1 Introduction
Methotrexate (MTX), a folate antagonist, is a competitive inhibitor of dihydrofolate reductase which converts dihydrofolate into tetrahydrofolate required for nucleic acid synthesis (Kremer, 2004; Koźmiński et al., 2020). However, it is used in high doses for managing various cancers such as acute lymphoblastic leukemia, osteosarcoma, and breast cancer (Ezhilarasan, 2021). Furthermore, it is used for gestational choriocarcinoma (Chin, 2023). Low-dose MTX is used as an anti-inflammatory and immunomodulatory drug as the first line and centerpiece therapy for rheumatoid arthritis and as maintenance therapy for Crohn’s disease and psoriasis (Pivovarov and Zipursky, 2019; Hsieh and Tsai, 2023). Unfortunately, however, adverse effects associated with MTX use such as gastrointestinal toxicity, hepatotoxicity, nephrotoxicity, pulmonary toxicity, and neurotoxicity limit its clinical application (Triantafyllou et al., 2010; Mahmoud et al., 2017b). Steatohepatitis, fibrosis, as well as cirrhosis, are forms of liver toxicity triggered by MTX (Romão et al., 2014). Among MTX-treated patients, the prevalence of hepatic fibrosis and cirrhosis was up to fifty percent and twenty-six percent, respectively (Roghani et al., 2020). Meanwhile, MTX’s acute hypersensitivity pneumonitis and interstitial lung inflammation which usually appear within the first year of therapy occur in approximately 8% of patients with an estimated mortality of 13%–17% (Romão et al., 2014). Moreover, lung fibrosis can also occur with MTX (Juge et al., 2021).
The pathogenesis of MTX-induced hepatotoxicity and lung damage is not well clarified. However, it was documented that oxidative stress is the critical factor, as it can initiate an inflammatory response and oxidative DNA damage, inducing apoptotic cell death (Mahmoud et al., 2017b; Chauhan et al., 2020; Ozmen et al., 2024). Furthermore, the profibrotic effects of MTX were identified in both liver and lung tissues (Mohamed et al., 2019). MTX activates fibroblasts, which then induce extracellular matrix synthesis, leading to tissue fibrosis and organ dysfunction (Fayez et al., 2018). That is why finding therapeutic agents that can be used with MTX to reduce the incidence and severity of its associated adverse effects is sought after.
Galectin-3 (Gal-3) is a multifunctional mammalian β-galactoside-binding lectin that is expressed on the cell surface, nucleus, cytoplasm, and extracellularly. It is mainly secreted by macrophages and participates in numerous biological events, for example, cell adhesion, migration, angiogenesis, and apoptosis (Nangia-Makker et al., 2000; Henderson et al., 2008; Li et al., 2014). Its key role in tissue inflammation and fibrosis was documented. Gal-3 activation in different fibrotic models in addition to abnormally elevated levels in patients who have liver, lung, or heart fibrosis have been reported (Nishi et al., 2007; Bayes-Genis et al., 2014; Sciacchitano et al., 2018).
Modified citrus pectin (MCP) is obtained from citrus fruit as a water-soluble dietary fiber. It is a natural inhibitor of Gal-3 by binding directly to its carbohydrate recognition domain (Glinsky and Raz, 2009; Gunning et al., 2009). Recently, MCP gained popularity due to its anti-cancer (Glinsky and Raz, 2009; Leclere et al., 2013; Garrido et al., 2024), anti-inflammatory, and antifibrotic effects (Kolatsi-Joannou et al., 2011; Abu-Elsaad and Elkashef, 2016; Li et al., 2018; Marín-Royo et al., 2018; Xu et al., 2020; Cui et al., 2022; Bouffette et al., 2023) in several diseases. It showed hepatoprotective effects against carbon tetrachloride (CCl4)-caused liver fibrosis in rats through antioxidant and Gal-3 blockade-mediated antifibrotic and antiapoptotic effects (Abu-Elsaad and Elkashef, 2016). Nevertheless, the involvement of Gal-3 inhibition in MTX-elicited liver and lung illnesses is still unclear.
