Oxaliplatin-Induced Liver Toxicity: Hepatic Transglutaminase 7 Upregulation Associates with Oxidative Stress, Inflammation, and Apoptosis

Abstract

Transglutaminases (TGs) are calcium-dependent enzymes that cross-link proteins, contributing to apoptosis, extracellular matrix (ECM) stabilization, and inflammation. While TG2 has been extensively studied in hepatic injury, the role of TG7 in oxaliplatin-induced liver responses remains unclear. Oxaliplatin, a third-generation platinum chemotherapeutic, effectively treats solid tumors but can induce hepatic stress through oxidative and pro-inflammatory signaling. Adult rats received intraperitoneal oxaliplatin (10 mg/kg weekly) for 6 weeks. qRT-PCR, immunohistochemistry (IHC), immunofluorescence (IF), and a TG activity assay assessed hepatic TG7 expression, localization, and activity. Oxidative stress indicators (serum malondialdehyde [MDA] and reduced glutathione [GSH]) and pro-inflammatory cytokine transcription (CASP3, interleukin-6 (IL-6), tumor necrosis factor α (TNF-α)) were evaluated. Oxaliplatin exposure markedly increased TG7 mRNA and protein levels, elevated TG enzymatic activity, raised MDA (+49.4%), depleted GSH (−18.6%), and upregulated CASP3, IL-6, and TNF-α. DNA fragmentation and microscopic observations from IHC- and IF-processed sections were consistent with apoptosis-associated DNA degradation and subtle stress-related structural variations. Immunostaining revealed altered TG7 distribution within hepatocytes and sinusoidal regions. In this oxaliplatin-exposed rat liver model, TG7 upregulation and increased TG activity were associated with oxidative stress, inflammatory cytokine induction, and apoptotic signaling. These findings identify TG7 as a stress-associated marker during oxaliplatin exposure and support further studies to clarify its mechanistic role and evaluate its potential as utility as a biomarker under chemotherapy-associated hepatic stress conditions.

INTRODUCTION

Transglutaminases (TGs) are a conserved family of Ca²⁺-dependent enzymes that catalyze post-translational protein cross-linking through covalent ε-(γ-glutamyl)-lysine isopeptide bonds formed between glutamine and lysine residues. This reaction increases protein stability and protects substrates from proteolytic degradation.¹⁾ Beyond cross-linking, TGMs also mediate polyamine conjugation, lipid esterification, and glutamine deamidation, thereby broadening their physiological impact.²⁾ To date, nine members of this family have been characterized TG1–TG7, coagulation factor XIII, and erythrocyte membrane protein band 4.2, each displaying distinct structural motifs and tissue-specific expression patterns.³⁾ Eight isoenzymes are catalytically active, whereas band 4.2 is enzymatically inactive due to a cysteine-to-alanine substitution but remains critical for maintaining erythrocyte membrane integrity and mechanical resilience.⁴⁾ TG1, TG3, and TG5 are abundant in epithelial tissues; TG4 is predominantly prostate-specific; TG6 is expressed in the testis, lung, and brain; and TG7 is widely distributed but especially enriched in the testis and lung.⁴⁾ TG2 and factor XIIIa have multifunctional roles: TG2 contributes to extracellular matrix (ECM) stabilization, wound repair, and apoptosis regulation, while factor XIIIa is essential for blood coagulation.^5,6) TG2 also exhibits Ca²⁺-independent GTPase, kinase, and protein disulfide isomerase activities, expanding its functional repertoire.^6,7) Dysregulated TG activity has been implicated in a range of pathological processes, including cancer, fibrosis, chronic inflammation, and neurodegenerative diseases.^8–10)

Among these isoenzymes, TG7 remains one of the least explored. Early evidence suggests that TG7 may be responsive to cellular stress and could participate in remodeling or apoptotic processes; however, its functional role has not been clearly defined, and experimental data remain limited.^4,9) Understanding how TG7 behaves during hepatic stress may provide new insight into chemotherapy-related stress responses, but its involvement in liver pathology has not been well established.

Oxaliplatin (OXA) is a third-generation, platinum-based chemotherapeutic agent widely used for colorectal cancer and as part of combination therapies for gastric, ovarian, and lung malignancies.^11–13) Acting as an alkylating compound, OXA generates intra- and inter-strand DNA cross-links, disrupting replication and transcription, ultimately triggering apoptosis.¹⁴⁾ In addition, it promotes reactive oxygen species (ROS) formation and depletes reduced glutathione (GSH), producing oxidative stress and mitochondrial dysfunction.^15,16) Despite its survival benefits in advanced cancer, OXA’s nonspecific distribution can lead to adverse effects such as peripheral sensory neuropathy, cytopenias, gastrointestinal toxicity, hypersensitivity reactions, and liver injury.^17,18)

The hepatotoxic potential of oxaliplatin is now widely acknowledged. Lu et al.¹⁹⁾ reported that OXA intensified hepatic oxidative stress, inflammation, and fibrosis in a non-alcoholic fatty liver disease model, while Kalemci et al.¹⁸⁾ described histopathological liver alterations following OXA administration in rats. Chen et al.¹⁴⁾ further demonstrated that oxidative stress contributes to the synergistic anticancer effects observed when OXA is combined with piperlongumine. Collectively, these findings imply that ROS generation, while integral to OXA’s chemotherapeutic action, can exacerbate hepatic injury.

Transglutaminases, particularly TG2, are known to influence apoptosis and chemoresistance. TG2 overexpression has been linked to enhanced cancer cell survival and reduced chemosensitivity, whereas TG2 activation under certain stress conditions can promote apoptosis.^9,20) This functional duality underscores the context-dependent nature of TG activity, which may vary with tissue type, stress level, or injury severity. However, the role of TG7 in chemotherapy-induced liver injury remains largely uncharacterized and no studies to date have defined how TG7 responds to oxaliplatin-associated hepatic stress. Given TG7′s expression profile and potential stress-responsive behavior,⁴⁾ investigating its expression and activity under oxaliplatin exposure is necessary to establish whether TG7 participates in hepatic stress responses.

