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. 2026 May 18;30(1):406–418. doi: 10.1080/19768354.2026.2671489

Cathepsin A deficiency exacerbates LPS-induced inflammatory liver injury and apoptosis

Hee Young Chae a, Ji Yeong Park a,b, Hyejin Hyung a, Jiwon Ko b, Su-Geun Lim b, Young Jin Lee a, Myoung Ok Kim c, Song Park d,e, Dong Kyu Choi a, Soyoung Jang e,f,CONTACT, Zae Young Ryoo a,c,✉, Soyeon Jang g,✉
PMCID: PMC13185071  PMID: 42164055

ABSTRACT

Hepatic inflammation plays a key role in acute and chronic liver diseases by driving hepatocellular injury and promoting disease progression. However, the molecular mechanisms that maintain hepatic immune homeostasis remain unclear. Cathepsin A (Ctsa), a lysosomal serine carboxypeptidase involved in protein degradation and enzyme stabilization, has been implicated in lysosomal storage disorders. However, its role in liver immunity is poorly understood. Given the emerging evidence that lysosomal proteases contribute to immune regulation and inflammatory signaling, Ctsa is a promising yet underexplored candidate for elucidating how lysosomal proteases influence hepatic inflammation. To address this gap, this study aimed to investigate Ctsa function using Ctsa knockout (Ctsa-/-) mice. Under basal conditions, Ctsa-/- mice exhibited splenic immune activation and relative hepatomegaly accompanied by histological alterations. Following lipopolysaccharide challenge, the mice developed aggravated liver injury with elevated aminotransferase levels, enhanced immune cell infiltration, and increased pro-inflammatory cytokine expression. These inflammatory changes were accompanied by increased hepatocellular apoptosis, as evidenced by elevated Bax/Bcl-2 ratio, cleaved caspase-3 expression, and increased TUNEL-positive cells. Collectively, these findings indicate that Ctsa contributes to the regulation of hepatic immune and cellular homeostasis, and its loss increases susceptibility to inflammatory liver injury.

KEYWORDS: CTSA, Cathepsin, liver inflammation, apoptosis, STAT3

GRAPHICAL ABSTRACT

graphic file with name TACS_A_2671489_UF0001_OC.jpg

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Introduction

The liver is a vital organ that coordinates nutrient storage, detoxification, and metabolic regulation. Furthermore, it plays a pivotal immunological role by producing acute-phase proteins, complement components, cytokines, and chemokines (Robinson et al. 2016). To maintain immune tolerance while defending against harmful stimuli, the liver relies on innate immune mechanisms mediated by distinct hepatic cell populations, including hepatocytes, Kupffer cells, hepatic stellate cells, and liver sinusoidal endothelial cells (Kolios et al. 2006; Gao 2016; Gong et al. 2022). In addition to innate immune mechanisms, the liver also contains diverse adaptive immune cells that participate in immune surveillance, tolerance, and chronic inflammation (Shuai et al. 2016; Zheng and Tian 2019). Disruption of this complex immune environment can lead to hepatocellular injury, fibrosis, and the progression of liver diseases, with even subtle disturbances in hepatic immune regulation potentially inducing exaggerated inflammatory responses (Heymann and Tacke 2016). Among hepatic immune cells, Kupffer cells, the resident hepatic macrophages, represent the largest population of tissue-resident macrophages and play a central role in immune surveillance and the clearance of bacteria, viruses, and other pathogens (Kolios et al. 2006). Under pathological conditions, these cells become activated and, together with infiltrating immune cells, help establish an inflammatory hepatic microenvironment that drives innate immune responses and liver injury (Krenkel and Tacke 2017).

To establish an experimental model of acute hepatic inflammation and innate immune activation, lipopolysaccharide (LPS), a prototypical component of Gram-negative bacterial cell walls, is commonly used. Systemic administration of LPS induces hepatic inflammatory responses characterized by hepatocellular injury, elevated serum aminotransferases, and the recruitment of innate immune cells to the liver (Hamesch et al. 2015; Riolo et al. 2025). This acute inflammatory response closely resembles key features of sepsis-associated liver dysfunction and endotoxin-driven hepatitis, conditions in which disruption of hepatocyte membranes results in the release of ALT and AST into circulation (Giannini et al. 2005). Additionally, LPS triggers the production of chemokines, which mediate monocyte and neutrophil recruitment and amplify hepatic inflammation. These features contribute to the widespread use of LPS as a model for studying the mechanisms underlying inflammatory liver injury (Deshmane et al. 2009; Shi and Pamer 2011).

Liver injury under inflammatory conditions is ultimately determined by the regulation of stress response pathways. Among these pathways, apoptosis plays a central role in maintaining tissue homeostasis by facilitating the removal of damaged cells through tightly regulated molecular mechanisms involving caspases and Bcl-2 family proteins, including Bcl-2, Bax, and Bak (Mustafa et al. 2024). However, excessive or dysregulated apoptosis aggravates tissue injury and promotes disease progression (Korsmeyer 1999; Guicciardi and Gores 2005). In the liver, apoptosis is not merely a terminal consequence of inflammation but an active driver of disease progression. Apoptotic hepatocytes release damage-associated molecular patterns (DAMPs) and apoptotic bodies that stimulate Kupffer cells and infiltrate immune cells, thereby reinforcing inflammatory cytokine production and sustaining hepatic inflammation (Canbay et al. 2003; Gong et al. 2022).