For the first time, our goal was to discover the role of Gal-3 in the pathogenesis of MTX-induced hepatotoxicity and lung toxicity and the possible protective effects of its natural inhibitor, MCP. We also explored the potential signaling pathways that could explain these suggested preservative effects. Finally, different human cancer cell lines were used here to analyze the impact of MCP on the cytotoxic effect of MTX as an in vitro part of the study.
2 Materials and methods
2.1 Drugs
Methotrexate was procured from the Austrian pharmaceutical company EBEWE Pharma, Ges.m.b.H. Nfg. KG. MCP (Pectasol) was obtained from EcoNugenics, Santa Rosa, CA, U.S.A.
2.2 Animals
2 Here, we procured adult male albino Wistar rats (180–220 g) from Helwan Farm in Cairo, Egypt, owned by Vacsera Company. Animals had free access to water and food throughout the 2 weeks of the acclimatization period and the experiment period. They were subjected to a 25°C ± 2 temperature and 12:12 h of dark/light cycles. With the approval number: MPEC-230506, the present study protocol complied with the regulations set out by the Research Ethics Committee at the Faculty of Pharmacy at Minia University in Egypt.
2.3 Experimental design
After randomly dividing the rats into four groups of eight, they were administered the following dosage schedule:
1. Control group: Rats were given distilled water (MCP vehicle) orally for 14 days and a single intraperitoneal saline (MTX Diluent) injection on day 9.
2. MCP group: Rats were administered MCP (200 mg/kg/day, orally for 14 days) and a single intraperitoneal saline injection on day 9.
3. MTX group: Rats received distilled water orally for 14 days and a single intraperitoneal MTX injection (40 mg/kg) on day 9.
4. MTX + MCP group: Rats were given MCP (200 mg/kg/day, orally for 14 days) and a single intraperitoneal MTX injection (40 mg/kg) on day 9.
The dosage and timing for MCP were determined based on our preliminary studies and prior pharmacological investigation (Li et al., 2021) which showing its protective impact against organ damage. Meanwhile, the MTX dose was chosen to be sufficient to elicit hepatic and pulmonary injury in male rats (Letertre et al., 2020; Dogra et al., 2021; Matouk et al., 2022; Matouk et al., 2023). MTX injection on day 9 was done based on prior research (Matouk et al., 2022). Furthermore, our preliminary study results indicated that MCP showed hepatic and pulmonary protective effects when given 200 mg/kg/day for 8 days before giving MTX on day 9.
2.4 Blood and tissue sampling
Rats were put under anesthesia 24 h following the final dose. A cardiac puncture was made to take samples of blood in clean centrifuge tubes and then centrifuged at 3,500 rpm for 10 min to get sera which were used freshly for liver function assessment. The liver and lungs were rapidly isolated and weighed after drying on filter paper. Relative liver and lung weights were calculated (Relative organ weight = ((organ weight/body weight) × 100) (Jakkula et al., 2000; Wan et al., 2021). Immunohistochemical and histopathological examinations were conducted on parts of the lower lobe of the right lung and the medial lobe of the liver after fixing them in 10% formalin. Before the biochemical analysis, the residual liver parts and left lung were kept at −80°C after being quickly frozen in liquid nitrogen.
2.5 Evaluation of microvascular permeability and lung edema
Rapidly after blood collection by cardiac puncture, we obtained the bronchoalveolar lavage fluid (BALF) through tracheal intubation and lavage of both lungs with 2 mL saline. The BALF samples underwent centrifugation at 1,000 rpm at 4°C for 10 min (Rajizadeh et al., 2024). The supernatant was used to measure the BALF total protein spectrophotometrically using a commercially available kit (BioMed, Cairo, Egypt) according to Kingsley (1939). Meanwhile, total leukocyte count was detected by Mindray Bc-20 s Auto Hematology analyzer after resuspending the cell pellet in 0.5 mL phosphate buffer saline (PBS) (Zhao et al., 2021). To assess the wet/dry (W/D) weight ratio as an indicator of lung edema, the right upper lobe weight was determined alone after lung separation, wet weight. Then 24 h of drying was done in an oven at 80°C, dry weight (Zhang et al., 2021).
