Irisin Potentiates 5-fluorouracil Cytotoxicity in HT-29 Colon Cancer Cells through Enhanced Apoptosis and Transcriptional Modulation of Adenosine Monophosphate-activated Protein Kinase/Mammalian Target of Rapamycin Signaling

Abstract

Background:

Although 5-fluorouracil (5-FU) is a standard chemotherapeutic agent for colorectal cancer (CRC), its efficacy is often limited by chemoresistance. Irisin, an exercise-induced myokine, has demonstrated anticancer potential; however, its role in CRC remains incompletely defined.

Materials and Methods:

Human HT-29 CRC cells were treated for 12 h with irisin (2, 20, and 200 ng/mL) alone or combined with 5-FU (50 μM). Cell cycle progression and apoptosis were assessed by flow cytometry. Expression of cell cycle regulators (CCND1, CCND3, p21, and p27), apoptotic markers (BCL2, Bcl-2-associated X protein, cytochrome c, caspase-8, caspase-9, and caspase-3), and metabolic mediators (protein kinase B [Akt], mammalian target of rapamycin [mTOR], adenosine monophosphate-activated protein kinase [AMPK], and phosphatase and tensin homolog [PTEN]) was analyzed by quantitative reverse transcription-polymerase chain reaction.

Results:

While 5-FU induced G0/G1 arrest with p21/p27 upregulation, irisin alone or in combination exerted minimal additional effects on cell cycle progression. In contrast, high-dose irisin (200 ng/mL) significantly increased apoptotic cell fractions and upregulated caspase-8 and caspase-3 without significant alteration of mitochondrial apoptotic gene expression. Combination treatment further enhanced apoptosis. Co-treatment also increased AMPK and PTEN mRNA levels while reducing Akt and mTOR mRNA levels.

Conclusion:

Under the experimental conditions used, irisin potentiated 5-FU cytotoxicity in HT-29 CRC cells primarily through enhanced apoptosis and transcriptional modulation of metabolic signaling components. These findings suggest a potential chemosensitizing role for irisin, warranting further validation at the protein and functional levels.

Introduction

Colorectal cancer (CRC) is a highly prevalent malignancy worldwide, ranking as the third most commonly diagnosed cancer and the second leading cause of cancer-related mortality, with approximately two million new cases and 900,000 deaths reported globally in 2022.[1,2] The increasing incidence of CRC has been strongly associated with modifiable lifestyle-related factors, including physical inactivity, obesity, hyperglycemia, and hyperinsulinemia.[3,4]

At the molecular level, CRC progression is characterized by dysregulated cell cycle control, including upregulation of cyclins (e.g., CCND1 and CCND3) and suppression of cyclin-dependent kinase inhibitors such as p21 and p27.[5,6,7] In parallel, apoptosis evasion is a key hallmark of tumorigenesis. Apoptosis is governed by two main pathways: the intrinsic (mitochondrial) and extrinsic (death receptor-mediated) mechanisms.[8,9] The key molecular regulators include caspase-9, Bcl-2-associated X protein (BAX), and cytochrome c, which all converge on the activation of caspase-3, the key executioner of programmed cell death.[8,9]

Clinically, early-stage CRC is primarily managed by surgical resection, whereas advanced disease relies heavily on 5-fluorouracil (5-FU)-based chemotherapy, which disrupts DNA synthesis, thereby inducing cell cycle arrest and apoptosis.[10,11] However, its therapeutic efficacy is frequently limited by chemoresistance driven by enhanced DNA repair, impaired apoptosis, and activation of survival signaling pathways.[10,11]

Metabolic reprogramming further contributes to CRC progression and drug resistance. CRC cells frequently exhibit increased reliance on aerobic glycolysis (the Warburg effect), promoting proliferation and survival under stress.[12,13,14] This metabolic phenotype is largely mediated through the activation of the protein kinase B (Akt)/mammalian target of rapamycin (mTOR) axis,[15,16] while key negative regulators, including phosphatase and tensin homolog (PTEN) and 5′ adenosine monophosphate-activated protein kinase alpha (AMPKα), are often downregulated.[15,16,17,18] Accordingly, targeting the Akt/mTOR axis and restoring metabolic checkpoints has therefore emerged as a promising strategy to enhance chemosensitivity and overcome 5-FU resistance in CRC.[10,11]

