of pyruvate dehydrogenase [32,33,34,35,36]; however, the concomitant downregulation of CPT2 and UCP3 indicates that downstream β-oxidation and controlled proton leaking are impaired [30,37,38]. Simply, tubular cells appear to “attempt” to enhance fatty acid oxidation at the entry point (CPT1A/PDK4), but fail to fully execute this program owing to the suppression of the inner membrane transport and mitochondrial coupling machinery (CPT2, UCP3). The reduction in PGC1α further indicates compromised mitochondrial biogenesis, while decreased DRP1 expression suggests impaired mitochondrial fission and quality control, both of which are essential for segregating and removing damaged mitochondria. Together, these changes are consistent with a state of energetically stressed, structurally compromised mitochondria that are unable to sustain efficient ATP production or maintain redox homeostasis. AAT treatment largely reversed this mitochondrial dysfunction profile, normalizing CPT1A and PDK4 expression toward control levels while restoring CPT2, UCP3, PGC1α, and DRP1. This coordinated re-adjustment implies that AAT does not merely dampen metabolic activity, but instead restores a coherent mitochondrial program, in which substrate utilization (CPT1A/PDK4), β-oxidation capacity (CPT2), bioenergetic fine-tuning and ROS limitation (UCP3), biogenesis (PGC1α), and mitochondrial dynamics/quality control (DRP1) are all brought back toward a normal physiological range. Consequently, AAT may prevent the transition from a reversible, stress-induced metabolic adaptation to irreversible organellar failure. The mitigation of mitochondrial dysfunction is particularly relevant, as mitochondrial ROS generation and energetic failure form a negative cycle that accelerates tubular injury. In this context, the restoration of an integrated network of mitochondrial enzymes and regulators likely represents an important contributor to the renoprotective effects of AAT in cisplatin-induced AKI.

Cisplatin also activates a broad inflammatory response mediated by IL-1β, chemokines, and downstream MAPK signaling, which together promote immune cell recruitment and tubular injury [1,2,3,14,39,40]. Indeed, this was reflected in our model by increased IL-1β expression and marked induction of OPN and F4/80. OPN is a stress-inducible cytokine-like protein that is minimally expressed in normal tubules but is robustly upregulated in injured epithelial cells, where the protein functions both as a tubular injury marker and as a chemoattractant that perpetuates macrophage recruitment and interstitial inflammation [41,42]. F4/80 is a well-established macrophage marker and directly reflects monocyte/macrophage infiltration into the injured kidney. Therefore, the concomitant increase in OPN and F4/80 expression indicates an active tubular–macrophage inflammatory axis in cisplatin-induced AKI. AAT treatment markedly attenuated this axis: IL-1β levels were reduced, OPN expression was suppressed, and F4/80-positive macrophage infiltration was significantly decreased. All these findings indicate that AAT dampens both tubular inflammatory activation and immune cell recruitment. These findings are consistent with previous experimental data showing that AAT exerts anti-inflammatory and cytoprotective effects in renal cells, including human proximal tubular cells [26]. AAT has been reported to attenuate proinflammatory cytokine production and oxidative stress-induced damage in tubular epithelial cells in vitro; furthermore, AAT therapy reduced inflammatory infiltrates and tubular necrosis in ischemia–reperfusion and calcineurin inhibitor nephrotoxicity models in vivo [18,19,20,21,22].

At the apoptotic level, cisplatin shifted the balance of Bcl-2 family proteins toward cell death, consistent with mitochondrial stress-driven apoptosis. AAT restored this balance by decreasing the Bax/Bcl-2 ratio, suggesting an antiapoptotic effect. The combination of restored Bax/Bcl-2 signaling and reduced MAPK phosphorylation supports the interpretation that AAT mitigates tubular cell death. This finding aligns with prior observations that AAT protects renal and peritoneal mesothelial cells from toxin- or stress-induced apoptosis [18,19,20]. Collectively, these data indicate that AAT suppresses a cisplatin-induced inflammatory and apoptotic network at multiple levels, including reducing IL-1β- and OPN-driven inflammatory amplification, limiting macrophage accumulation, and restoring prosurvival signaling in tubular epithelial cells.

Persistent inflammation, oxidative stress, mitochondrial dysfunction, and apoptotic injury converge to stimulate TGF-β1 signaling, promote EMT, and drive extracellular matrix accumulation during cisplatin-induced AKI. Importantly, fibrotic remodeling is recognized as a key pathological process linking acute tubular injury to CKD progression, rather than merely a late or passive consequence of injury [43,44]. In this context, fibrosis represents an integrative and maladaptive repair response that perpetuates structural and functional deterioration following AKI. In this study, AAT effectively mitigated fibrosis-related changes, including reduced expression of fibronectin and α-SMA, as well as decreased collagen deposition on histological staining. These antifibrotic effects should be interpreted as the cumulative outcome of the upstream actions of AAT, including suppression of oxidative stress and NOX-dependent ROS generation, stabilization of mitochondrial metabolism and biogenesis, attenuation of inflammatory signaling, and restoration of antiapoptotic balance. Thus, by simultaneously restraining these mechanistic drivers, AAT interrupts the transition from acute tubular injury to maladaptive fibrotic repair, thereby limiting the fibrogenic shift that underlies AKI-to-CKD progression. This integrative antifibrotic role is consistent with prior findings that AAT limits TGF-β1-induced EMT and extracellular matrix production in renal epithelial cells [23], reinforcing the concept that AAT may function as both an acute cytoprotective agent and a modulator of longer-term renal repair and disease progression.

From a translational perspective, AAT is an attractive therapeutic candidate due to its established clinical use and favorable safety profile in AAT deficiency [14,16]. Moreover, clinical associations linking higher circulating AAT levels to AKI severity [24,25] suggest that AAT may function as a protective factor and an endogenous stress-responsive biomarker. However, this study has several limitations. We evaluated a single cisplatin dose and time point; therefore, the long-term effects of AAT on renal recovery and chronic fibrosis remain to be determined. In addition, although the coordinated impact of AAT on oxidative stress, mitochondrial function, inflammation, and apoptosis was identified, our assessment of oxidative stress relied primarily on tissue-level oxidative damage markers and gene expression analyses rather than direct measurements of antioxidant enzyme activities. Thus, post-translational regulation of antioxidant enzymes could not be fully addressed. Furthermore, this study focused on delineating the pleiotropic protective effects of AAT across multiple pathogenic pathways, including oxidative stress, inflammation, mitochondrial dysfunction, and apoptosis, which limited the extent of in-depth mechanistic analyses within each pathway. More detailed mechanistic investigations, such as caspase activation, mitophagy, and NAD⁺/NADH metabolism, are therefore required to delineate the full underlying mechanisms. Future studies should also assess long-term functional recovery, examine the effects of AAT on chronic fibrotic outcomes, and determine whether AAT preserves the antitumor efficacy of cisplatin, which is an essential requirement for clinical translation.

5. Conclusions

In conclusion, AAT provides comprehensive protection against cisplatin-induced AKI by reducing oxidative stress, stabilizing mitochondrial metabolism, suppressing inflammatory and apoptotic pathways, and inhibiting profibrotic remodeling. Meanwhile, by targeting multiple interconnected mechanisms of cisplatin nephrotoxicity, AAT may represent a promising therapeutic agent to mitigate cisplatin-induced nephrotoxicity and reduce the burden of drug-induced kidney injury.