2.6 Determination of liver function markers
The serum alanine aminotransferase (ALT) and aspartate aminotransferase (AST) levels were assessed using commercial kits purchased from Biodiagnostic, Cairo, Egypt as described by Reitman and Frankel (1957).
2.7 Liver and lung histopathology
After fixation of the liver and lung sections in 10% formalin, they were embedded in paraffin blocks after dehydration with graded ethanol series, cleared with xylene, and then sectioned into 4–6 µm thickness slices. For histopathological examination, deparaffinization and hematoxylin and eosin (H&E) staining of the produced sections were performed following Bancroft and Gamble (2008). The damage was evaluated according to the scoring system by Plaa et al. (1994) in the liver and by Eldh et al. (2012) in the lung. Moreover, Masson’s trichrome staining was done to evaluate organ fibrosis. Fibrosis was assessed semi-quantitatively in ×200 magnification. The area percentage (%) of tissue with fibrotic changes on Masson’s trichrome-stained sections was evaluated using ImageJ software (Van De Vlekkert et al., 2020).
2.8 Assessment of oxidative stress parameters in liver and lung homogenates
Lipid peroxidation in hepatic and lung homogenates was analyzed as thiobarbituric acid reactive species (TBARS) named malondialdehyde (MDA) employing the methodology established by Buege and Aust (1978). Reduced glutathione (GSH) content and superoxide dismutase (SOD) activity have been investigated as markers of endogenous antioxidant defense. Ellman (1959) methodology was used to measure GSH content. Whereas, SOD activity was assessed following Marklund (1985).
2.9 Assessment of Nrf2, TLR-4, NF-κB, and c-caspase-3 using western blot analysis
Hepatic and pulmonary tissues were homogenized using a protease inhibitor cocktail (Biospes, China) and tris lysis buffer for 30 min at 4°C following the method described by Ali et al. (2018). After that, centrifugation at 10,000 rpm for 10 min at 4°C was done to remove residual tissue. Total protein concentrations were assessed using the Biuret method Wang et al. (1996). Utilizing 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis, equivalent quantities of protein, 30 μg of total protein for each lane, underwent electrophoresis and then transferred to a polyvinyl difluoride membrane (Millipore, USA) applying semi-dry transfer methods (Towbin et al., 1979). To block the membranes, they were incubated for 60 min at room temperature with 5% non-fat milk in tris-buffered saline Tween 20.
Next, the membranes were incubated with the primary antibodies for the target proteins overnight at 4°C; toll-like receptor-4 (TLR4) (dilution 1:1,000) (Santa Cruz Biotechnology, Inc., sc-293072), nuclear factor erythroid 2-related factor 2 (Nrf2) (dilution 1:500), nuclear factor-kappa B (NF-κB p65) (dilution 1:500), cleaved caspase-3 (c-caspase-3) (dilution 1:500) (Chongqing Biospes Co., Ltd., China, YPA1865, BBP1066, and YPA2210, respectively) and β-actin (dilution 1:3,000) (Elabscience Biotechnology, Inc., E-AB-20031). The membranes were mixed with an alkaline phosphatase-conjugated secondary antibody (dilution 1:5000) for 1 hour obtained from Biospes, China. Band visualization was achieved by BCIP/NBT substrate detection Kit obtained from Genemed Biotechnologies, United States of America. Analysis of the produced bands compared to the internal control β-actin bands was conducted utilizing ImageJ® software (National Institutes of Health, Bethesda, Maryland, United States of America).
2.10 Assessment of TNF-α using enzyme-linked immunoassay (ELISA) technique
A rat TNF-α ELISA kit was used to assess tumor necrosis factor-alpha (TNF-α). It employs the sandwich ELISA principle, wherein samples were added to precoated microwells with TNF-α specific antibody. The microplate was then incubated with avidin-horseradish peroxidase (HRP) conjugate and biotinylated antibody. Substrate solution was added, causing a blue color. A stop solution was used to end the reaction, turning blue to yellow. At 450 nm, the optical density was measured which is directly proportional to the TNF-α concentration.