Irisin, an exercise-induced myokine cleaved from fibronectin type III domain-containing protein 5, has gained attention for its metabolic regulatory properties.[19,20,21] Irisin interacts with the components of the AMPK/Akt/mTOR signaling network, suggesting a potential role in modulating cancer cell metabolism and survival.[19,20,21,22] Clinical evidence has reported altered circulating irisin levels in CRC patients,[23] and in vitro studies in various malignancies have demonstrated anti-proliferative and pro-apoptotic effects for the myokine.[24,25,26,27] Nevertheless, its direct impact on CRC cell cycle regulation, apoptosis, and chemotherapy responsiveness remains incompletely characterized.

Therefore, the present study investigated whether irisin modulates cell cycle progression, apoptosis, and transcriptional components of the AMPK/Akt/mTOR pathway in HT-29 CRC cells, which harbor a mutant p53 phenotype associated with reduced sensitivity to 5-FU. Furthermore, we examined whether irisin enhances 5-FU cytotoxicity, aiming to clarify its potential role as a metabolic modulator and chemosensitizing agent in CRC.

Materials and Methods

Chemicals and reagents

The human HT-29 colon cancer cell line (ATCC^® HTB-38™) was obtained from the American Type Culture Collection (ATCC, Virginia, USA). Recombinant, biologically active irisin protein (Cat. #HY-P70664) was sourced from MedChemExpress LLC (Princeton, NJ, USA) and 5-FU (≥95% purity) was purchased from Hospira Australia Ltd (Melbourne, Australia).

Routine cell culture was performed using GlutaMAX™ RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS) and 1% antibiotic–antimycotic solution (Thermo Fisher Scientific, Waltham, MA, USA). Additional cell culture reagents and consumables included sterile 96-well and 6-well plates, propidium iodide (PI), RNase A, the MTT Cell Viability Assay Kit (#M6494), and the Annexin V/Alexa Fluor 488 and PI Apoptosis Detection Kit (#V13245), also obtained from Thermo Fisher.

For gene expression analysis, PureLink RNA Mini Kit (#12183025), High-Capacity cDNA Reverse Transcription Kit (#4374967), Power SYBR™ Green PCR Master Mix (#4368708), MicroAmp™ Fast Optical 96-Well Plates (#4346907), and Optical Adhesive Film (#4311971), were purchased from Thermo Fisher Scientific. Primers [Table 1] were procured from Integrated DNA Technologies (IDT, Leuven, Belgium).

Table 1.

Sequences of polymerase chain reaction primers used for detecting human GAPDH, CCND1, CCND3, CDKN1A, CDKN1B, BCL2, Cytochrome c, BAX, Caspase-8, Caspase-9, Caspase-3, AKT1, mTOR, PTEN, and AMPK-α (PRKAA1) mRNAs in human HT-29 coloncancer cell including their corresponding gene accession numbers and amplicon sizes