Reliable indicators of oxidative stress, such as malondialdehyde (MDA) and the antioxidant GSH, are routinely used to assess lipid peroxidation and redox homeostasis. Elevated MDA and diminished GSH have been observed in rat liver mitochondria following OXA exposure,¹⁵⁾ and antioxidant therapies have been shown to mitigate OXA-induced peripheral neuropathy.¹⁶⁾ Evaluating these markers alongside TG activity, TG7 expression, and protein localization offers a comprehensive assessment of OXA’s hepatic effects. Protein aggregation and cross-linking, processes commonly associated with TG activity, are frequently observed under cellular stress and may represent adaptive responses to injury.²¹⁾

The present study was designed to investigate the influence of oxaliplatin on TG activity, TG7 gene expression, and TG7 protein localization in rat liver tissue. By examining oxidative stress markers, inflammatory cytokine expression, and apoptotic indicators, this work aims to characterize the hepatic stress responses associated with TG7 upregulation during oxaliplatin exposure.

MATERIALS AND METHODS

Biological Samples and Experimental Design

Twenty adult male albino Wistar rats, Rattus norvegicus (7–8 weeks old; 150–200 g) were obtained from the Faculty of Pharmacy, Qassim University. All animal procedures were strictly followed by institutional ethical standards and adhered to the Guide for the Care and Use of Laboratory Animals.²²⁾ The Institutional Animal Care and Use Committee (IACUC) at Qassim University approved the study protocol (Approval No.: 25-37-22).

Rats were maintained in well-ventilated cages under controlled environmental conditions at 25 ± 1°C with a 12-h light/dark cycle and free access to standard laboratory chow and water. Following a 1-week acclimatization period, the animals were randomly allocated into two groups of ten rats each. The control group received no treatment and was maintained on a standard diet, whereas the oxaliplatin-treated group received intraperitoneal injections of oxaliplatin at a dose of 10 mg/kg body weight per week, dissolved in 5% glucose solution, for 6 consecutive weeks as described by Ashry et al.²³⁾ At the conclusion of the treatment period, all animals were fasted for 24 h, anesthetized with sodium phenobarbital (200 mg/kg b.wt. diluted 1 : 3 with phosphate-buffered saline, IP), and then sacrificed by capitation. Sodium phenobarbital was administered only once as a terminal anesthetic immediately before sacrifice and was not used during the oxaliplatin treatment period; therefore, it was not expected to induce hepatic drug-metabolizing enzymes or influence oxaliplatin metabolism during the experimental exposure phase. Blood samples were collected, and sera were separated by centrifugation before storage at −80°C for subsequent biochemical analyses. Livers were harvested and snap-frozen in liquid nitrogen for molecular and enzymatic investigations. Carcasses were disposed of at the Qassim University Veterinary Hospital in compliance with international biosafety and ethical disposal standards.

Ethics Approval

All experimental procedures were reviewed and approved by the Committee of Research Ethics at the Deanship of Scientific Research, Qassim University (Approval No. 25-03-17). All protocols were conducted following institutional guidelines and the ethical principles outlined in the Declaration of Helsinki.

Tissue Preparation and Immunofluorescence

To determine the localization of TGs in normal rat liver and oxaliplatin-treated tissues (OXP-TT), samples were fixed in 10% formaldehyde, embedded in paraffin, and sectioned at a thickness of 3 μm. Sections were deparaffinized in xylene and rehydrated through graded ethanol solutions. Immunofluorescence (IF) was performed according to the manufacturer’s instructions (Elabscience, Houston, TX, U.S.A.). The slides were fixed with 3.7% (w/v) paraformaldehyde in phosphate-buffered saline (PBS) for 15 min at room temperature, permeabilized with 0.1% (v/v) Triton X-100 in PBS for 15 min, and rinsed three times with PBS for 5 min after each step. Blocking was performed with 3% (w/v) BSA for 1 h at room temperature. For detection, slides were incubated with TG7 primary pAb, followed by anti-rabbit–Cy5 secondary antibody (red). Nuclei were counterstained with DAPI (blue). Negative controls were processed without the primary antibody. Fluorescence images were captured at 10× or 20× magnification using an EVOS FL AMF4300 fluorescence microscope (Life Technologies, Carlsbad, CA, U.S.A.). Quantification was performed on five randomly selected fields per biological sample (n = 10) animals per group using ImageJ software to quantify staining intensity and protein distribution.

Tissue Histology and Immunohistochemistry

Paraffin sections of normal and OXP-TT tissues were prepared as described above. After deparaffinization, antigen retrieval was carried out using sodium citrate buffer (pH 6.0) heated to 100°C for 10 min. Endogenous peroxidase activity was blocked with 0.3% hydrogen peroxide for 10 min at room temperature, after which the sections were incubated with a protein-blocking buffer for an additional 10 min. Primary antibodies against TG7 (1 : 1000; Abbexa, Cambridge, U.K.) were applied, and detection was performed using an HRP-conjugated polymer detection system. Visualization was achieved with a DAB substrate, and counterstaining was performed with hematoxylin. Negative controls were processed without the primary antibody. Quantification was performed on three randomly selected fields per biological sample (n = 10) animals per group using ImageJ software.

Protein and RNA Extraction from Liver Tissue

Liver specimens were prepared for homogenization by trimming away connective tissue and briefly rinsing in ice-cold PBS to remove residual blood. Approximately 50–100 mg of tissue was transferred to a chilled processing tube and disrupted with a Kinematica Polytron^® 1300D Homogenizer (Brinkmann Instruments, Westbury, NY, U.S.A.) under continuous cooling to prevent heat buildup. For protein isolation, the minced tissue was washed twice with 1 mL of cold PBS and centrifuged at 500 × g for 5 min. Pellets were resuspended in 1 mL of ice-cold Total Protein Extraction Buffer (TPEB) supplemented with a protease inhibitor cocktail (proteinSafe^™ Protease Inhibitor Cocktail, EDTA-free, 100×; DE101, Transgenbiotech, Beijing, China) and homogenized for 6–10 cycles in a pre-chilled glass homogenizer. Between samples, the homogenizer probe was thoroughly cleaned, rinsed three times with distilled water, once with 95% ethanol, and once with fresh lysis buffer, to prevent cross-contamination.