2.11 Assessment of Gal-3 and TGF-β using immunohistochemistry
After deparaffinizing and rehydrating the liver and lung tissue sections, they were soaked in hydrogen peroxide and washed in buffer to inhibit peroxidase activity. Non-specific background staining was blocked using Ultra V block. Antibodies targeting Gal-3 (Novocastra laboratories, UK, clone 9c4) and transforming growth factor-beta (TGF-β1) (Chongqing Biospes Co., Ltd., China, YPA1196) were incubated with liver and lung tissue sections exactly as directed by the manufacturer. Afterward, they were incubated at room temperature for 10 min with a primary antibody enhancer and then for 15 min with HRP polymer. Hematoxylin was used as a counterstain to contrast the chromogen color (Shan et al., 1999).
Immunoreactivity was assessed semi-quantitatively in high microscopic power fields (X400). The area percentage (%) of positively stained cells was evaluated by using Fiji ImageJ software (Schindelin et al., 2012).
2.12 <italic>In vitro</italic> analysis
Cell culture was done at Vacsera-cell culture laboratory, Cairo, Egypt. The cytotoxicity assay was evaluated on the tested drugs using breast cancer (MCF7), acute lymphoblastic leukemia (Nalm6), and choriocarcinoma (JEG3) cell lines. The American Type Culture Collection in Manassas, Virginia, United States of America was the source of all the cell lines. These cells were grown in RPMI 1640 media provided with 10% fetal bovine serum, 1% penicillin, and 1% streptomycin, and then incubated with 5% CO2 at 37°C.
2.13 Assessment of the cytotoxic effect of MCP and MTX using MTT assay
An in vitro toxicological assay kit (Sigma Aldrich, Inc., M-5655) which is MTT-based was utilized to assess MTX, MCP, and their combination cytotoxicity on MCF7, Nalm6, and JEG3 cell lines. The cancer cells were seeded into a 96-well plate containing 100 μL of the culture media at 1 × 104 cells/mL density and incubated for 1 day. Afterwards, the culture media was exchanged with 100 μL of a new media with different concentrations of the tested drugs; MTX and MCP (0.4, 1.6, 6.3, 25, 100 μg/mL) and vehicle control, 0.01% dimethyl sulfoxide (DMSO), for 48 h. Each well of the plates was incubated for another 4 hours with 20 μL of MTT in 200 μL of media. Eliza microplate reader (BDR206, Bioline Technology, India) was used for optical density determination, with a 570 nm absorbance. Each concentration of each drug was done in triplicate wells for assessment of IC50 (the concentration required to inhibit 50% of cell growth) of MTX and MCP. After the assessment of the IC50 of each drug for each cell line, another experiment was carried out. The IC50 of MTX was added in combination with that of MCP to the three types of cancer cells for measuring the percentage of cell viability of each cell line to study the effect of MCP on MTX cytotoxicity.
2.14 Statistical analysis
Results representation was done as mean ± standard error of the mean (SEM). One-way analysis of variance (ANOVA) was performed to figure out the statistically significant differences. Thereafter, the comparison between the means of all groups was done using the Tukey-Kramer post-analysis test. A P-value below 0.05 was deemed significant by using Version 9.00 for Windows of GraphPad Prism® (GraphPad Software, United States of America, https://www.graphpad.com/).
4 Discussion
Despite being a commonly prescribed chemotherapeutic and immunosuppressant agent (Pivovarov and Zipursky, 2019; Koźmiński et al., 2020), MTX use is limited due to several adverse effects such as liver and lung toxicity which represent a major clinical challenge (Kim et al., 2009; Ezhilarasan, 2021). For the first time, we reported the potential protective effects of MCP against MTX-elicited liver and lung toxicity in rats as evidenced by improving markers of liver and lung functions and restoring normal liver and lung structure. Mitigating oxidative stress, inflammation, fibrosis, and apoptosis might also contribute to MCP’s therapeutic impact.