Genes	Forward	Reverse	Amplicon size (bp)
GAPDH (NCBI: NM_002046.5)	5’-CACATGGCCTCCAAGGAGTAA-3’	5’-TGA GGGTCCTCTCTTCCTCTTGT-3’	74
CCND1 (NCBI: NM_053056.2)	5’-TGACCCCGCACGATTTCATT-3’	5’-CATGGAGGGCGGATTGGAAA-3’	143
CCND3 (NCBI: NM_001136017.3)	5’-GGTGCAATCCTCTCCTCG C-3’	5’-TAGTTCATGGCCAGGGGGAA-3’	183
CDKN1A (NCBI:NM_000389.4)	5’-AGTCAGTTCCTTGTGGAGCC-3’	5’-GCATGGGTTCTGACGGACAT-3’	109
CDKN1B (NCBI: NM_004064.4)	5’-CTGGCCTCAGAAGACGTCAAA-3’	5’-AGGATGTCCATTCCATGAAGTCAG-3’	147
BCL2 (NCBI: NM_000633.3)	5’-TTT CGGTGACTTCCGCATCA-3’	5’-CGGTCTCCTAAAAGCAGGCA-3’	79
CYCS (NCBI: NM_018947.6)	5’-CGTTGTGCCAGCGACTAAAA-3’	5’-TGGCACTGGGAACACTTCAT-3’	88
BAX (NCBI: NM_001291428.1)	5’-TCGCCCTTTTCTACTTTGCCA-3’	5’-GTCCTGGAGACAGGGACATCA-3’	195
Caspase-8 (NCBI: NM_001228.4)	5’-CTGTACCTTTCTGGCGGAGG-3’	5’-TCCTTCTCCCAGGATGACCC-3’	200
Caspase-9 (NCBI: NM_001229.5)	5’-CAGGCCCCATATGATCGAGG-3’	5’-GGCCTGTGTCCTCTAAGCAG-3’	140
Caspase-3 (NCBI:NM_004346.3)	5’-CTC TGGTTTTCGGTGGGTGT-3’	5’-CCACTGAGTTTTCAGTGTTCTCC-3’	90
AKT1 (NCBI: NM_001382431.1)	5’-CTCAGTGTCGTCAGAGCCC-3’	5’-ATGGAAAGCAGGCCAGACTC-3’	100
mTOR (NCBI: NM_004958.4)	5’-GACGAGAGATCATCCGCCAG-3’	5’-ACAAGGGACCGCACCATAAG-3’	97
PTEN (NCBI: NM_000314.8)	5’-CTC AGCCGTTACCTGTGTGT-3’	5’-AGGTTTCCTCTGGTCCTGGT-3’	129
PRKAA1 (NCBI: NM_006251.6)	5’-ATGCGCAGACTCAGTTCCTG-3’	5’-CTTCACTTTGCCGAAGGTGC-3’	126

CYCS: Cytochrome c

Cell culture and cell cycle analysis

HT-29 colon cancer cells, which harbor a mutant p53 phenotype, were routinely cultured in RPMI-1640 medium supplemented with 10% FBS and 1% antibiotic-antimycotic. Cells were incubated at 37°C in 5% CO₂ under the humidified conditions. A previously established IC₅₀ concentration of 5-FU (50 μM), determined after 24 h of treatment, was selected for use in all relevant experiments.[28,29] For irisin treatments, three concentrations were applied: 2 ng/mL (IR2), 20 ng/mL (IR20), and 200 ng/mL (IR200).

The irisin concentrations applied in this study (2, 20, and 200 ng/mL) were selected based on previously published in vitro investigations evaluating irisin-mediated modulation of cancer cell signaling and apoptosis.[20] Moreover, reported circulating irisin levels in humans vary substantially depending on physiological status and detection methods.[30,31] In light of this variability, the lower concentrations (2 and 20 ng/mL) were selected to approximate reported systemic levels, whereas the higher dose (200 ng/mL) was included to evaluate potential dose dependent and pharmacological effects in a controlled in vitro setting.[20]

A 12-h exposure period was used for all monotherapy and combination treatments, consistent with prior studies assessing the early effects of 5-FU-based therapies on cell cycle progression, apoptosis, and expression of key regulatory genes.[28,29] This time frame also enables the evaluation of early transcriptional modulation of apoptosis- and metabolism-related genes before the development of extensive secondary cytotoxic effects that may complicate interpretation of primary regulatory events.[32,33] Accordingly, analysis at the 12-h timepoint allows the assessment of early mechanistic responses rather than late-stage transcriptional changes associated with advanced cell death.[34]

HT-29 cells were seeded into 6-well plates and allowed to adhere for 24 h before treatment. Experimental groups included untreated control cells (CT), monotherapies with 5-FU, IR2, IR20, IR200, and combination regimens of 5-FU with IR2 (IF2), IR20 (IF20), and IR200 (IF200).

After treatment, cells were trypsinized, washed twice with PBS, and incubated with RNase A (20 μg/mL) for 15 min at the room temperature. Following further PBS washes, PI (2 μg/mL) was added immediately before flow cytometric analysis using the ACEA NovoCyte 3000 system (Agilent Technologies, CA, USA). Data acquisition involved 20,000 events per sample, and cell cycle distribution was determined using NovoExpress software, following standard algorithms for DNA content analysis.[28,29] Data were reported as mean ± standard deviation (SD) from three biological replicates.