The homogenates were transferred to 1.5 mL microcentrifuge tubes, kept on ice to maintain protein integrity, and vortexed every 10 min during a 30-min incubation. After centrifugation at 14000 × g for 10 min at 4°C, the clarified supernatants were collected and stored at −80°C for downstream assays. Protein concentrations were determined using a BCA Protein Assay Kit (P310125-1250, Biofargo, Henrico, VA, U.S.A.) according to the manufacturer’s instructions.

For RNA extraction, total RNA and protein were also obtained using TRIzol^® Reagent (Life Technologies, Cat. No. 15596-026). Frozen liver tissues were homogenized in 1 mL of TRIzol^®, mixed with chloroform, incubated for 3 min, and centrifuged at 12000 × g for 15 min at 4°C. The aqueous phase was transferred to a clean tube, combined with 0.5 mL of 100% isopropanol, incubated for 10 min, and centrifuged again at 12000 × g for 10 min at 4°C. The resulting RNA pellet was washed, air-dried, and resuspended in RNase-free water. Before downstream applications, RNA purity and concentration were measured using a NanoDrop ND-2000c spectrophotometer (Thermo Fisher Scientific, Waltham, MA, U.S.A.).

Quantitative Real-Time PCR (qRT-PCR)

Specific primers for TG2, TG7, TG6, IL6, TNF, and GAPDH (Table 1) were designed with the PrimerQuest Tool (Integrated DNA Technologies, Coralville, IA, U.S.A.) to span exon–exon junctions and include known alternatively spliced variants. Reverse transcription and amplification were performed using the ABScript II One-Step SYBR Green qRT-PCR Kit (RK20404; ABclonal Technology, Wuhan, China), with GAPDH serving as the internal control. Reactions were run on an AriaMx Real-Time PCR System (Agilent Technologies, Santa Clara, CA, U.S.A.) in 20 μL mixtures containing SYBR Green buffer, 10 μL ABScript II enzyme mix, 0.8 μL forward primer (10 μM), 0.4 μL reverse primer (10 μM), 0.4 μL ROX dye (50×), 0.4 μL RNA, and 2 μL RNase-free water. Thermocycling conditions included reverse transcription at 42°C for 5 min, pre-denaturation at 95°C for 1 min, and 40 amplification cycles at 95°C for 5 s and 60°C for 30–34 s. Each sample was analyzed in duplicate, and all experiments were repeated independently at least three times. Cycle threshold (Ct) values were obtained using the Agilent AriaMx real-time PCR system. Relative gene expression was calculated using the comparative ΔΔCt method with Agilent AriaMx software. For each target gene, Ct values were first normalized to the internal reference gene GAPDH (ΔCt), and expression levels in untreated control liver tissue were used as the calibrator and set to 1. Expression levels in oxaliplatin-treated samples are presented as fold change relative to this control. Fold-change values were subsequently exported to GraphPad Prism for statistical analysis.

Table 1. Accession Numbers and Primer Sequences for qRT-PCR of TG Isoforms and Related Genes (5′→3′)

Gene/accession number	FWD set 1	REV set 1
TG2 NM_019386.3	TGTTGGTCAGAGGAGTGATTG	GGAGTGGACCTTGTGGTTATT
TG4 NM_022713.2	GATGCTGTGGAGCCTTAGTT	GCTCTTGAATCTGCCCTCATA
TG6 XM_063284796.1	CATCCTGAACATCTGCCTCTC	TCGGTCGTTGTTGCTGTT
TG7 XM_008762241.4	GGGTCTTCGCCTCTGTTATG	ATCTCGGCATTTCGGTCATAG
CASP3 NM_012922.3	CCACGGAATTTGAGTCCTTCT	CACTCCCAGTCATTCCTTTAG
TNF NM_012675.3	ACCTTATCTACTCCCAGGTTCT	GGCTGACTTTCTCCTGGTATG
IL6 NM_012589.2	GCCAGAGTCATTCAGAGCAATA	TAGGAGAGCATTGGAAGTTGG
GAPDH NM_017008.4	GACCACTTTGTCAAGCTCATTTC	CTCTCTTCCTCTTGTGCTCTTG

TG Activity Assay

TG activity was quantified using a colorimetric assay kit (Elabscience^®, Cat. No. E-BC-K840-M). The assay detects enzyme-mediated acyl transfer between glutamine residues and primary amines, producing a chromogenic compound measured at 525 nm. Measurements were performed in triplicate for each biological sample. In all, 60 μL samples were incubated with 200 μL of freshly prepared measuring solution at 37°C for 1 h in the dark. After the addition of 200 μL of chromogenic solution, the mixture was incubated for 10 min, centrifuged at 12000 × g for 10 min, and 200 μL of the supernatant was transferred to a 96-well plate for absorbance measurement. TG activity (U/g protein) was calculated from a standard curve using the formula:

  

$$TGActivity=\left(\Delta A525-b\right)/a\cdot Cpr\cdot {\displaystyle \frac{T}{f}}$$

where ΔA₅₂₅ is the difference between the sample and control optical densities, a and b are derived from the standard curve, Cpr is the protein concentration, f is the dilution factor, and T is the reaction time (1 h).