Consistent with previous experimental and clinical studies (Ali et al., 2014; Mori et al., 2018; Cao et al., 2019; Karlsson Sundbaum et al., 2019; Roghani et al., 2020), MTX-induced hepatotoxicity was presented through a pronounced elevation in serum ALT and AST levels. Besides the damaged liver histological structure; disorganized hepatic cords, hepatocyte ballooning degeneration with nuclear pyknosis, and hepatic apoptosis (Mahmoud et al., 2017a; Al Kury et al., 2020). The elevated serum liver enzymes may be ascribed to hepatocellular degeneration, loss of hepatocytes structural integrity, and leak of their contents into the blood (McGill, 2016; Rizk et al., 2018).
The MCP hepatoprotective effect against MTX was verified here by the decline in the elevated serum ALT and AST levels alongside the improved liver architecture. As reported before (Kelleni et al., 2016; Kalantari et al., 2019), MTX led to hepatomegaly in rats which was attenuated, here, by MCP. MCP hepatoprotective influence was reported in CCl4-induced liver fibrosis in rats (Abu-Elsaad and Elkashef, 2016).
Regarding MTX-caused pulmonary toxicity, the relative lung weight and lung W/D weight ratio in addition to total protein content and leukocyte count in BALF were notably elevated in the MTX group which are features of acute lung injury (Abraham, 2003; Poitout-Belissent et al., 2021) consistent with previous studies (Rajizadeh et al., 2023; Rajizadeh et al., 2024). The BALF analysis of patients with MTX-induced pneumonitis revealed lymphocytosis and elevated neutrophil counts (D'Elia, 2014). Lymphocyte proliferation and hypersensitivity pneumonitis triggered by alveolitis are linked to cellular immune response and cytokine release (Kim et al., 2009). The histopathological findings showed lymphocytes and macrophage infiltration, disrupted lung architecture, thickened inter-alveolar septa, and capillary congestion as reported before (Arpag et al., 2018; Ozmen et al., 2024).
Here, MCP ameliorated lung edema indices, BALF total protein content, and leukocyte count, and improved the histological architecture, suggesting its protective effects against MTX-induced lung toxicity.
Despite no obvious mechanism illustrating MTX-induced organ dysfunction, accumulation of MTX polyglutamate inside hepatocytes, the metabolized form of MTX, has been reported as the key factor of MTX-induced hepatotoxicity. It decreases the folic acid reservoir which consequently induces several pathological events associated with oxidative stress, inflammation, apoptosis, and fibrosis (Hawwa et al., 2015; Yamamoto et al., 2016).
The impairment of the tissue oxidant/antioxidant balance remains the hallmark cause of MTX-induced organ toxicity resulting in oxidative damage (Gao and Horie, 2002; Huang et al., 2005; Chang et al., 2013). This oxidative damage of protein and DNA in addition to lipid peroxidation causes disarrangement of the lipid bilayer membrane, deactivation of membrane-bound receptors and enzymes, and, in turn, enhances tissue permeability (Halliwell and Gutteridge, 2015; Malayeri et al., 2022) which can explain the elevated liver enzymes with MTX (Dalaklioglu et al., 2013).
The pathogenesis of MTX-induced hepatotoxicity (Cetin et al., 2008; Çakır et al., 2011; Pınar et al., 2018; Kalantar et al., 2019) and lung toxicity (Kurt et al., 2015; Saygin et al., 2016; Arpag et al., 2018) is known to involve oxidative stress. An elevation in MDA levels, a marker of lipid peroxidation, and a decrease in GSH levels as well as inhibition of SOD antioxidant activity, in liver and lung tissues were demonstrated in the MTX group consistent with earlier studies (Demiryilmaz et al., 2012; Mohamed et al., 2019; Hussein et al., 2020; Goudarzi et al., 2021; Zaki et al., 2021; Parthasarathy and Prince, 2023; Abdalhameid et al., 2024). GSH reduction was explained previously by MTX inhibitory effect on cytosolic reduced nicotinamide adenine dinucleotide phosphate (NADPH) (Vardi et al., 2010) which is required for GSH maintenance by glutathione reductase (Ali et al., 2017).
