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Article

Roburic Acid as a Therapeutic Candidate: Antiproliferative Activity and Secondary Cell Death Response in Colorectal Cancer Cells

by
Adrianna Gielecińska
1,2,
Mateusz Kciuk
1,*,
Renata Gruszka
1,
Sebastian Wawrocki
3 and
Renata Kontek
1
1
Department of Molecular Biotechnology and Genetics, University of Lodz, Banacha 12/16, 90-237 Lodz, Poland
2
Doctoral School of Exact and Natural Sciences, University of Lodz, Banacha Street 12/16, 90-237 Lodz, Poland
3
Laboratory of Microscopic Imaging and Specialized Biological Techniques, University of Lodz, Banacha Street 12/16, 90-237 Lodz, Poland
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2026, 27(5), 2478; https://doi.org/10.3390/ijms27052478
Submission received: 9 February 2026 / Revised: 4 March 2026 / Accepted: 6 March 2026 / Published: 8 March 2026
(This article belongs to the Special Issue Phytochemicals: Mechanisms and Applications in Cancer Prevention and Therapy)
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Abstract

Natural compounds are increasingly recognized as valuable sources of pharmacologically active agents for cancer therapy. Among them, plant-derived triterpenoids attract attention due to their structural diversity and broad biological activity. Roburic acid (RA), a tetracyclic triterpenoid, has previously been shown to exert antiproliferative effects in colorectal cancer (CRC) cells with limited cytotoxicity. In the present study, we investigated the cellular mechanisms underlying RA activity in CRC cells, focusing on cell cycle regulation, mitochondrial function, apoptosis, oxidative stress, and DNA integrity. RA treatment markedly suppressed CRC cell proliferation, resulting in G0/G1 cell cycle arrest and downregulation of key proliferation markers. Mitochondrial analysis revealed an early reduction in mitochondrial membrane potential (MMP) following RA exposure, indicating mitochondrial dysfunction. Importantly, these effects occurred in the absence of intracellular reactive oxygen species (ROS) generation and without induction of DNA strand breaks, demonstrating a non-pro-oxidant and non-genotoxic profile of RA. Apoptotic features were observed mainly at higher concentrations and after prolonged exposure and were strongly dependent on cell line and assay type. Overall, RA limits CRC cell growth predominantly through cytostatic mechanisms, including cell cycle arrest and mitochondrial modulation, while apoptosis is a secondary, context-dependent response. The lack of oxidative stress and genotoxicity distinguishes RA from many conventional cytotoxic agents and supports its further investigation as a non-genotoxic anticancer compound.