Serum MDA and GSH Estimation

Serum MDA and GSH concentrations were determined using commercial kits (MyBioSource, U.S.A.) supplied by Ejadah Trading Corporation, following the manufacturer’s instructions. MDA was measured using the thiobarbituric acid reactive substances (TBARS) method, in which MDA reacts with thiobarbituric acid at 95–100°C under acidic conditions to form a pink chromogen detected at 532 nm. Concentrations, expressed as nmol/mL, were calculated from an MDA standard curve. GSH levels were determined by reaction with 5,5′-dithiobis-(2-nitrobenzoic acid) (DTNB) to form 5-thio-2-nitrobenzoic acid, which was measured at 412 nm. GSH concentrations, expressed as μmol/mL, were derived from a standard curve. All samples were analyzed in duplicate, and kit quality controls were included to ensure assay reliability.

DNA Fragmentation Assay

Approximately 50–100 mg of liver tissue was homogenized in 1 mL of TRIzol^™ Reagent and allowed to stand for 5 min at room temperature to ensure complete dissociation of nucleoprotein complexes. Chloroform (0.2 mL per 1 mL TRIzol^™) was then added, mixed thoroughly, incubated for 2–3 min, and centrifuged at 12000 × g for 15 min at 4°C. After phase separation, the upper aqueous layer was discarded. To precipitate DNA from the interphase, 0.3 mL of 100% ethanol (per 1 mL TRIzol^™) was added, mixed gently, incubated for 2–3 min, and centrifuged at 2000 × g for 5 min at 4°C to collect the DNA pellet.

The pellet was washed twice with 0.1 M sodium citrate in 10% ethanol (pH 8.5) and once with 75% ethanol, followed by a 10–20 min incubation. After a final centrifugation step, the pellet was air-dried and resuspended in 8 mM NaOH. The solution was centrifuged at 12000 × g for 10 min, and the pH of the supernatant was adjusted with HEPES buffer. DNA samples were stored overnight at 4°C for short-term use or at −20°C (pH 7–8, 1 mM EDTA) for long-term storage. DNA concentrations and purity were measured using a NanoDrop^™ spectrophotometer.

For the assessment of fragmentation, a 1.5% (w/v) agarose gel was prepared in 1× TBE buffer containing SYBR Safe stain. All samples (control and oxaliplatin-treated) were processed identically to minimize extraction-related shearing. DNA samples were mixed with loading dye, loaded alongside a molecular weight ladder, and electrophoresed at 100 V for approximately 1 h in 1× TBE. After electrophoresis, gels were visualized under UV illumination. DNA fragmentation was evaluated based on the presence of smeared or laddered patterns after loading 20 μg of genomic DNA.

Statistical Analysis

Data were analyzed using GraphPad Prism (10.2.3). Statistical significance between the two groups was evaluated using unpaired two-tailed t-tests. Gene expression data were analyzed using multiple unpaired two-tailed t-tests with Holm–Šídák correction for multiple comparisons, with each gene analyzed independently by comparing control and oxaliplatin-treated groups. The results are presented as mean ± standard error of the mean (S.E.M.), and values of p < 0.05 were considered statistically significant.

RESULTS AND ANALYSIS

Gene Expression and Enzymatic Activity of TGs in Normal and Oxaliplatin-Treated Rat Liver Tissue

To investigate the effects of oxaliplatin on hepatic TGs, qRT-PCR was used to assess TG2, TG4, TG6, and TG7 mRNA levels in control and oxaliplatin-treated liver tissue (Fig. 1A). Messenger RNA levels were quantified relative to GAPDH and analyzed using multiple unpaired t-tests with Holm–Šídák correction. Oxaliplatin exposure produced only minor, non-significant changes in TG2, TG4, and TG6 expression compared with controls. In contrast, TG7 mRNA was markedly upregulated, rising to approximately 270–300-fold relative to control, representing a highly significant induction (****p < 0.0001). This strong induction indicates that TG7 is the most responsive isoform to oxaliplatin exposure under these experimental conditions, whereas other TG isoforms remained largely unchanged.

Fig. 1. Gene Expression and Enzymatic Activity of Transglutaminases (TGs) in Normal and Oxaliplatin-Treated Rat Liver Tissue

(A) mRNA was extracted from normal (Control) and oxaliplatin-treated liver (OXP.TT) samples and subjected to RT-qPCR using TGs-specific primers. Relative mRNA expression of TG2, TG4, TG6, and TG7 normalized to GAPDH and expressed as fold change relative to untreated control liver tissue (set to 1). Data are presented as mean ± S.E.M. (n = 10). Statistical analysis was performed using multiple unpaired two-tailed t-tests with Holm–Šídák correction for multiple comparisons. ****p < 0.0001. (B) Proteins extracted from the same liver tissues were subjected to a colorimetric TG activity assay. Data points represent mean ± S.E.M. TG-specific activity and were analyzed using an unpaired t-test to compare normal and oxaliplatin-treated liver tissue samples (n = 10; **p < 0.01). These results demonstrate selective induction of TG7 expression and marked elevation of TG activity following oxaliplatin exposure.

Complementing the transcriptional data, a colorimetric TG activity assay demonstrated a statistically significant elevation in TG-specific enzymatic activity in oxaliplatin-treated liver extracts compared with normal controls (mean ± S.E.M., n = 10; **p < 0.01) (Fig. 1B). Because this assay measures overall TG activity rather than TG7-specific activity, the increase is consistent with, but does not solely attribute, the enzymatic enhancement to TG7 induction. Together, these findings show that oxaliplatin treatment is associated with strong TG7 upregulation and increased hepatic TG activity.

TG7 Protein Expression and Localization in Normal and Oxaliplatin-Treated Liver Tissue

To investigate the protein expression and localization of TG7 following oxaliplatin treatment, immunohistochemical (IHC) staining was performed on liver tissue samples. The results revealed clear differential expression and localization of TG7 between the treated and control groups (Fig. 2). The presence of a strong positive IHC reaction in oxaliplatin-treated tissues (OXP-TT) compared with the control indicates an upregulation of TG7 protein expression in response to chemotherapy. Quantitative analysis of TG7-positive staining confirmed this observation, showing a significant increase in the treated samples (****p < 0.0001) (Fig. 2B).