Conversely, MCP significantly rebalanced the oxidative status in both tissues, indicating its antioxidant properties. The antioxidant activity of MCP was revealed in various animal studies including diabetes-associated cognitive impairment (Yin et al., 2020), CCl4-induced liver fibrosis (Abu-Elsaad and Elkashef, 2016), doxorubicin-induced cardiotoxicity (Tian et al., 2020), diabetes-induced nephropathy (Mahmoud et al., 2024), diet-induced obesity (Marín-Royo et al., 2018), and in vitro model of mouse monocytes (Ramachandran et al., 2017).
The master and emerging regulator of cellular antioxidant defense, Nrf2, induces the transcription of antioxidant enzymes and enzymes involved in GSH and NADPH regeneration (Ma, 2013). Although Nrf2 is activated during oxidative stress, the generation of huge amounts of reactive oxygen species (ROS) suppresses its expression (Mahmoud et al., 2017a; Mahmoud et al., 2017b). In agreement with other studies (Mukherjee et al., 2013; Bu et al., 2018; Fayez et al., 2018; Kawami et al., 2022), MTX greatly downregulated Nrf2 expression in hepatic and pulmonary tissues parallel to the findings of MDA, SOD, and GSH. Conversely, MCP prevented Nrf2 downregulation which, subsequently, reduced oxidative stress and improved antioxidant defense. These antioxidant effects of MCP can play a significant role in its protective effects against MTX-induced hepatic and pulmonary toxicity.
Galectin-3 is pivotal in fibrosis and inflammation (Mackinnon et al., 2012; An et al., 2021; Slack et al., 2021; Boutin et al., 2022; Lima et al., 2023). As a proinflammatory protein, Gal-3 initiates and amplifies acute inflammatory response through the recruitment of macrophages to the injury site and perpetuating a chronic inflammatory state through the induction of proinflammatory pathways (Bouffette et al., 2023). The embroilment of inflammation was documented in the pathogenesis of MTX-induced liver (Ali et al., 2017; Mahmoud et al., 2017a; Al Kury et al., 2020) and lung (Mammadov et al., 2019; Zaki et al., 2021; Ozmen et al., 2024) toxicity. This study aimed to demonstrate the significance of Gal-3, as a therapeutic target, in MTX-induced liver and lung toxicity, which has yet to be explored, using MCP as a natural Gal-3 inhibitor.
Galectin-3 was reported as an endogenous paracrine ligand and activator of TLR-4 inducing an inflammatory response (Burguillos et al., 2015). TLR-4, a member of the pattern recognition receptors, is an important sensor of the innate immune response that can interact with exogenous molecules such as lipopolysaccharide (LPS) of Gram-negative bacteria which are recognized as pathogen-associated molecular patterns. Additionally, it can be triggered by endogenous molecules of damaged or necrotic cells such as heat shock protein after oxidative stress which are recognized as damage-associated molecular patterns (Lu et al., 2008; Gill et al., 2010). This interaction eventually leads to an inflammatory cascade through the activation of NF-κB and elevated transcription of proinflammatory cytokines such as TNF-α (Miller et al., 2005).
Recently, the contribution of the TLR-4/NF-κB signaling pathway in MTX-induced liver toxicity was documented (Matouk et al., 2022; Manna et al., 2023). To the best of our knowledge, its importance in MTX-induced lung toxicity has yet to be established. Here, high expression of Gal-3 along with TLR-4/NF-κB/TNF-α signaling pathway upregulation was observed in the hepatic and lung tissues of the MTX group. The activation of TLR-4/NF-κB signaling in LPS-caused injury in chondrocytes mediated the inflammatory and proapoptotic actions of Gal-3 while Gal-3 silence resulted in apoptosis inhibition through inhibition of the inflammatory response (Wang et al., 2019).
Crosstalk between TLR-4 and oxidative stress has also been reported. ROS may lead to TLR-4 activation which, in turn, may increase ROS production through direct interaction between NADPH oxidase and TLR-4 (Gill et al., 2010). Reports discussing relation between NF-κB and Nrf2 revealed that Nrf2 is a negative regulator of NF-κB signaling dampening NF-κB activation as well as limiting the transcription and overproduction of proinflammatory cytokines (Ahmed et al., 2017). Moreover, Nrf2 activation may mitigate TLR-4-induced inflammation under pathological conditions (Huang et al., 2014; Marinovic et al., 2015). This crosstalk may also be a result of the Nrf2/NF-κB relationship (Wardyn et al., 2015; Mohan and Gupta, 2018). In a model of ischemic-reperfusion liver injury, it was stated that induction of Nrf2 attenuated TLR-4-induced liver inflammation and ameliorated oxidative stress (Huang et al., 2014).