Signal transducer and activator of transcription 3 (STAT3) is a key transcription factor that integrates inflammatory and stress-related signals to regulate hepatocellular homeostasis. Upon IL-6 activation, STAT3 promotes acute-phase protein synthesis, limits excessive tissue damage, and supports hepatocyte survival (Taub 2003; Taub 2004). Under basal conditions, this pathway helps maintain metabolic and immunological stability. Transient STAT3 activation is generally protective; however, sustained or dysregulated signaling can amplify inflammation, increase cellular stress and impair regenerative processes. Aberrant STAT3 activation has been associated with heightened susceptibility to endotoxin-induced liver injury, the impaired resolution of inflammation, and disruption of hepatocyte integrity (Gao et al. 2012; Kasembeli et al. 2018).

Cathepsins are a family of lysosomal proteases classified into serine, aspartic, and cysteine subgroups. Cathepsin A (Ctsa), a lysosomal serine protease, degrades excess substrates and forms a protective complex with β-galactosidase and neuraminidase (NEU1), contributing to their stabilization and catalytic activity (Hiraiwa 1999). Although Ctsa has traditionally been studied in the context of lysosomal homeostasis and tumor progression, emerging evidence indicates broader functions in immune regulation (Masuhara et al. 2009; Park and Lee 2025). However, its involvement in inflammatory liver injury remains poorly understood.

Given the emerging associations between lysosomal proteases, inflammatory signaling, and cell death regulation, we hypothesized that Ctsa plays a critical role in maintaining hepatic immune and cellular homeostasis under inflammatory stress. In this study, we investigated the function of Ctsa in LPS-induced acute inflammatory liver injury using Ctsa-deficient mice. The results revealed that loss of Ctsa exacerbates hepatocellular damage and inflammatory responses, accompanied by dysregulated apoptosis and altered STAT3 signaling. These findings suggest that Ctsa contributes to the maintenance of hepatic immune homeostasis and cell survival pathways under inflammatory stress conditions.

Materials and methods

Animals

Wild type (WT) and Ctsa knockout (Ctsa-/-) mice on a C57BL/6J background, aged 8–10 weeks, were used in the experiments. Both male and female mice were included in this study (WT: 12 males and 10 females; Ctsa-/-: 13 males and 11 females). Ctsa-/- mice were generated by targeting exon 2 of the Ctsa gene using the CRISPR/Cas9 system. The guide RNA sequences were as follows: sgRNA1 (5’- GGCCCGGAGTTGGATCCGAGCGG-3’) and sgRNA2 (5’- GAATTCTAACGACCCCAGGCAGG-3’). Genotyping was performed by PCR. The PCR products were separated through electrophoresis on a 1.5% agarose gel and distinguished by size: 668 bp for WT and 277 bp for Ctsa-/-. PCR amplification was performed using Ctsa primers (F: 5’- CAC TGC TCT TGT TGC TGC TC-3’, R: 5’-GTC CGA TGC TCT GAG GTA GC-3’). LPS-induced liver injury was established by intraperitoneal injection of 5 mg/kg LPS (L2630, Sigma-Aldrich, Inc., St. Louis, USA). An equal volume of saline was injected into the control group. Mice were euthanized 12 h after injection via intraperitoneal administration of avertin (300 mg/kg), and tissue samples were harvested for further analysis. All mice were maintained under a 12 h light/12 h dark with free access to food and water. All animal experiments adhered to the guidelines for animal experimentation of the Kyungpook National University Animal Care and Use Committee and were approved by the Institutional Ethics Committee (Daegu, Korea; approval number 2024-0096). All experiments were performed in a manner consistent with the ARRIVE guidelines (https://arriveguidelines.org).

Western blot analysis

Total protein was extracted from liver tissue using PRO-PREP lysis buffer (iNtRON Biotechnology, Korea) supplemented with phosphatase inhibitor cocktail (GenDEPOT, TX, USA). Proteins were separated by SDS-PAGE on a 10% or 15% polyacrylamide gel and then subsequently transferred onto PVDF membranes (Immobilon®-P PVDF Membrane, IPVH08100, 0.45 μm). Membranes were blocked with 5% non-fat dry milk or 5% bovine serum albumin for 1 h and incubated overnight with primary antibodies at 4°C. Following this, membranes were washed and incubated with horseradish peroxidase (HRP)-conjugated secondary antibody for 2 h at room temperature. Protein signals were detected using West-Q Pico ECL solution (GenDEPOT, W3652-020). The primary antibodies were used: CTSA (1:2000, Cell Signaling Technology, Danvers, MA, USA, 88193), cleaved caspase 3 (1:2000, Cell Signaling Technology, 9664), Bax (1:2000, Santa Cruz Biotechnology, Dallas, TX, USA, sc-7480), Bcl-2 (1:2000, Santa Cruz Biotechnology, sc-509), STAT3 (1:2000, Cell Signaling Technology, 4904), p-STAT3 (1:2000, Cell Signaling Technology, 9145), β-actin (1:5000, Santa Cruz Biotechnology, sc-47778) and GAPDH (1:5000, Cell Signaling Technology, 2118). Goat anti-mouse IgG (H + L; 1:5000, Invitrogen, 31430) and goat anti-rabbit IgG (H + L; 1:5000, Invitrogen, 31463) were used as HRP-conjugated secondary antibodies. Protein levels were normalized to β-actin or GAPDH and analyzed using ImageJ software.