Cell apoptosis analysis

Apoptotic response was evaluated using the Annexin V/Alexa Fluor 488 and PI Apoptosis Kit, according to the manufacturer’s protocol. HT-29 cells (treated and control) were washed twice in cold PBS and resuspended in 100 μL of Annexin V binding buffer. Next, 5 μL of Annexin V–Alexa Fluor 488 and 1 μL of PI were added. After a 15-min dark incubation at the room temperature, 400 μL binding buffer was added and samples were analyzed using the NovoCyte 3000 cytometer. Apoptotic populations were classified based on dual staining into viable (unstained), early apoptotic (Annexin V⁺/PI−), late apoptotic (Annexin V⁺/PI⁺), and necrotic or dead (Annexin V⁻/PI⁺) cells. Data were shown as mean ± SD from three biological replicates.[28,29]

Gene expression

Total RNA was extracted using the PureLink RNA Mini Kit, as per the manufacturer’s instructions. Synthesis of cDNA was performed with the High-Capacity Reverse Transcription Kit. Quantitative real-time polymerase chain reaction (qPCR) was carried out using SYBR Green Master Mix on the QuantStudio™ 3 System (Thermo Fisher Scientific). Each 10 μL reaction contained 3 μL cDNA, 5 μL SYBR Green Master mix, and 1 μL each of forward and reverse primers. The amplification cycle included 40 cycles of 95°C for 15 s and 60°C for 1 min.

No-template and no- reverse-transcription (RT) controls were included to verify the absence of contamination or genomic DNA. Expression levels of genes including CCND1, CCND3, p21, p27, BCL2, BAX, Cytochrome c, Caspase-3, Caspase-8, Caspase-9, AKT1, mTOR, AMPK-α, and PTEN were normalized to GAPDH, and fold change was calculated using the 2^−∆∆Ctmethod.[35] Primer specificity was verified by melt curve analysis, demonstrating single amplification products. Amplification conditions were optimized to ensure consistent efficiency across target and reference genes, supporting the use of the 2^−∆∆Ct method for relative quantification.

Statistical analysis

Statistical analyses were performed using the SPSS software version 25 (IBM Corp., Armonk, NY, USA). Data distribution normality was evaluated using the Kolmogorov–Smirnov test, while Levene’s test assessed homogeneity of variances. Intergroup comparisons were conducted using the one-way ANOVA, followed by Tukey’s HSD or Games–Howell post hoc tests, depending on variance equality. Results were expressed as mean ± SD. P <0.05 was considered statistically significant.

Results

Effects of 5-fluorouracil and/or irisin on cell cycle and expression of cell cycle-regulatory genes

Compared with untreated controls, 5-FU monotherapy significantly increased the HT-29 Sub-G1 cell population (1.5-fold; P < 0.0001), which commonly reflects DNA fragmentation associated with cell death [Figure 1A]. Treatment with IR20 and IR200 also significantly elevated the Sub-G1 fraction, whereas IR2 had no significant effect. However, co-treatment regimens (IF2, IF200) did not exceed the effect observed with 5-FU monotherapy.

Figure 1.

Distribution of HT-29 CRC cells across different cell cycle phases following 12h treatment with 5-fluorouracil (5-FU; 50 μM) and/or irisin at 2 ng/mL (IR2), 20 ng/mL (IR20), and 200 ng/mL (IR200), administered as monotherapies or in combination (IF2, IF20, IF200). Cell cycle was assessed by flow cytometry following propidium iodide staining, with quantification of: (A) Sub-G1 phase, (B) G0/G1-phase, (C) S-phase, and (D) G2/M-phase. Data are presented as mean ± standard deviation (SD) from three independent experiments. Statistical analysis was performed using one-way ANOVA with Tukey’s post hoc test (statistical significance: a = P < 0.05 compared to untreated cells; b = P < 0.05 compared to 5-FU group; c = P < 0.05 compared to IR2 group, d = P < 0.05 compared to IR20 group; e = P < 0.05 compared to IR200 group; f = P < 0.05 compared to IF2 group, and g = P < 0.05 compared to IF20 group).

Analysis of cell cycle distribution demonstrated significant G0/G1 phase accumulation following 5-FU treatment [1.7-fold; Figure 1B], accompanied by corresponding reductions in S phase [Figure 1C] and G2/M phase [Figure 1D] populations. Irisin monotherapy induced comparatively minor changes in cell cycle distribution. Although combination treatments significantly increased G0/G1 arrest compared to untreated cells, the magnitude of this effect remained comparable to that of 5-FU alone.