We found that the MCP-induced blockade of Gal-3 protected against MTX-induced liver and lung toxicity by suppressing TLR-4 and its downstream regulator, NF- κB. Hence, one of the critical mediators of inflammation and apoptosis was decreased, TNF-α. Previous studies indicated that the MCP-induced Gal-3 inhibition exhibited anti-inflammatory effects in animal models analyzing erectile dysfunction (Wang et al., 2024), cerebral-ischemia reperfusion injury (Cui et al., 2022), and myocardial fibrosis (Xu et al., 2020) via the downregulation TLR-4/NF-κB signaling pathway. Collectively, the inhibition of the TLR-4/NF-κB/TNF-α signaling by MCP could be explained by both inhibition of Gal-3 and activation of Nrf2. The anti-inflammatory effects of MCP can explain, to a certain extent, its protective effects against MTX toxicity.
Unresolved inflammation and abnormal tissue repair can result in tissue fibrosis (Bouffette et al., 2023). As a profibrotic protein, Gal-3 is identified as a biomarker for the progression of fibrosis (Li et al., 2014) and its expression was increased in the bleomycin-caused pulmonary fibrosis mouse model and patients with idiopathic pulmonary fibrosis (Nishi et al., 2007; Mackinnon et al., 2012) or liver fibrosis (Mackinnon et al., 2023). MTX-elicited fibrosis in the liver (Taskin et al., 2017; Cao et al., 2019; Ahmad et al., 2021) and lung (Saygin et al., 2016; Abdalhameid et al., 2024; Manie et al., 2024) tissues were identified. In this work, MTX led to elevated expression of TGF-β, a profibrotic cytokine. TGF-β stimulation eventually activates tissue fibroblasts into active myofibroblasts leading to extracellular matrix synthesis (Biernacka et al., 2011; Meng et al., 2016). TGF-β is significant in MTX-induced pulmonary fibrosis which is at least partially mediated by epithelial-mesenchymal transition (EMT) at which myofibroblasts originate from the injured epithelial cells (Ohbayashi et al., 2014). Moreover, MTX-induced downregulation and reduced activity of Nrf-2 are also involved in MTX-caused EMT in alveolar epithelial cell lines (Kawami et al., 2022). Gal-3 has an important role in the regulation of EMT induction (Mackinnon et al., 2012).
Gal-3 is mandatory for TGF-β-induced myofibroblast activation and extracellular matrix production (Henderson et al., 2006). Gal-3 has recently been shown to activate TGF-β in human pulmonary fibroblasts and its inhibition can prevent TGF-β activation (Calver et al., 2024). Moreover, it was identified that the pharmacological inhibition of Gal-3 in the NAFLD mice model downregulated TGF-β (Lee et al., 2022). MCP counteracted the profibrotic effects of Gal-3 in different disease models (Kolatsi-Joannou et al., 2011; Calvier et al., 2013; Martínez-Martínez et al., 2015; Vergaro et al., 2016; Li et al., 2018; Ibarrola et al., 2019; Yin et al., 2020). Consistent with previously mentioned findings, MCP-induced Gal-3 inhibition downregulated hepatic and pulmonary TGF-β in MTX-treated rats. So, the inhibition of the fibrotic Gal-3/TGF-β pathway by MCP can protect against MTX-induced liver and lung fibrosis.
Hepatic and pulmonary fibrosis induced by MTX were additionally confirmed by Masson’s trichrome staining which showed the proliferation of fibrous tissue, as previously reported (Tag, 2015; Mohamed et al., 2019; Abdalhameid et al., 2024). However, MCP treatment decreased collagen deposition in both liver and lung tissues in line with the results of Gal-3 and TGF-β. Similarly, MCP decreased collagen content demonstrated by Masson’s trichrome staining in the cisplatin-induced nephrotoxicity model (Li et al., 2018).