Quantitative real-time PCR

Total RNA was isolated from spleen and liver tissue using the QIAzol lysis reagent (QIAGEN). RNA was reverse transcribed into cDNA using the PrimeScript 1st strand cDNA synthesis kit (TAKARA). Quantitative real-time PCR was performed using TB Green® Premix Ex Taq™ II (Tli RNaseH Plus) (TAKARA) on a LightCycler 96 system. The following gene-specific primers were used: MCP-1 (F: 5’- GCA GCA GGT GTC CCA AAG AA −3’, R: 5’- ATT TAC GGG TCA ACT TCA CAT TCA −3’), CXCL1 (F: 5’- CTG GGA TTC ACC TCA AGA ACA TC −3’, R: 5’- CAG GGT CAA GGC AAG CCT C −3’), TNF-α (F: 5’- TGT GCT CAG AGC TTT CAA CAA C −3’, R: 5’- GCC CAT TTG AGT CCT TGA TG −3’), IL-6 (F: 5’- TGT CTA TAC CAC TTC ACA AGT CGG AG −3’, R: 5’- GCA CAA CTC TTT TCT CAT TTC CAC −3’), IL-1β (F: 5’- TGC CAC CTT TTG ACA GTG ATG C-3’, R: 5’- TGA TGT GCT GCT GCG AGA TT −3’), IL-10 (F: 5’- TTT GAA TTC CCT GGG TGA GAA −3’, R: 5’- TGC TCC ACT GCC TTG CTC TT −3’) and β-actin (F: 5’- TCT GGC ACC ACA CCT TCT ACA −3’, R: 5’- TTT TCA CGG TTG GCC TTA GG −3’). Relative mRNA levels were calculated using the 2−ΔΔCt method and normalized to the level of β-actin.

Flow cytometry

Mouse spleens were mechanically dissociated through 70-μm cell strainers using the plunger of a syringe to obtain single-cell suspensions. Red blood cells were removed using 1×RBC lysis buffer prepared from 10× RBC lysis buffer stock (BioLegend, 420302). Cells were fixed with 4% paraformaldehyde and permeabilized with 90% methanol in PBS for 15 min. Fc receptors were blocked with anti-CD16/32 antibody (BioLegend, 156604) for 10 min, cells were stained with FITC-anti-CD4 (Invitrogen, 11-0042-82), PE-anti-IL-17A (BD Pharmingen™, 559502), and PE-anti-Foxp3 (eBioscience™, 12-4771-82, San Diego, California) antibodies. Stained cells were analyzed using FACS Aria Fusion (BD Biosciences, CA, USA).

Hematoxylin and eosin (H&E) staining

Liver tissues were fixed in 4% paraformaldehyde, embedded in paraffin and sliced into 4-μm sections. The sections were deparaffinized, rehydrated, stained with hematoxylin (StatLab, Clearview Hematoxylin) and eosin (StatLab, Clearview eosin). Slides were observed using a Motic EasyScan One (Motic Asia, Kowloon, Hong Kong).

Immunohistochemistry

Liver tissues were embedded in paraffin, sliced into 4-μm sections, deparaffinized, and then rehydrated. The sections were subsequently treated with 3% H₂O₂ for 20 min at room temperature to block endogenous peroxidase activity. Heat-induced antigen retrieval was performed using boiling citrate buffer. Following a 5 min wash in phosphate-buffered saline (PBS), sections were blocked with 10% normal goat serum for 1 h at room temperature and incubated overnight at 4 °C with primary antibody. Immunohistochemistry staining was performed using anti-CD68 (1:400, Cell Signaling Technology, 97778) and Ly6G (1:100, Abcam, ab25377, Cambridge, UK) antibodies. After three washes (5 min each) with PBS containing 0.1% Triton X-100 (PBST), sections were incubated with an anti-rabbit secondary antibody (VECTASTAIN ABC kit, Vector Labs, CA, USA) for 30 min at room temperature. Signals were detected using 3,3’-diaminobenzidine tetrahydrochloride-dihydrate peroxidase substrate (Vector Labs) staining. Nuclei were stained for 30 s using a 1:4 dilution of hematoxylin. Slides were observed using a Motic EasyScan One (Motic Asia).

Immunofluorescence

After permeabilization in PBST for 10 min, the sections were washed three times with PBS for 5 min each and blocked with 1% BSA at room temperature for 1 h. The sections were then incubated overnight at 4°C with the primary antibody CD11b (1:100, BioLegend, San Diego, USA, 101205). Following an additional three washes (5 min each) with PBS, the nuclei were stained using ProLong™ Gold Antifade mountant with DNA Stain DAPI (Invitrogen, P36931). Slides were observed using a fluorescence microscope (Leica DMI3000B, Leica, Germany), and cells were quantified using ImageJ.

TUNEL assay

Apoptotic cell death in liver tissues was evaluated using a TUNEL assay with the In Situ Cell Death Detection Kit (11684795910, Roche Diagnostics, Mannheim, Germany) according to the manufacturer’s protocol. Tissue sections were deparaffinized, rehydrated, and permeabilized prior to TUNEL labeling. The sections were then incubated with the TUNEL reaction mixture at 37 °C for 60 min in a humidified chamber. After washing with PBS, nuclei were counterstained with DAPI using ProLong™ Gold Antifade mountant with DNA Stain DAPI (Invitrogen, P36931), and TUNEL-positive cells were visualized using a fluorescence microscope.

ELISA

Serum was isolated from the blood of WT and Ctsa-/- mice 12 h after intraperitoneal injection of LPS and prior to euthanization. The serum was stored at −80°C until further analysis. Levels of the pro-inflammatory cytokines TNF-α (Invitrogen, 88-7324), IL-6 (Invitrogen, 88-7064), and IL-1β (Invitrogen, 88-7013A) were measured using commercial kits according to the manufacturer's instructions. Cytokine concentrations were measured using standard curve based on the absorbance values at 450 nm. The concentrations of each sample were calculated according to the standard curve.