At the transcriptional level, CCND1 expression remained unchanged across all treatment conditions [Figure 2a]. In contrast, CCND3 was significantly downregulated in response to 5-FU alone and in all combination groups [Figure 2b]. Expression of p21 [Figure 2c] and p27 [Figure 2d] was markedly upregulated following 5-FU monotherapy (P < 0.0001). Irisin monotherapy had no significant effect on these genes; however, co-treatment with 5-FU further enhanced p21 and p27 mRNA expression compared with untreated controls.

Figure 2.

Relative gene expression levels of cell cycle regulators (a) CCND1, (b) CCND3, (c) p21, and (d) p27 assessed by quantitative reverse transcription-polymerase chain reaction following 12 h treatment with 5-fluorouracil (5-FU; 50 μM) and/or irisin at 2 ng/mL (IR2), 20 ng/mL (IR20), and 200 ng/mL (IR200), administered either as monotherapies or in combination (IF2, IF20, IF200). Gene expression was normalized to GAPDH and presented as mean ± standard deviation from three independent experiments. Statistical analysis was performed using one-way ANOVA with Tukey’s post hoc test (statistical significance: a = P < 0.05 compared to untreated cells; b = P < 0.05 compared to 5-FU group; c = P < 0.05 compared to IR2 group, d = P < 0.05 compared to IR20 group; e = P < 0.05 compared to IR200 group; f = P < 0.05 compared to IF2 group, and g = P < 0.05 compared to IF20 group). 5-FU: 5-fluorouracil

Effects of 5-fluorouracil and/or irisin on cell apoptosis and expression of apoptosis-related genes

Monotherapy with 5-FU significantly reduced viable HT-29 cell numbers and increased both early and late apoptotic populations relative to untreated controls [Figure 3; P < 0.0001]. IR2 and IR20 monotherapies produced minimal apoptotic effects, whereas IR200 significantly increased apoptotic cell fractions and reduced viability (P < 0.0001). Notably, IR200 monotherapy induced a stronger apoptotic response than 5-FU alone (P < 0.001). Although the IF200 co-treatment further reduced cell viability compared with 5-FU monotherapy, the magnitude of apoptosis did not exceed that observed with IR200 alone [Figure 3].

Figure 3.

Percentage distribution of HT-29 colorectal cancer cells across different apoptotic stages following 12-h treatment with 5-fluorouracil (5-FU; 50 μM) and/or irisin at 2 ng/mL (IR2), 20 ng/mL (IR20), and 200 ng/mL (IR200), administered either as monotherapies or in combination (IF2, IF20, IF200). Apoptosis was assessed using Annexin V/Alexa Fluor 488 and PI staining, with flow cytometric quantification of: (a) viable cells, (b) early apoptotic cells, (c) late apoptotic cells, and (d) dead cells. Data represent mean ± standard deviation from three independent experiments. Statistical analysis was performed using one-way ANOVA followed by Tukey’s post hoc test (statistical significance: a = P < 0.05 compared to untreated cells; b = P < 0.05 compared to 5-FU group; c = P < 0.05 compared to IR2 group, d = P < 0.05 compared to IR20 group; e = P < 0.05 compared to IR200 group; f = P < 0.05 compared to IF2 group, and g = P < 0.05 compared to IF20 group). 5-FU: 5-fluorouracil

At the gene expression level, BCL2 was significantly downregulated following 5-FU and IR200 monotherapies, with the most pronounced reduction observed in the IF200 combination group [Figure 4a]. In contrast, expression levels of BAX, cytochrome c, and caspase-9 remained largely unchanged across treatment conditions [Figure 4b-d]. However, caspase-8 and caspase-3 transcripts were significantly upregulated in several treatment groups, with the highest expression observed in the IF200 combination [Figure 4e and f].

Figure 4.