Fig. 2. Transglutaminase (TG7) Protein Expression, Location, and Distribution in Rat Normal and Oxaliplatin-Treated Liver Tissue

(A) Immunohistochemical (IHC) staining of TG7 in liver tissue using a polyclonal antibody against TGM7. Panels (a–c) show representative sections from normal liver tissue, whereas panels (d–f) show representative sections from oxaliplatin-treated liver tissue. Representative negative-control IHC sections processed without primary antibody are shown in (c) for normal liver tissue and (f) for oxaliplatin-treated liver tissue. Images a, c, d, f acquired at 4× magnification (scale bar = 500 μm), and b and e (40×; scale bar = 50 μm). (B) the percentage quantification of positive TG7 IHC staining (%). An unpaired t-test compared the mean difference between normal and Oxaliplatin-treated liver tissue samples (n = 10, **p < 0.002).

Notably, TG7 localization was marked by yellow to brownish cytoplasmic staining, highlighting regions of elevated protein expression within hepatic tissue (Fig. 2A). The yellow areas suggest active TG7 expression, while the deeper brown coloration indicates sites of higher protein accumulation. In normal liver tissue, TG7 protein was weakly expressed and largely confined to scattered hepatocyte cytoplasm, with limited distribution around central veins. In contrast, oxaliplatin-treated liver tissue displayed intensified TG7 staining distributed broadly across hepatocyte cytoplasm and surrounding sinusoids. The broader and more intense TG7 staining in oxaliplatin-treated liver reflects greater TG7 protein abundance within hepatocytes and sinusoidal areas, consistent with the marked upregulation observed at the mRNA level.

Within IHC-processed sections used for TG7 localization, control liver tissue showed preserved cellular organization, whereas oxaliplatin-treated sections displayed subtle structural irregularities, such as mild variation in hepatocyte appearance and sinusoidal spacing. Because conventional H&E staining was not performed, classical histopathological features such as necrosis or inflammatory infiltration cannot be assessed, and these observations should be interpreted as qualitative structural features visible within immunostained sections rather than definitive histopathological alterations.

Immunofluorescent Analysis of TG7 in Normal and Oxaliplatin-Treated Liver Tissue

To further confirm TG7 protein expression and localization following oxaliplatin treatment, IF staining was performed on liver tissue samples. The results revealed a marked difference in TG7 signal intensity and distribution between control and oxaliplatin-treated tissues (Fig. 3A). Positive IF reactions were observed in oxaliplatin-treated liver (OXP.TT) compared with normal control, indicating a substantial upregulation of TG7 protein in response to chemotherapy. Quantitative analysis of TG7-positive staining supported this finding, showing a significant increase in TG7 levels in the treated group compared with control (****p < 0.0001) (Fig. 3B).

Fig. 3. Immunofluorescence Analysis of Transglutaminase 7 (TG7) and Its Levels in Normal and Oxaliplatin-Treated Liver Tissue

Immunostaining of transglutaminase 7 TG7 and its levels in Liver tissue and Oxaliplatin-treated Liver tissue. Paraffin-embedded Liver tissue for TG7 using a polyclonal antibody against TGM7. A) Negative IF reaction for TG7 in (–Ve Con.) normal and Oxaliplatin-treated Liver tissue (–Ve OXP.TT); Positive IF reaction for TG7 in (Con.) normal and Oxaliplatin-treated liver tissue (OXP.TT). The original magnification of the images was 10×, and 20×, respectively and the scale bar was 200 or 400 μm. B) the percentage quantification of positive TG7 IF staining. An unpaired t-test compared the mean difference between normal and Oxaliplatin-treated Liver tissue samples (n = 10, ****p < 0.0001). DAPI (blue), TG7 (red (Cy5)).

TG7 localization was visualized as distinct red fluorescence (Cy5) overlaid with DAPI-stained nuclei (blue) (Fig. 3A). In the control tissue, TG7 staining appeared faint and sparsely distributed within hepatocyte cytoplasm. In contrast, oxaliplatin-treated liver exhibited intensified red fluorescence with widespread cytoplasmic distribution, indicating enhanced TG7 accumulation and broader tissue involvement. Negative controls (–Ve Con. and –Ve OXP.TT) confirmed minimal background signal, validating staining specificity.

Evaluation of IF-processed sections revealed preservation of general tissue organization in the controls, whereas oxaliplatin-treated samples showed localized regions of altered cellular arrangement associated with increased TG7 fluorescence intensity. These features were modest and do not indicate overt necrosis, but are consistent with early cellular stress responses observable during TG7 localization, rather than definitive histopathological injury.

Apoptotic and Inflammatory Responses in Normal and Oxaliplatin-Treated Rat Liver Tissue

To evaluate oxaliplatin-induced inflammatory and apoptotic responses, CASP3, IL6, and TNFα mRNA expression was quantified by qRT-PCR in liver tissue. Compared with the control group (untreated control liver tissue), oxaliplatin treatment caused significant upregulation of all three genes (Fig. 4A). IL6 showed a moderate but significant increase (**p < 0.01), while CASP3 and TNFα exhibited more robust induction, with TNFα demonstrating the highest elevation among the assessed markers (****p < 0.0001). Statistical analysis was performed using one-way ANOVA followed by Dunnett’s multiple comparisons test (n = 10). These transcriptional findings indicate that oxaliplatin exposure is associated with increased expression of apoptotic and inflammatory markers in liver tissue.