Blood biochemical test

Serum ALT and AST levels in WT and Ctsa-/- mice were measured at the Preclinical Center of the Daegu Gyeongbuk Advanced Medical Industry Promotion Foundation.

Statistical analysis

Data were analyzed using GraphPad Prism (version 10.2) and are presented as mean ± standard deviation (SD). Comparisons between two groups were performed using a Student’s t-test, and comparisons among multiple groups were conducted using two-way analysis of variance (ANOVA). Significance levels are reported as *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Results

Generation of Ctsa knockout mice and splenic immune activation with altered Th17 populations

Transgenic Ctsa-/- mice were generated by targeting exon 2 of the Ctsa gene using the CRISPR-Cas9 system (Figure 1A). Deletion of Ctsa was confirmed by genotyping polymerase chain reaction (PCR), which distinguished WT, heterozygous, and homozygous knockout alleles based on the expected fragment sizes (Figure 1B). The absence of Ctsa protein expressions in the spleens of Ctsa-/- mice was confirmed by western blot analysis based on a comparison with WT mice (Figure 1C). Spleen weight did not differ significantly between the two groups; however, the spleen-to-body weight ratio was significantly higher in Ctsa-/- mice (Figure 1D). An increased spleen-to-body weight ratio is commonly associated with altered splenic homeostasis and chronic immune activation. To determine whether Ctsa deficiency is associated with basal immune activation and dysregulated inflammatory signaling in the spleen, the expression levels of the splenic inflammation-associated cytokines such as tumor necrosis factor (TNF)-α, interleukin (IL)−6, IL-1β, and IL-10 were assessed using quantitative reverse transcription real-time polymerase chain reaction (qRT)-PCR. Consistent with the increased spleen-to-body weight ratio, splenic analysis showed a significant upregulation of IL-1β, IL-6, and IL-10 transcripts in Ctsa-/- mice relative to the WT controls (Figure 1E). To further evaluate immune cell populations in the spleen, splenocytes were isolated and subjected to flow cytometric analysis (Park and Lee 2025). Similarly, analysis of the distributions of immune cell populations within splenocytes showed significantly larger Th17 populations in Ctsa-/- mice than in WT mice, whereas regulatory T cell (Treg) populations did not significantly differ (Figure 1F). This selective increase in Th17 cells indicated a shift in the Th17/Treg balance toward a proinflammatory immune axis, as is commonly observed in the early stages of autoimmune and chronic inflammatory diseases. Collectively, these findings demonstrate that Ctsa deficiency induces a state of systemic immune activation and proinflammatory priming even under unstimulated conditions.

Figure 1.

Six visuals: one gene editing diagram, one gel image, one western blot, nine bar charts, and one flow cytometry plot comparing WT and Ctsa knockout mice. The figure shows six visuals summarizing generation and analysis of cathepsin A deficient mice. A schematic diagram depicts the Ctsa gene with exon 1, exon 2, and two guide ribonucleic acid target sites flanking exon 2, with short nucleotide sequences labeled beneath. A gel image labeled wild type, Ctsa + or minus, and Ctsa minus minus shows a larger band near 600 base pairs for the wild type allele and a smaller band near 300 base pairs for the knockout allele. A western blot image compares spleen lysates from wild type and Ctsa minus minus mice, with upper bands labeled cathepsin A around 54 kilodaltons and lower bands labeled beta actin around 43 kilodaltons; cathepsin A bands are present in wild type lanes and absent in knockout lanes, while beta actin bands appear in all lanes. Three grouped bar charts display body weight, spleen weight, and spleen to body weight ratio for wild type and Ctsa minus minus mice with individual data points, error bars, and Student t test * marks; all data are approximate. A set of four bar charts shows relative messenger ribonucleic acid levels of tumor necrosis factor alpha, interleukin 1 beta, interleukin 6, and interleukin 10 in wild type and Ctsa minus minus spleens with error bars and *s; all data are approximate. Flow cytometry dot plots of interleukin 17A versus CD4 and Foxp3 versus CD4 for wild type and Ctsa minus minus splenocytes appear on the left, and two bar charts on the right plot percentages of T helper 17 and regulatory T cell populations for each genotype with error bars and *s; all data are approximate.

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Generation of Ctsa knockout mice and splenic immune activation in Ctsa-deficient mice. (A) Ctsa-/- mice were generated using the CRISPR/Cas9 system. (B) Genotyping PCR of WT, heterozygous (Ctsa+/–), and CTSA-deficient (Ctsa–/–) mice was performed using allele specific primers. The WT allele produced a ∼600 bp band, whereas the KO allele yielded a ∼300 bp fragment. (C) Ctsa expression levels in spleen tissue were analyzed by western blot to confirm the successful deletion of Ctsa in Ctsa-/- mice. (D) Body weight and spleen weight were measured in WT and Ctsa-/- mice to assess physiological changes associated with Ctsa deletion. Spleen weight was normalized to body weight for comparative analysis (n = 4 per group). (E) Relative TNF-α, IL-1β, IL-6 and IL-10 mRNA levels were measured by qRT-PCR to assess inflammatory responses in the mouse spleen (n = 4 per group). (F) Th17 and Treg cell populations in splenocytes from WT and Ctsa-/- mice were measured using flow cytometry (n = 5 per group). Data are expressed as the mean ± SD. Statistical analysis was performed using Student’s t-test. *p < 0.05, **p < 0.01, and ***p < 0.001.