1. Introduction

Colorectal cancer (CRC) remains one of the most frequently diagnosed malignancies worldwide and represents a major global health burden. According to the most recent GLOBOCAN 2022 estimates, CRC accounts for approximately 1.9 million new cases and over 930,000 deaths annually, ranking third in global cancer incidence and second in cancer-related mortality [1]. Although incidence rates are stabilizing or declining in some high-income countries due to screening programs, CRC incidence continues to rise in many low- and middle-income regions. Notably, an alarming increase in early-onset CRC has been observed in individuals under 50 years of age, particularly in developed countries, suggesting shifts in environmental, lifestyle, and metabolic risk factors [2]. Despite advances in surgical techniques, chemotherapy, targeted therapies, and immunotherapy, treatment efficacy remains limited by the frequent development of drug resistance and the occurrence of severe adverse effects, often compromising patient outcomes and quality of life. In this context, increasing attention has been directed toward therapeutic approaches that suppress tumor progression while minimizing cytotoxicity. In particular, cytostatic agents that inhibit cancer cell proliferation without inducing extensive cell death are gaining interest, as they may reduce systemic toxicity and limit selective pressure favoring resistant tumor clones [3]. Natural compounds have emerged as a promising source of such agents due to their remarkable chemical diversity, multitargeted mechanisms of action, and generally favorable safety profiles. Consequently, they have attracted growing interest as potential anticancer therapeutics capable of modulating multiple pathways involved in cancer progression and therapy resistance.
Roburic acid (RA) is a tetracyclic triterpenoid isolated from the roots of Gentiana species and from oak galls (Figure 1). It has been reported to exhibit strong anti-inflammatory, anti-bacterial, anti-viral, and anti-atherogenic activities [3,4,5,6]. These effects are thought to stem from mechanisms that include the inhibition of cyclooxygenase enzymes and direct binding to tumor necrosis factor-alpha (TNF-α), thereby modulating pathways such as nuclear factor kappa B (NF-κB), which is associated with inflammatory responses, regulation of apoptotic proteins, and oncogenic transformation [4,7,8]. In our previous study, we demonstrated that RA significantly reduces metabolic activity, inhibits colony formation, and suppresses DNA synthesis across multiple cancer cell lines, with CRC models (DLD-1, HCT-116, and HT-29) showing relatively greater sensitivity [9]. These findings support an antiproliferative (cytostatic), rather than cytotoxic mode of action of this natural compound. However, the cellular and molecular mechanisms underlying this growth-inhibitory effect have not yet been fully elucidated.
Cytostatic responses in cancer cells may arise from diverse mechanisms, including alterations in cell cycle progression, activation of checkpoint pathways, mitochondrial dysfunction, or DNA damage signaling, even in the absence of pronounced cytotoxic effects. Therefore, a comprehensive analysis integrating functional, morphological, and molecular approaches is required to clarify the basis of RA-mediated growth inhibition. In the present study, we aimed to further elucidate the mechanisms of action of RA in CRC cells by evaluating its effects on cell proliferation, cell cycle regulation, DNA integrity, oxidative stress levels, mitochondrial function, and apoptotic responses. To this end, we employed a combination of fluorescence-based assays, cytometric analyses, and gene expression profiling to characterize RA activity across CRC cell lines representing distinct mutational backgrounds and chemoresistance profiles. Interestingly, our findings differ from some previously reported observations [7,10], potentially reflecting differences in experimental models, treatment conditions, or cellular contexts.
Based on these considerations, we hypothesized that RA exerts its antiproliferative effects in CRC cells through modulation of cell cycle progression and regulation of key molecular components involved in cell cycle control. To test this hypothesis, we performed a comprehensive mechanistic analysis evaluating RA-induced changes in cell cycle distribution, proliferation markers, mitochondrial function, oxidative stress levels, DNA integrity, and apoptotic responses in CRC cell lines representing distinct molecular backgrounds. Elucidating the mechanisms underlying RA-mediated growth inhibition may provide important insights into its potential as a novel anticancer candidate and contribute to a broader understanding of regulatory mechanisms controlling tumor cell proliferation in CRC.

2. Results

2.1. Ki-67 Immunofluorescence Assay

Building on the antiproliferative effects of RA previously reported by our group [9], the impact of RA on the expression of the proliferation marker Ki-67 was evaluated in CRC cell lines. Following 48 h of exposure to RA at 0.5 × IC50 and IC50 concentrations, Ki-67 immunofluorescence was significantly altered in all analyzed models (Figure 2). In DLD-1, treatment with RA at 0.5 × IC50 resulted in a modest but statistically significant increase in Ki-67 fluorescence compared with untreated controls (p < 0.0001), whereas exposure at IC50 led to a pronounced reduction in the Ki-67 signal (p < 0.0001). In HCT-116 and HT-29 cells, RA treatment resulted in a significant reduction in Ki-67 fluorescence at both 0.5 × IC50 and IC50 concentrations, indicating a robust antiproliferative response across CRC models. Quantitative analysis confirmed a statistically significant reduction in Ki-67 signal in RA-treated cells (p < 0.05), demonstrating a decreased proportion of actively cycling cells. These findings corroborate earlier observations obtained using bromodeoxyuridine incorporation assays and support the notion that RA limits CRC cell growth predominantly by inhibiting proliferative activity [7].