Fig. 4. Apoptotic and Inflammatory Responses in Normal and Oxaliplatin-Treated Rat Liver Tissue

(A) mRNA was extracted from untreated (Control) and oxaliplatin-treated liver (OXP.TT) samples and subjected to RT-qPCR using specific primers. Relative mRNA expression of CASP3, IL6, and TNF-α was normalized to GAPDH and expressed as fold change relative to untreated liver tissue (set to 1). Data are presented as mean ± S.E.M. (n = 10). Statistical analysis was performed using multiple unpaired two-tailed t-tests with Holm–Šídák correction. **p < 0.01, ***p < 0.001, ****p < 0.0001. (B) Genomic DNA fragmentation analysis. Lane M, molecular weight marker; lanes 1–2, control liver DNA; lanes 3–6, oxaliplatin-treated (OXP.TT) liver DNA. Equal amounts of genomic DNA (20 μg per lane) were electrophoresed on an agarose gel. Control lanes display predominantly intact high-molecular-weight DNA, whereas oxaliplatin-treated lanes show increased low-molecular-weight DNA smearing below the loading wells, consistent with apoptosis-associated DNA degradation when interpreted together with CASP3 upregulation. The 25 kb reference position is indicated. Minor background irregularities likely reflect technical variation inherent to genomic DNA extraction and electrophoresis and do not alter the overall qualitative pattern.

Genomic DNA fragmentation analysis revealed predominantly intact high-molecular-weight DNA in control samples (lanes 1–2), whereas oxaliplatin-treated liver samples (lanes 3–6) exhibited increased low-molecular-weight DNA smearing below the loading wells (Fig. 4B). This pattern is consistent with apoptosis-associated DNA degradation rather than classical internucleosomal laddering. Although minor background irregularities were observed, all samples were processed identically, and the qualitative increase in DNA fragmentation in oxaliplatin-treated samples aligns with the concurrent upregulation of CASP3 expression observed at the transcript level.

Oxaliplatin-Induced Oxidative Stress: Increased MDA and Reduced GSH Levels

Intraperitoneal administration of oxaliplatin (10 mg/kg body weight/week for 6 weeks) resulted in a significant increase in serum MDA levels compared with the control group (27.09 ± 0.90 vs. 18.13 ± 0.40 nmol/mL; ****p ≤ 0.0001) (Table 2). In contrast, oxaliplatin treatment caused a significant reduction in serum GSH levels (69.78 ± 0.40 vs. 85.77 ± 0.46 μmol/mL), representing an 18.64% decrease relative to controls. These changes indicate higher lipid peroxidation and decreased antioxidant capacity in oxaliplatin-treated rats.

Table 2. Mean Serum Oxidative Stress Markers in Control and Oxaliplatin-Treated Rats

Group	MDA (nmol/mL)	GSH (μmol/mL)
Control	18.13 ± 0.40	85.77 ± 0.46
Oxaliplatin	27.09 ± 0.90****	69.78 ± 0.40****
% Change	+49.4%	−18.64%

Data are expressed as mean ± S.E.M. (n = 10 per group). Statistical significance between control and oxaliplatin-treated groups was assessed using an unpaired two-tailed Student’s t-test; ****p ≤ 0.0001. GSH, reduced glutathione; MDA, malondialdehyde.

DISCUSSION

The present study shows that TG7 is markedly upregulated in the livers of oxaliplatin-treated rats, indicating that this isoform is particularly responsive to oxaliplatin-associated hepatic stress. Platinum drugs such as oxaliplatin generate reactive oxygen species (ROS) and deplete GSH, creating oxidative stress that drives mitochondrial dysfunction and lipid peroxidation^19,24,25) Elevated ROS levels, disrupted Ca²⁺ homeostasis, and GSH depletion are established triggers for TG activation.^26,27) Comparable xenobiotic modulation of TG activity has been observed with organophosphates, which induce TG2 activation and covalent adduct formation in neuronal cells,^10,28) demonstrating that chemical stress can directly influence TG function.

Within the TG family, TG2 has been more thoroughly investigated in hepatic injury and fibrosis. TG2′s cross-linking activity contributes to ECM stabilization, reinforcing collagen and fibronectin deposition in fibrotic liver,^29,30) which promotes tissue stiffening and sinusoidal narrowing. Although TG7 has received less attention in hepatic research, its robust induction in our oxaliplatin model highlights it as an isoform that responds strongly to chemotherapeutic stress. Under oxidative conditions, cellular proteins undergo structural alterations that can increase their susceptibility to TG-mediated modification; however, whether TG7 acts on these proteins in the liver remains to be clarified. If TG7 activity increases under oxidative stress, it could theoretically contribute to stabilizing altered proteins or forming protein aggregates; however, this has not been demonstrated in the present study and remains a potential mechanism for future investigation. Stress-induced activation of TG isoforms has been previously reported in extrahepatic tissues,³¹⁾ and this is consistent with our finding that oxaliplatin-treated liver displayed a significant increase in TG7 expression together with elevated total TG enzymatic activity. However, although TG7 activation was clearly observed in this model, the upstream regulatory mechanisms driving this activation in hepatocytes were not investigated and remain to be clarified.

Although routine serum biomarkers of hepatocellular injury (ALT, AST, and bilirubin) were not reassessed in the current experiment, the biological activity and systemic toxicity of this oxaliplatin regimen have been validated previously by our laboratory group. Using the same dose, route, and 6-week exposure protocol, an earlier study from our group demonstrated significant oxidative, inflammatory, and apoptotic damage in testicular tissue, confirming that this dosing regimen reliably induces systemic cytotoxic stress.²³⁾ Based on this established systemic effect, the present study focused specifically on hepatic TG7 regulation, oxidative stress, and apoptotic signaling, rather than repeating previously characterized general toxicity endpoints.

Oxaliplatin exposure has been reported to elicit a spectrum of hepatic stress responses, including oxidative imbalance, endothelial perturbation, and ECM alterations.¹⁹⁾ TG2′s role in liver fibrosis through TGF-β signaling and ECM cross-linking has been confirmed in recent studies.⁶⁾ TG7′s pronounced induction in this model raises the possibility that it may contribute to early changes that precede ECM remodeling, although direct evidence of TG7-mediated ECM modification was not assessed in this study. Although TGs have been reported in other systems to influence cellular signaling through protein cross-linking, the functional consequences of TG7 induction in hepatocytes were not investigated here and remain speculative. It is therefore possible, but unproven, that TG7 upregulation could participate in stress-related cellular changes observed during oxaliplatin exposure. Previous hepatic injury studies show that changes in ECM behavior can influence inflammatory and stress–response pathways.³²⁾ Although ECM remodeling was not directly assessed here, the clear and selective induction of TG7 observed in oxaliplatin-treated liver suggests that TG7 may be positioned upstream of processes that potentially lead to such alterations, a possibility that warrants dedicated future investigation. Given its robust induction alongside oxidative stress, inflammation, and apoptosis, TG7 may act as an early responder to oxaliplatin-induced cellular injury, positioning it as a plausible upstream modulator of subsequent hepatic remodeling processes that require further investigation.