Hepatic morphological alterations in Ctsa-deficient mice

Given the presence of splenic immune activation, we investigated whether Ctsa deficiency was associated with alterations in liver morphology and basal hepatic inflammatory status. Although absolute liver weight did not differ significantly between groups, the liver-to-body weight ratio was significantly increased in Ctsa-/- mice, indicating relative hepatomegaly (Figure 2A). Liver enlargement is associated with compensatory responses to hepatic tissue stress and impaired liver function (Ludwig et al. 1980; Sharma and Arora 2020). H&E staining, performed to examine histological alterations associated with Ctsa deficiency, revealed inflammatory cell infiltration and inflammatory foci in the livers of Ctsa-/- mice (Figure 2B). Despite these hepatic structural abnormalities, hepatic expression of pro-inflammatory cytokines, including TNF-α, IL-1β, and IL-6, was not significantly increased under basal conditions (Figure 2C). These results indicate that, although Ctsa deficiency leads to systemic immune activation, the liver exhibits early morphological abnormalities and immune cell infiltration without overt induction of inflammatory cytokines expression at baseline.

Figure 2.

Five bar charts and four microscopy images comparing WT and Ctsa deficient mouse livers, with higher liver ratio and scattered infiltrates. The figure shows quantitative charts and microscopy images comparing livers from wild type mice and Ctsa deficient mice. The upper left chart is a bar chart labeled Liver weight in grams on the vertical axis from 0.0 to 1.5 in 0.5 increments and wild type and Ctsa deficient on the horizontal axis. Both groups have similar bars around 0.9 to 1.1 with scatter points and error bars. The upper middle chart is a bar chart labeled Liver to body weight in percent on the vertical axis from 0 to 8 in steps of 2 and wild type and Ctsa deficient on the horizontal axis. The Ctsa deficient bar reaches about 7 while the wild type bar is around 4 with individual points and a line of *s above the groups. On the lower left, three vertical bar charts show relative messenger ribonucleic acid levels in liver for tumor necrosis factor alpha, interleukin 1 beta, and interleukin 6. Each chart has wild type and Ctsa deficient categories on the horizontal axis and relative messenger ribonucleic acid level on the vertical axis, with values below about 5 for tumor necrosis factor alpha, below about 2.5 for interleukin 1 beta, and below about 25 for interleukin 6. Bars include error bars and scattered points. The right side displays four hematoxylin and eosin stained liver microscopy images: an overview and a magnified view for wild type and for Ctsa deficient tissue. The Ctsa deficient higher magnification image contains a clustered focus of small cells indicated by an arrow, whereas the wild type image shows a more uniform parenchyma. All data are approximate.

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Histological and physiological analyses in Ctsa-deficient mice. (A) Liver weight was measured in WT and Ctsa-/- mice to assess the physiological changes associated with Ctsa deletion. Liver weight was normalized to body weight for comparative analysis (n = 4 per group). (B) Mouse liver tissue sections were stained with H&E to evaluate histopathological changes. Scale bars: 200 and 120 μm (n = 3 per group). (C) Relative TNF-α, IL-1β and IL-6 mRNA levels in mouse liver were measured by qRT-PCR (n = 4 per group). Data are expressed as the mean ± SD. Statistical analysis was performed using Student’s t-test. ****p < 0.0001.

Ctsa deficiency aggravates the LPS-induced inflammatory response

Based on the relative hepatomegaly, hepatic structural alterations, and the increased levels of pro-inflammatory cytokines in the spleen of Ctsa-/- mice, we examined whether Ctsa deficiency exacerbates liver injury in response to an inflammatory challenge.

LPS treatment induced a significantly greater increase in CXCL1 levels in Ctsa-/- mice compared with saline-treated controls. Furthermore, under LPS-induced hepatic inflammatory conditions, the livers of Ctsa-deficient mice exhibited markedly higher expression of pro-inflammatory cytokines (TNF-α, IL-1β, and IL-6) and chemokine (MCP-1) compared with WT mice (Figure 3A). Histological analysis of H&E-stained liver tissue sections revealed considerably greater inflammatory cell infiltration and focal inflammatory cell aggregation in Ctsa-/- mice following LPS administration (Figure 3B). Given the pronounced histological signs of inflammatory cell infiltration, and the increased inflammatory cytokines and chemokines known to promote liver injury and immune cell recruitment, we further examined immune cell infiltration in the liver. Hepatic macrophage accumulation was assessed using CD68 immunohistochemistry, and inflammatory myeloid cell infiltration was evaluated by CD11b immunofluorescence. Under inflammatory challenge, Ctsa-/- mice exhibited a higher number of infiltrating immune cells compared to WT controls (Figure 3C–E). Neutrophil infiltration in the liver was examined by immunohistochemical staining for LY6G. Following LPS administration, the number of infiltrating neutrophils was higher in Ctsa-/- mice than in WT mice (Figure 3F). These results demonstrate that, under LPS-induced inflammatory conditions, Ctsa deficiency enhances the recruitment of innate immune cells and promotes the hepatic inflammatory response.

Figure 3.