2.2. Cell Cycle Analysis

Cell cycle profiling revealed that RA treatment induced marked alterations in cell cycle distribution in all examined CRC cell lines. After 24 h of exposure, an increased proportion of cells in the G0/G1 phase was observed in DLD-1, HCT-116, and HT-29 cells compared with untreated controls (Figure 3A). This increase reached statistical significance at the IC50 concentration in DLD-1 (p = 0.0029) and HCT-116 cells (p = 0.0305), whereas in HT-29 cells, a significant accumulation in the G0/G1 phase was already detected at 0.5 × IC50 (p = 0.0002) and remained significant at IC50 (p < 0.0001). These shifts were accompanied by a reduction in the S-phase population, which was statistically significant in selected conditions (DLD-1: p = 0.0155; HCT-116: p = 0.0315; HT-29: p < 0.0001 for both concentrations), indicating impaired cell cycle progression. Following prolonged exposure to RA (48 h), alterations in cell cycle distribution became more pronounced (Figure 3B). All three cell lines exhibited a significant increase in the G0/G1 fraction (DLD-1: p < 0.0001 for both concentrations; HCT-116: p = 0.001 for 0.5 × IC50 and p = 0.05 for IC50; HT-29: p < 0.0001 for IC50), particularly at the IC50 concentration, along with a concomitant decline in the S-phase population (HCT-116: p = 0.0002; HT29: p < 0.0001). In addition, a sub-G1 fraction became detectable after 48 h of treatment, most notably in HT-29 cells at the higher RA concentration (p = 0.0015), suggesting the emergence of DNA fragmentation in a subset of cells. Overall, these findings indicate that RA disrupts cell cycle progression in CRC cells, with a predominant accumulation in the G0/G1 phase. The magnitude and statistical significance of these effects varied between cell lines and treatment conditions, consistent with a primarily cytostatic response that becomes more pronounced with prolonged exposure.

2.3. Annexin V-FITC Binding Assay

Given the appearance of a sub-G1 population following prolonged RA exposure, apoptotic responses were further evaluated using annexin V conjugated with fluorescein isothiocyanate (FITC) binding. After 24 h of treatment, a statistically significant increase in the proportion of annexin V–positive cells were detected only at the highest RA concentration (2 × IC50) in DLD-1 (32.13 ± 5.10%; p < 0.0001) and HCT-116 cells (28.07 ± 3.80%; p < 0.0001), whereas no significant changes were observed in HT-29 cells at this time point (Figure 4A). Prolonged exposure to RA markedly enhanced apoptotic responses (Figure 4B). After 72 h of treatment at 2 × IC50, annexin V–positive cells accounted for more than half of the cell population in all examined CRC cell lines (57.9 ± 8.0% in DLD-1, 51.6 ± 1.42% in HCT-116, and 51.8 ± 4.07% in HT-29; p < 0.05). In addition, treatment with RA at IC50 resulted in statistically significant increases in apoptotic cell fractions in HCT-116 (31.1 ± 3.15%; p < 0.0001) and HT-29 (20.73 ± 0.68%; p = 0.0019). Importantly, the fraction of necrotic cells remained low and did not reach statistical significance (p < 0.05) under any of the tested conditions (Supplementary Figures S1 and S2), indicating that RA-induced cell death was predominantly apoptotic. Together, these results indicate that apoptosis is not an immediate response to RA exposure but rather emerges following prolonged treatment and/or higher exposure conditions, consistent with a predominantly cytostatic effect at earlier time points.