Relative gene expression levels of cell apoptosis regulators (a) BCL2, (b) Bcl-2-associated X protein, (c) Cytochrome c, (d) Caspase-9, (e) Caspase-8, and (f) Caspase-3 assessed by quantitative reverse transcription-polymerase chain reaction following 12 h treatment with 5-fluorouracil (5-FU; 50 μM) and/or irisin at 2 ng/mL (IR2), 20 ng/mL (IR20), and 200 ng/mL (IR200), administered either as monotherapies or in combination (IF2, IF20, IF200). Gene expression was normalized to GAPDH and presented as mean ± standard deviation from three independent experiments. Statistical analysis was performed using one-way ANOVA with Tukey’s post hoc test (statistical significance: a = P < 0.05 compared to untreated cells; b = P < 0.05 compared to 5-FU group; c = P < 0.05 compared to IR2 group, d = P < 0.05 compared to IR20 group; e = P < 0.05 compared to IR200 group; f = P < 0.05 compared to IF2 group, and g = P < 0.05 compared to IF20 group). 5-FU: 5-fluorouracil. 5-FU: 5-fluorouracil

Effects of 5-fluorouracil and/or irisin on adenosine monophosphate-activated protein kinase/mammalian target of rapamycin pathway gene expression

Monotherapy with either 5-FU or varying concentrations of irisin had minimal impact on Akt [Figure 5a] and mTOR [Figure 5b] mRNA expression. In contrast, co-treatment with 5-FU and the highest irisin concentration (IF200) resulted in marked downregulation of both Akt and mTOR transcripts compared with other treatment groups.

Figure 5.

Relative gene expression levels of (a) protein kinase B, (b) mammalian target of rapamycin, (c) adenosine monophosphate-activated protein kinase alpha, and (d) phosphatase and tensin homolog assessed by quantitative reverse transcription-polymerase chain reaction following 12 h treatment with 5-fluorouracil (5-FU; 50 μM) and/or irisin at 2 ng/mL (IR2), 20 ng/mL (IR20), and 200 ng/mL (IR200), administered either as monotherapies or in combination (IF2, IF20, IF200). Gene expression was normalized to GAPDH and presented as mean ± standard deviation from three independent experiments. Statistical analysis was performed using one-way ANOVA with Tukey’s post hoc test (statistical significance: a = P < 0.05 compared to untreated cells; b = P < 0.05 compared to 5-FU group; c = P < 0.05 compared to IR2 group, d = P < 0.05 compared to IR20 group; e = P < 0.05 compared to IR200 group; f = P < 0.05 compared to IF2 group, and g = P < 0.05 compared to IF20 group). 5-FU: 5-fluorouracil

Similarly, none of the monotherapy protocols significantly altered AMPK-α [Figure 5c] or PTEN [Figure 5d] expression. However, the IF200 co-treatment group demonstrated the highest mRNA expression levels of AMPK-α and PTEN relative to other treatments.

Discussion

This study assessed the anticancer potential of irisin, both as a monotherapy and in combination with 5-FU, in HT-29 human CRC cells. The main objective was to investigate whether irisin could enhance 5-FU cytotoxicity by promoting cell cycle arrest and apoptosis alongside modulating the AMPK/mTOR pathway, which is commonly implicated in chemoresistance.

Consistent with established mechanisms, 5-FU monotherapy induced G0/G1-phase arrest, downregulated CCND3, and significantly upregulated p21 and p27, confirming effective inhibition of the G1/S transition.[36,37] In contrast, treatment with various irisin concentrations, alone or combined with 5-FU, exerted negligible effects on cell cycle distribution and expression of cell cycle regulatory genes.

Currently, limited and conflicting evidence exists on the role of irisin in CRC. While a clinical study reported markedly lower serum irisin levels in CRC patients compared to healthy controls,[23] another demonstrated increased irisin protein expression in malignant colonic specimens relative to adjacent noncanceroussites.[38] To our knowledge, this is the first study to evaluate the effect of irisin, alone and combined with chemotherapy, on cell cycle progression and its regulatory genes in CRC. The present findings demonstrate that irisin, either as monotherapy or combined with 5-FU, did not significantly alter cell cycle distribution in HT-29 cells beyond the effects induced by 5-FU alone. These results are consistent with previous in vitro studies reporting minimal effects of irisin on proliferation in human HT-29 colon, KLE and RL95-2 endometrial, SW579 thyroid, and OE33 esophageal cancer cell lines.[39] Although cell cycle arrest has been observed at G0/G1 phase in pancreatic[40] and at G2/M phase in glioblastoma[41] cancer cells following irisin treatment, our data indicate that, under the present experimental conditions, irisin does not exert a measurable impact on CRC cell cycle progression.