Six visuals: six bar charts and four set of microscopy images comparing LPS and saline effects on WT and Ctsa knockout mouse livers. The figure shows six visuals comparing wild type and Ctsa knockout mouse livers after saline or lipopolysaccharide treatment. The first visual is a group of five bar charts for interleukin 1 beta, tumor necrosis factor alpha, interleukin 6, monocyte chemoattractant protein 1, and chemokine C X C motif ligand 1 messenger ribonucleic acid levels. The x axis of each chart lists saline and lipopolysaccharide for wild type and Ctsa knockout, and the y axis shows relative messenger ribonucleic acid level from 0 to about 120 with ticks every 20 units. Bars for lipopolysaccharide treated Ctsa knockout mice are higher than the other groups. All data are approximate. The second visual is a set of four hematoxylin and eosin stained liver microscopy images arranged by genotype and treatment, with arrows indicating immune cell clusters. The third visual shows four immunohistochemistry liver images stained for macrophage marker C D 68. The fourth visual presents four immunofluorescence liver images stained with nuclear dye and C D 11 b, alongside merged views. The fifth visual is a single bar chart of C D 11 b positive area, showing the highest bar for lipopolysaccharide treated Ctsa knockout liver. All data are approximate. The sixth visual shows four liver immunohistochemistry images stained for neutrophil marker L y 6 G.

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Ctsa deficiency exacerbates LPS-induced hepatic inflammation and promotes innate immune cell infiltration. WT and Ctsa-/- mice were intraperitoneally injected with LPS (5 mg/kg) and euthanized 12 h later. (A) Relative mRNA expression levels of cytokines (TNF-α, IL-1β, IL-6) and chemokines (MCP-1, CXCL-1) in mouse liver tissue were measured by qRT-PCR (n≥5 per group). (B) H&E-stained liver tissue sections were evaluated for histopathological changes, including immune cell infiltration and morphological alterations. Scale bar: 200 μm (n = 3 per group). (C) Macrophage infiltration in mouse liver tissue was assessed immunohistochemically using CD68, a macrophage marker. Scale bar: 100 μm (n = 3 per group). (D, E) Immunofluorescence staining of mouse liver tissue was performed to detect CD11b expression, indicating infiltrating myeloid cells. Scale bar: 50 μm (n = 3 per group). (F) Neutrophil infiltration in mouse liver tissue was evaluated by immunohistochemical staining for Ly6G, a neutrophil marker. Scale bar: 300 μm (n = 3 per group). Data are presented as the mean ± SD. Statistical significance was determined using Student’s t-test or two-way ANOVA. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001.

Ctsa deficiency exacerbates apoptotic cell death in the liver

Serum levels of TNF-α, IL-1β, and IL-6 were analyzed to evaluate the development and progression of acute inflammatory liver injury and changes in systemic inflammatory responses (Figure 4A). These findings indicate that Ctsa deficiency is associated with heightened systemic and hepatic inflammatory responses and increased liver injury following LPS challenge. Consistent with these observations, serum ALT and AST levels were significantly elevated in LPS-challenged Ctsa-/- mice, indicating exacerbated hepatocellular injury (Figure 4B). To determine whether the aggravated liver injury in Ctsa-deficient livers was accompanied by increased apoptotic signaling, we next examined the expression of Bcl-2 family proteins and markers of caspase activation. Western blot analysis of Bax and Bcl-2 confirmed these findings at the protein level, with a significantly elevated Bax/Bcl-2 ratio in Ctsa-deficient mice, suggesting that apoptosis is promoted under LPS-induced inflammation conditions. Executioner and initiator caspases, as downstream effectors of apoptosis, were also examined (Ludwig et al. 1980). Western blot analysis revealed increased levels of cleaved caspase-3, a key executioner caspase. The accumulation of cleaved caspase-3 indicates active apoptotic cell death in Ctsa-deficient livers. Quantitative analysis showed markedly elevated levels of activated apoptosis proteins in Ctsa-/- mice compared to WT controls (Figure 4C, D). To confirm hepatocellular apoptosis in vivo, we performed TUNEL staining on liver tissue sections. TUNEL positive cells were increased in the livers of Ctsa-/- mice following LPS challenge compared with WT controls (Figure 4E). These results indicate that Ctsa deficiency exacerbates hepatocyte apoptosis during LPS-induced liver injury.

Figure 4.

Five visuals: seven bar charts, one western blot image, one TUNEL microscopy panel comparing WT and Ctsa knockout mouse livers after LPS. The figure shows experimental data comparing wild type and Ctsa knockout mouse livers after lipopolysaccharide challenge. The first visual is a set of three grouped bar charts with treatment on the horizontal axis and cytokine concentration on the vertical axis in picograms per milliliter from 0 to 1500 or 0 to 50000 at 250 or 10000 intervals. Bars for Ctsa knockout with lipopolysaccharide are higher than wild type with lipopolysaccharide for tumor necrosis factor alpha, interleukin 1 beta, and interleukin 6. The second visual is two grouped bar charts of serum alanine aminotransferase and aspartate aminotransferase, with activity units on the vertical axis from 0 to about 200 at 50 unit intervals, showing higher values in lipopolysaccharide treated Ctsa knockout mice. The third visual is a western blot image with four horizontal bands labeled B cell lymphoma 2, Bax, cleaved caspase 3, and beta actin across wild type saline, wild type lipopolysaccharide, Ctsa knockout saline, and Ctsa knockout lipopolysaccharide lanes. The fourth visual is two bar charts plotting Bax to B cell lymphoma 2 ratio and cleaved caspase 3 to beta actin ratio, both higher with lipopolysaccharide in Ctsa knockout mice. The fifth visual is a TUNEL staining microscopy grid showing more TUNEL positive nuclei in Ctsa knockout lipopolysaccharide liver sections than in other groups. All data are approximate.