The marked induction of IL-6 and TNF-α in oxaliplatin-treated liver indicates that inflammatory cytokines play a significant role in the hepatic stress response observed in this model. Oxaliplatin-generated oxidative stress is known from previous studies to activate pathways such as nuclear factor-kappaB (NF-κB) and STAT3, which drive IL-6 and TNF-α expression; although these signaling events were not directly assessed here, the increased cytokine levels observed in this study are consistent with such ROS-responsive mechanisms.^18,19) Although TG7-specific regulation has not yet been fully defined, the cytokine elevations observed here suggest that similar inflammatory feedback mechanisms may influence TG7 during oxaliplatin-induced stress. Pro-inflammatory cytokines such as IL-6 and TNF-α have been shown in previous studies to modulate TG expression and activity, particularly for TG2 in liver injury models.²⁶⁾ Bioinformatic annotations indicate that the TG7 gene contains potential regulatory elements, including sites associated with NF-κB and SP1 activity. While these motifs were not experimentally examined in this study, their presence suggests that TG7 could be responsive to inflammatory or stress-related signaling pathways. Stress-responsive kinase pathways such as PKC and PKA are known to modulate TG2 expression and activity in hepatic injury models.³³⁾ Although TG7-specific regulation by these kinases has not yet been established, the pronounced TG7 induction observed here raises the possibility that similar signaling cross-talk may contribute to its activation during oxaliplatin-induced stress. Notably, PKC- and PKA-related signaling pathways are active targets in current antifibrotic drug development.³⁴⁾ While TG7 regulation by these pathways has not been defined, their involvement in hepatic stress signaling raises the possibility that future therapeutic strategies targeting upstream kinase networks could indirectly influence TG7 activity.

The concurrent upregulation of TG7 and CASP3 in oxaliplatin-treated liver strongly suggests that TG7 participates in apoptotic remodeling during hepatic stress. Within the TG family, several isoforms are known to be cleaved by caspases to alter their activity, and can cross-link apoptotic substrates and compact chromatin to stabilize apoptotic bodies.^27,32) The concurrent induction of TG7 and CASP3 suggests a potential functional relationship, where TG7 may influence apoptotic signaling or participate in structural stabilization during the execution phase of apoptosis; however, this possibility remains hypothetical and is presented as a mechanistic interpretation rather than a measured outcome. Findings from organophosphate exposure studies demonstrate that TG family members can function as cellular stress sensors and effectors.^10,28) These observations support the possibility that TG7 may participate in analogous stress–response pathways in hepatocytes following oxaliplatin exposure. Additionally, the documented high expression of TG7 in organs such as the lung and testis³⁵⁾ suggests that this enzyme may participate in conserved cellular stress–response pathways across different tissues, supporting the broader physiological relevance of its induction in the liver.

DNA fragmentation analysis in this study was intended as a qualitative indicator of apoptosis-associated DNA degradation rather than as a demonstration of classical internucleosomal laddering. In solid tissues, particularly liver, apoptotic DNA fragmentation frequently appears as diffuse smearing rather than distinct internucleosomal ladders on agarose gels, because only a subset of cells undergoes apoptosis at any given time and the degree of internucleosomal cleavage can vary with cell type and extraction conditions, limiting the visibility of discrete ladder patterns in heterogeneous samples^36,37) Accordingly, the DNA fragmentation data are interpreted as supportive evidence when integrated with molecular apoptotic markers, rather than as a standalone diagnostic assay. Together, these complementary observations support the presence of apoptotic signaling under oxaliplatin-induced hepatic stress while acknowledging the technical limitations inherent to genomic DNA electrophoresis.

In established liver disease models, hepatotoxic insults can evolve from acute stress into chronic fibrotic remodeling, and persistent TG activity, well described for TG2 and other family members, can promote ECM cross-linking, generate proteolysis-resistant matrix structures, and sustain inflammatory signaling.²⁹⁾ Recent fibrosis models show that reducing TG activity, particularly TG2, attenuates fibrotic remodeling,^38,39) supporting the concept that TG-mediated cross-linking contributes to early matrix stiffening; given the marked induction of TG7 in the present study, it is plausible that TG7 may participate in similar remodeling processes during oxaliplatin-induced hepatic stress. As hepatocytes become surrounded by a progressively stiffened ECM, alterations in mechano-transduction pathways can further sustain NF-κB activation, amplifying pro-inflammatory signaling and increasing susceptibility to apoptotic priming. These reinforcing feedback loops between ECM remodeling, inflammatory signaling, and apoptotic activation are well recognized as key drivers of chronic liver disease progression.²⁷⁾

Evidence from extrahepatic systems supports this interpretation; for example, in neuronal models, organophosphate exposure increases TG activity, which contributes both to stress adaptation and to the accumulation of cross-linked damaged proteins, ultimately exacerbating cellular toxicity.^10,28) A similar mechanism could occur in hepatic tissue, where TG7 cross-links oxidized or alkylated proteins after oxaliplatin exposure, forming proteolysis-resistant aggregates that impede recovery.