Despite limited cell cycle modulation, irisin exerted pronounced effects on apoptosis. High-dose irisin (200 ng/mL), alone or combined with 5-FU, significantly increased early and late apoptotic cell populations and upregulated caspase-8 and caspase-3 transcripts. These findings are consistent with prior studies reporting irisin-induced apoptosis in breast,[24] pancreatic,[26] prostate,[42,43] ovarian,[44] and cervical[45] human cancer cells. Moreover, the current data support prior research demonstrating that irisin potentiates chemosensitivity of doxorubicin in breast[24] paclitaxel in lung[46] and gemcitabine in pancreatic[47,48] cancer cells, suggesting a broader chemosensitizing role for the myokine. While these findings support the growing interest in integrating physical exercise or exercise-induced molecules such as irisin into multimodal cancer treatment strategies,[20,21] further mechanistic studies incorporating protein expression and mitochondrial functional assays are necessary to fully characterize its pro-apoptotic effects in CRC.

Interestingly, mitochondrial apoptotic markers (BCL2, BAX, cytochrome c, and caspase-9) remained largely unchanged. While similar patterns have been described in breast[24] and pancreatic[47] cancer cells, other reports showed irisin-induced modulation of mitochondrial pro-apoptotic regulators in pancreatic,[26] prostate,[43] and cervical[45] cancer cell lines. One potential explanation relates to the molecular characteristics of HT-29 cells, which harbor a mutant p53 phenotype (R273H).[49] Given the established role of wild-type p53 in regulating intrinsic mitochondrial apoptosis,[50] the limited modulation of mitochondrial markers observed here may reflect the underlying p53-mutant background.[49,50] Accordingly, extrinsic pathway activation may represent a more prominent apoptotic mechanism in this model. Validation in CRC cell lines with distinct p53 status, together with protein-level and functional apoptosis analyses, is required to determine whether this pathway preference is generalizable.[9,20,21]

In addition to apoptotic modulation, irisin was associated with transcriptional changes in key components of the AMPK/Akt/mTOR axis. High-dose irisin, particularly in combination with 5-FU, increased AMPKα and PTEN mRNA levels while reducing Akt and mTOR transcripts. Previous studies have demonstrated that irisin modulates AMPK and mTOR signaling in several cancer types.[25,26,27,40] Given the established inhibitory roles of AMPK and PTEN in regulating the oncogenic PI3K/Akt/mTOR pathway,[10,11,28,51,52] these findings suggest potential involvement of metabolic regulatory pathways in the enhanced cytotoxic effects of irisin against CRC.[25,26,27,40] However, it is important to emphasize that AMPK, Akt, and mTOR activity is predominantly regulated at the posttranslational level through phosphorylation-dependent mechanisms.[15,16,17,18] Therefore, transcriptional alterations do not confirm functional pathway activation or inhibition. Protein expression analysis and phosphorylation profiling will be necessary to validate whether irisin directly modulates this signaling axis in CRC cells.

This study has several limitations. First, only a single CRC cell line was examined. HT-29 cells harbor a mutant p53 phenotype and exhibit relative resistance to 5-FU compared with p53 wild-type models.[49,50] Given the role of p53 in mitochondrial apoptosis and chemotherapy responsiveness, this genetic background may have influenced the apoptotic signaling profile observed. Validation in additional CRC models with distinct molecular characteristics, including p53 wild-type lines such as HCT116, is therefore required to determine whether irisin-mediated effects are consistent across different genetic contexts.[49,50] Second, all mechanistic conclusions are based on mRNA expression data without protein or functional validation. Third, the study did not include time-course analyses or hypoxic conditions, which may influence irisin bioactivity. Finally, in vivo validation is necessary to determine therapeutic relevance.

Conclusion

The present findings demonstrate that under the current experimental conditions, irisin enhances apoptotic responses and potentiates 5-FU cytotoxicity in p53-mutant HT-29 CRC cells, while exerting minimal influence on cell cycle progression. These effects are associated with transcriptional modulation of apoptotic mediators and PI3K/Akt/mTOR signaling components. Further mechanistic and translational studies are required to establish the broader applicability of irisin as a potential adjunct in CRC therapy.

Ethical clearance

The study was approved by the institutional Ethics Committee of UQU, Saudi Arabia (HAPO-02-K-012-2022-11-1268).

Conflicts of interest

There are no conflicts of interest.

Funding Statement

Nil.