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Ctsa deletion exacerbates apoptosis and liver injury following LPS challenge. (A) TNF-α, IL-1β, and IL-6 levels in mouse serum were measured by ELISA (n ≥ 4 per group). (B) Serum levels of ALT and AST in LPS-treated mice were measured to assess liver function (n ≥ 4 per group). (C, D) Liver lysates from WT and Ctsa-/- mice were analyzed by western blot to assess c-caspase-3, Bax, and Bcl-2 protein levels, as markers of apoptosis (n ≥ 5 per group). (E) TUNEL positive cells were measured in liver tissue. Scale bar: 100 μm (n = 3 per group). Data are presented as the mean ± SD. Statistical significance was determined using two-way ANOVA. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001.

Ctsa deficiency is associated with altered hepatic STAT3 signaling following LPS challenge

Because IL-6 is a major upstream activator of STAT3 signaling, we examined the activation of the STAT3 pathway through western blot analysis. Western blot analysis showed increased STAT3 phosphorylation following LPS challenge, which appeared more prominent in Ctsa-deficient mice. However, total STAT3 protein levels were reduced in Ctsa−/− livers under both basal and inflammatory conditions. Consequently, the relative phosphorylation ratio (p-STAT3/STAT3) was elevated in Ctsa-deficient mice after LPS administration (Figure 5A, B). These findings indicate that CTSA deficiency is associated with altered STAT3 signaling in the liver during inflammatory stress.

Figure 5.

Two visuals, a western blot and a bar chart, comparing liver STAT3 phosphorylation in WT and Ctsa deficient mice with saline or LPS. The figure shows 2 visuals summarizing liver signal transducer and activator of transcription 3, or STAT3, analysis in wild type and Ctsa deficient mice treated with saline or lipopolysaccharide. The left visual is a western blot labeled A. Three horizontal bands are shown for phospho STAT3, total STAT3, and beta actin. Above the blot, 4 treatment groups are labeled from left to right as wild type saline, wild type lipopolysaccharide, Ctsa knock out saline, and Ctsa knock out lipopolysaccharide. For phospho STAT3, bands are faint in saline lanes and darker in lipopolysaccharide lanes, with the strongest bands in the Ctsa knock out lipopolysaccharide group. For total STAT3, bands appear relatively uniform in wild type lanes and lighter in Ctsa knock out lanes. Beta actin bands look similar across lanes. Molecular weight markers on the right indicate 86 kilodalton and 79 kilodalton for phospho STAT3 and STAT3, and 43 kilodalton for beta actin. The right visual is a bar chart labeled B titled phospho STAT3 divided by STAT3. The x axis lists 4 groups, wild type saline, wild type lipopolysaccharide, Ctsa knock out saline, and Ctsa knock out lipopolysaccharide. The y axis is labeled phospho STAT3 divided by STAT3 and ranges from 0 to 3 with tick marks at 1 and 2. Each bar has 5 overlaid dots and an error bar. Bars for saline groups are near 0. Bars for lipopolysaccharide groups are higher, with the tallest bar in the Ctsa knock out lipopolysaccharide group. Three asterisks above this group indicate a reported p value less than 0.001. All data are approximate.

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Ctsa deficiency alters hepatic STAT3 phosphorylation following LPS challenge. (A, B) Liver lysates from WT and Ctsa-/- mice were analyzed by western blot to examine the STAT3 pathway (n≥5 per group). Data are presented as the mean ± SD. Statistical significance was determined using two-way ANOVA. ***p < 0.001.

Discussion

Maintaining hepatic immune homeostasis is essential for preventing excessive inflammatory responses during systemic stress. In this study Ctsa was identified as a critical regulator of hepatic immune balance under inflammatory stress. Although previous studies on Ctsa, a lysosomal serine carboxypeptidase involved in enzyme stabilization and lysosomal substrate degradation, have primarily focused on lysosomal storage disorders and tumor biology, this study identified a previously unrecognized immunoregulatory role in the liver (Hiraiwa 1999; Caciotti et al. 2013; Wang et al. 2021). Specifically, our results demonstrate that Ctsa is involved in the regulation of inflammatory signaling, altered immune cell populations, and apoptosis. These findings highlight a previously underappreciated role for lysosomal CTSA in coordinating immune and stress-response pathways during inflammatory challenges.

Ctsa deficiency induced significant phenotypic and immunological changes, including relative hepatomegaly and hepatic structural alterations, without resulting in an overt elevation of hepatic cytokines. This is consistent with the intrinsically tolerogenic immune environment of the liver (Zheng and Tian 2019). These hepatic structural alterations were accompanied by an increase in cytokine expression in the spleen and a higher proportion of Th17 cells in splenocytes, indicating that the spleen serves as a central organ for immune cell activation and systemic cytokine regulation. Thus, under basal conditions, Ctsa deficiency may establish a pre-activated systemic immune state despite relatively quiescent hepatic cytokine levels (Heymann and Tacke 2016; Krenkel and Tacke 2017). Previous studies have shown that a Th17/Treg imbalance is strongly associated with autoimmunity and chronic inflammation (Miossec and Kolls 2012; Lee 2018; Lee et al. 2026). Given the liver’s role in maintaining immune tolerance to continuous antigenic exposure, the splenic upregulation observed in this study suggests that loss of CTSA disrupts the hepatic–splenic immune axis (Jenne and Kubes 2013). The resulting imbalance may create a permissive immunological state that sensitizes the host to subsequent inflammatory challenges (Racanelli and Rehermann 2006).