Therapeutically, interventions that reduce oxidative stress or modulate TG activity have shown benefit in experimental models, suggesting that antioxidant supplementation or TG-directed strategies may help attenuate oxaliplatin-induced hepatic stress. Glutathione supplementation and other ROS-scavenger approaches have been shown to reduce oxaliplatin-induced hepatotoxicity in experimental studies, underscoring the central role of oxidative stress in mediating the observed liver injury.^24,40) TG inhibitors currently studied in TG2-driven fibrotic models may eventually be adapted to target TG7 once isoform-selective tools become available, highlighting a potential avenue for future therapeutic development.^38,39) Upstream signaling pathways such as PKC and PKA are known regulators of TG2 activity under oxidative or inflammatory conditions,³³⁾ raising the possibility that similar kinase-dependent mechanisms may influence TG7, although this has not yet been experimentally demonstrated. Combination approaches that incorporate antioxidants, kinase modulators, or TG-targeted agents may offer a promising strategy to lessen oxaliplatin-associated hepatic injury while preserving the drug’s antitumor effectiveness.

Finally, the schematic model in Fig. 5 provides a hypothesis-based summary integrating our experimental findings with established knowledge of TG biology. In the present model, oxaliplatin exposure increased ROS levels and depleted GSH, indicating oxidative stress, a condition known to influence TG family activation. Although NF-κB/STAT3 signaling was not experimentally assessed in this study, these pathways are well documented in oxidative stress-associated hepatic injury and are therefore included in the figure strictly as conceptual intermediates rather than demonstrated mechanisms. In this context, TG7 upregulation may be associated with stress-related cellular processes such as altered protein cross-linking capacity, mechano-transduction sensitivity, or increased apoptotic susceptibility; however, these downstream roles remain speculative and are shown as hypothetical links. The observed increase in CASP3 expression together with qualitative DNA fragmentation supports the presence of apoptotic signaling, which may interact with inflammatory pathways during oxaliplatin exposure. Overall, the model illustrates a potential framework in which TG7 is positioned at the interface of oxidative stress, inflammation, and apoptosis during oxaliplatin-associated hepatic stress, while emphasizing that these mechanistic relationships require further experimental validation.

Fig. 5. Hypothesis-Based Schematic Model of TG7 Involvement in Oxaliplatin-Induced Hepatic Stress Responses

Solid arrows indicate pathways supported by the experimental findings of this study, including oxaliplatin-induced oxidative stress (ROS increase and GSH depletion), TG7 upregulation, increased transglutaminase activity, pro-inflammatory cytokine induction (IL-6 and TNF-α), and CASP3-associated apoptotic signaling. Dashed arrows denote hypothetical or literature-based mechanisms, including Ca²⁺ influx, NF-κB/STAT3 activation, TG7-mediated cross-linking of intracellular and extracellular substrates, and potential contributions to early fibrotic remodeling. This model integrates the present findings with established transglutaminase biology and is intended as a conceptual framework rather than experimentally confirmed signaling cascades^6,18,19,26).

Limitations and Future Directions

A limitation of this study is that classical hepatotoxicity endpoints, including serum transaminases (ALT/AST) and H&E histopathology, were not assessed; therefore, conclusions are restricted to oxidative, inflammatory, and apoptosis-associated stress responses observed under oxaliplatin exposure. This study used a single oxaliplatin dose and time point, limiting our ability to characterize temporal progression or dose-dependent effects. We did not identify the specific protein substrates of TG7 nor experimentally validate upstream regulators that may drive its induction. Although TG7 upregulation coincided with oxidative stress and apoptotic markers, the relationship is correlative rather than causal. Future studies should include TG7 knockout or knockdown models to clarify functional contributions, alongside proteomic mapping of TG7-mediated cross-links. Promoter and pathway analyses under cytokine or kinase modulation would also help define upstream regulatory mechanisms. Longitudinal studies across multiple chemotherapeutic agents are needed to determine whether TG7 acts primarily as a stress responder, a fibrogenic mediator, or a biomarker of hepatotoxicity. Phenobarbital was used only as a single terminal anesthetic dose immediately before sacrifice; thus, it is unlikely to have influenced hepatic enzyme induction during oxaliplatin exposure.

CONCLUSION

This work identifies TG7 as a selectively responsive isoform in oxaliplatin-treated liver tissue. Oxaliplatin exposure was associated with oxidative stress, increased inflammatory cytokine expression, and activation of apoptotic pathways, with TG7 showing markedly greater induction than other TG family members. These findings highlight TG7 as a potentially important component of the hepatic response to oxaliplatin-induced stress. By situating TG7 within established TG biology, our study provides a conceptual foundation for investigating TG7 in apoptosis, stress adaptation, and ECM remodeling. Although further mechanistic work is required, the current data support continued exploration of TG7 as a candidate biomarker and as a possible target for strategies aimed at reducing chemotherapy-associated hepatotoxicity. Future studies employing TG7-specific loss-of-function models and protein-interaction mapping will be essential to advance this translational potential.

Acknowledgments

The authors thank the Deanship of Graduate Studies and Scientific Research at Qassim University for financial support (QU-APC-2025-6/26). They also gratefully acknowledge Prof. Methat Rehan, Professor of Molecular Biotechnology, College of Agriculture, Qassim University, for his generosity in providing access to essential laboratory facilities and for his support and trust during key phases of the experimental work.

DECLARATIONS

Funding

The authors gratefully acknowledge Qassim University, represented by the Deanship of Graduate Studies and Scientific Research, for the financial support for this research under the number (QU-I-PG-2-2025- 54152) during the academic year 1446 AH/2024 AD.

Author Contributions

H. M. A., I. S. A., and H. F. G. designed the experiments; H. M. A., I. S. A., and H. F. G. performed the experiments and collected data; H. M. A., I. S. A., and H. F. G. discussed the results and strategy; I. S. A. and H. F. G. supervised, directed, and managed the study; H. M. A., I. S. A., and H. F. G. gave final approval of the version to be published.

Conflict of Interest

The authors declare no conflict of interest.

Use of Generative AI

During manuscript preparation, AI tools were used only for language improvement; the authors take full responsibility for the content.

Data Availability

All data are available upon reasonable request.

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