Several studies reported that the induction of apoptosis may mediate MTX-induced liver (Mahmoud et al., 2017a; Khafaga and El-Sayed, 2018; Türk et al., 2022) and lung (Kurt et al., 2015; Abosrea et al., 2023; Ozmen et al., 2024) damage. Consistent with previous reports (Ali et al., 2014; Rajizadeh et al., 2024), MTX caused c-caspase-3 overexpression in both examined tissues which can be related to the elevated levels of ROS and proinflammatory cytokines (Simon et al., 2000; Jaeschke, 2011). It is worth mentioning that Gal-3 may have a function in regulating apoptosis. It may act as an antiapoptotic factor due to its sequence homology to B-cell lymphoma 2 (Bcl-2), an apoptosis suppressor (Yang et al., 1996). However, a previous study showed that extracellular Gal-3 acts as a proapoptotic factor triggering apoptosis in activated T-cells leading to mitochondrial apoptosis involving the release of cytochrome c and activation of caspase-3 (Fukumori et al., 2003). In the present, MCP showed antiapoptotic effects through downregulating c-caspase-3 which may result from the Gal-3 inhibition, anti-inflammatory and antioxidant influences of MCP parallel to earlier studies reporting the antiapoptotic properties of MCP (Abu-Elsaad and Elkashef, 2016; Li et al., 2018; Tian et al., 2020; Mahmoud et al., 2024).
Galectin-3 has a tumor-promoting effect in different tumors (Eliaz and Raz, 2019). It promotes cancer cell resistance to chemotherapeutic agents acting as a potent inhibitor of the intrinsic apoptosis pathway (Nakahara et al., 2005; Fukumori et al., 2007; Navarro et al., 2020). Several previous reports have shown that MCP, through Gal-3 inhibition, modulates multiple rate-limiting steps of cancer metastasis (Glinsky and Raz, 2009; Eliaz and Raz, 2019). The anticancer activity of MCP was demonstrated before such as in the colon (Liu et al., 2008; Wu et al., 2018), prostate (Yan and Katz, 2010), ovarian (Hossein et al., 2013), and breast (Garrido et al., 2024) cancers. According to these findings, MCP may be used as a natural chemosensitizer with chemotherapeutic agents (Chauhan et al., 2005; Johnson et al., 2007). Accordingly, our findings of the MTT cell viability assay showed that both MTX and MCP significantly reduced, in a concentration-dependent way, the cell viability of MCF7, Nalm6, and JEG3 at which the expression of Gal-3 in these cell lines was previously documented (Simone et al., 2014; Jovanović et al., 2024; Li et al., 2024). Moreover, combining MCP (IC50) with MTX (IC50) enhanced MTX cytotoxicity efficacy by decreasing cell viability by less than 50% compared to MTX alone in Nalm6 and JEG3 cells. The MCP-induced inhibition of Gal-3 could demonstrate this.
In conclusion, this study demonstrates that MCP protects against MTX-caused hepatic and pulmonary toxicity through anti-inflammatory, antiapoptotic, antifibrotic, and antioxidant properties. The antioxidant effects are evidenced by the upregulation of Nrf2 expression, a decrease in MDA levels, and an increase in SOD activity and GSH levels. Moreover, MCP downregulated the inflammatory signaling pathway Gal-3/TLR-4/NF-κB pathway. Additionally, MCP decreased c-caspase-3, TGF-β, and collagen levels in liver and lung tissues. These effects were reflected in the improved liver and lung functional markers and histological structure. Moreover, the cytotoxicity of MTX was enhanced by MCP in different human cell lines. A limitation of the study is the necessity for in vitro and further in vivo studies to investigate additional protective mechanistic pathways of MCP against MTX-induced hepatic and pulmonary toxicity.
The study outcomes are significant clinically where MTX is commonly prescribed. As a natural product, MCP can be combined with MTX treatment protocols to decrease the incidence and severity of its associated adverse effects thus enhancing the patient’s outcome. Clinical trials are required before practical application to assess efficacy, safety, appropriate dosage, and time required for pretreatment to offer its protective effects.