Upon LPS challenge, Ctsa-deficient mice exhibited exacerbated liver injury, characterized by elevated aminotransferase levels, enhanced immune cell infiltration, and the upregulation of inflammatory mediators. The heightened observed response in the absence of Ctsa suggests that basal immune priming lowers the threshold for inflammatory activation, thereby amplifying tissue injury in Ctsa-/- mice. Collectively, these findings suggest that Ctsa may function as an important modulator of inflammatory signaling and cellular stress responses, contributing to the maintenance of hepatic homeostasis under inflammatory conditions.

Alterations in STAT3 signaling were observed in Ctsa-deficient livers following the LPS challenge. STAT3 is widely recognized as a hepatoprotective transcription factor that promotes cell survival and limits inflammatory injury (Taub 2003; Taub 2004; Gao et al. 2012; Kasembeli et al. 2018). In the present study, Ctsa-deficient livers exhibited substantially elevated IL-6 levels and increased STAT3 phosphorylation following LPS administration. Notably, this increase in STAT3 phosphorylation occurred alongside a reduction in total STAT3 protein levels. Therefore, the elevated p-STAT3/STAT3 ratio observed in Ctsa-deficient livers should be interpreted with caution, as it may reflect both a relative enrichment of phosphorylated STAT3 and a reduction in total STAT3 abundance rather than a definitive increase in overall STAT3 signaling activity.

Under these conditions, liver damage and apoptotic signaling were significantly exacerbated, as evidenced by an increased Bax/Bcl-2 ratio, higher cleaved caspase-3 expression, and increased TUNEL-positive cells. Although changes in STAT3 phosphorylation were observed in Ctsa-deficient livers, the present findings do not establish a direct causal relationship between STAT3 signaling and hepatocyte apoptosis. Instead, the aggravated liver injury observed in Ctsa-deficient mice was accompanied by increased mitochondrial apoptotic signaling, suggesting that hepatocytes may be more vulnerable to inflammatory injury under CTSA-deficient conditions. In this context, the observed STAT3 activation may represent a compensatory response to heightened inflammatory stress. Taken together, these results suggest that Ctsa deficiency may increase susceptibility to hepatocellular injury under inflammatory conditions, while the observed changes in STAT3 phosphorylation likely reflect alterations in inflammatory signaling. Further studies investigating the relationship between STAT3 signaling and hepatocyte survival will be required to better understand the precise role of STAT3 in the context of Ctsa deficiency.

Although this study was based on an acute LPS-induced liver injury model, the use of global Ctsa knockout mice limits the ability to distinguish liver-intrinsic effects of Ctsa deficiency from those driven by systemic immune dysregulation. Notably, the immune alterations observed in the spleen, including an increased proportion of Th17 cells, suggest that systemic immune activation may partially contribute to the hepatic inflammatory phenotype. Therefore, future studies using liver-specific Ctsa knockout models will be essential to determine whether Ctsa acts locally within hepatic parenchymal cells or resident immune cells to regulate inflammatory signaling and hepatocyte survival. Additionally, although the present study demonstrates enhanced hepatic inflammation in Ctsa-deficient mice, it does not define the specific contribution of livers-resident macrophages to this phenotype. Future studies using isolated Kupffer cells from WT and Ctsa knockout mice will be necessary to determine whether Ctsa deficiency directly alters macrophage inflammatory response to LPS. Furthermore, extending these observations to models of chronic liver injury and fibrosis may provide valuable insight into whether Ctsa deficiency promotes persistent inflammation, fibrotic progression, and/or parenchymal remodeling. Such studies would help determine whether the loss of Ctsa leads to sustained immune cell activation, prolonged cytokine production, and long-term Th17/Treg imbalance, all of which are well-established drivers of chronic inflammation and fibrotic progression (Sun et al. 2014; He et al. 2017). To establish the clinical relevance of Ctsa in liver pathology, studies examining Ctsa expression and enzymatic activity in well-characterized patient cohorts, including those with autoimmune hepatitis, nonalcoholic steatohepatitis, and sepsis-associated liver injury will be important. Correlating Ctsa activity with disease stage and clinical prognosis may help elucidate whether the dysregulation observed in experimental mouse models mimics human disease.

In summary, our study suggests that Ctsa may function as a previously unrecognized regulator of hepatic immune homeostasis, integrating inflammatory signaling and apoptotic regulation during inflammatory stress. By demonstrating that Ctsa deficiency exacerbates liver injury enhanced hepatocellular apoptosis, accompanied by enhanced inflammatory responses and changes in STAT3 phosphorylation. These findings establish a mechanistic link between lysosomal protease function and inflammatory liver pathology. These results also highlight Ctsa as an unrecognized potential regulator and biomarker candidate for inflammatory liver diseases.

Author contributions

HYC, JYP, and HH performed the experiments and conducted data analysis. JK, SGL, and YJL assisted with data analysis. MOK, SP, and DKC critically reviewed the manuscript. SJ (Soyoung Jang), ZYR, and SJ (Soyeon Jang) participated in study design and coordination. HYC and JYP drafted the manuscript. All authors read and approved the final version of the manuscript.

Funding Statement

This work was supported by the National Research Foundation of Korea (NRF), funded by the Korean government (MSIT) (Grant Nos. RS-2025-23963601, RS-2024-00462878, RS-2024-00452147, RS-2024-00412716 and 2022R1C1C2005834).

Disclosure statement

No potential conflict of interest was reported by the author(s).

Ethics approval and consent to participate

All animal experiments were conducted in accordance with the guidelines and rules of the Institutional Animal Care and Use Committee of Kyungpook National University. The experimental protocol was approved by the same committee (Approval Number: KNU2024-0096).

Availability of data and material

All data generated and analyzed in this study are included in this published article.

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Data Availability Statement

All data generated and analyzed in this study are included in this published article.

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