2.4. Dual Acridine Orange/Ethidium Bromide Fluorescent Staining

Apoptotic morphology was evaluated in CRC cell lines using dual acridine orange/ethidium bromide (AO/EB) staining. After 24 h of RA exposure, a statistically significant increase (p < 0.05) in the proportion of apoptotic cells was observed only in DLD-1 cells treated with the highest RA concentration (2 × IC50), reaching 16.75 ± 3.0% (p = 0.001) compared with the untreated control (3.5 ± 2.65%) (Figure 5A). No significant changes were observed in HCT-116 or HT-29 cells at this time point. Prolonged exposure to RA (72 h) resulted in a more pronounced apoptotic response (Figure 5B). A statistically significant increase in apoptotic cells was observed in HCT-116 and HT-29 cell lines treated with RA at 2 × IC50 (14.75 ± 2.5%; p = 0.0387; and 22.75 ± 5.19%; p = 0.0003, respectively) compared with the untreated control (5.75 ± 4.57% and 8.13 ± 3.4%, respectively). In addition, HT-29 cells also exhibited a significant increase in apoptotic morphology at the IC50 concentration (18.00 ± 1.41%; p = 0.002). Overall, AO/EB staining confirmed that morphologically evident apoptosis occurs primarily following prolonged exposure and at higher RA concentrations, consistent with a delayed and secondary apoptotic response.

2.5. Mitochondrial Membrane Potential Assay

Alterations in mitochondrial membrane potential (MMP; ΔΨm) were examined in CRC cell lines following RA exposure. After 24 h of treatment, RA induced a reduction in MitoTracker Red CMXRos fluorescence intensity in DLD-1, HCT-116, and HT-29 cells, indicating mitochondrial membrane depolarization (Table 1, Figure 6). This effect was observed across most tested conditions and cell lines, except for HCT-116 cells treated with 0.5 × IC50 RA, where the decrease in ΔΨm did not reach statistical significance (p = 0.0501), although a clear downward trend was evident. The magnitude of ΔΨm reduction varied among the cell lines and concentrations. The most pronounced decrease in mitochondrial membrane potential was observed in HT-29 cells treated with RA at 2 × IC50, reaching 75.09 ± 2.67% (p < 0.0001) of the control value. Overall, the observed changes indicate that RA affects mitochondrial function within 24 h of exposure.

2.6. Determination of Intracellular Reactive Oxygen Species Level

Intracellular reactive oxygen species (ROS) levels were evaluated in CRC cell lines following RA exposure. Treatment with RA did not result in statistically significant changes (p > 0.05) in ROS production in DLD-1, HCT-116, or HT-29 cells at any of the tested concentrations compared with untreated controls (Figure 7). This lack of ROS induction was consistently observed throughout the monitoring period. In contrast, exposure to hydrogen peroxide, used as a positive control, led to a marked increase in fluorescence signal, confirming the sensitivity and responsiveness of the assay. Overall, these results indicate that RA does not promote intracellular ROS accumulation under the experimental conditions employed.

2.7. Genotoxicity—Alkaline Comet Assay

The genotoxic potential of RA was examined using the alkaline comet assay in CRC cell lines. After 24 h of exposure, RA did not induce a significant increase in DNA damage in DLD-1, HCT-116, or HT-29 cells at any of the tested concentrations compared with untreated controls (Figure 8). At the IC50 concentration, median DNA content in comet tails reached 3.82% in DLD-1 cells, 2.86% in HCT-116 cells, and 2.68% in HT-29 cells, values comparable to those observed in control samples. No statistically significant differences were detected between RA-treated cells and the negative control across all concentrations tested, indicating the absence of detectable DNA strand breaks under the experimental conditions employed. Representative comet images illustrating the lack of RA-induced DNA damage are shown in Figure 9, whereas cells treated with bleomycin, used as a positive control, exhibited pronounced DNA fragmentation. Importantly, no genotoxic effects of RA were detected in non-cancerous human cells (CCD-841 CoN and WI-38), further supporting the compound’s non-genotoxic profile (Supplementary Figure S3). Overall, these results demonstrate that RA does not exhibit genotoxic activity in CRC cells after 24 h of exposure.

2.8. Gene Expression Analysis by Quantitative Real-Time PCR

The expression of genes associated with cell proliferation and cell cycle regulation was analyzed by quantitative real-time PCR (RT-qPCR) following RA treatment at the IC50 concentration. Gene expression was normalized to the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and evaluated after 24 and 72 h of exposure. Across all examined CRC cell lines, RA treatment resulted in a statistically significant (p < 0.05) decrease in the proliferation marker Kiel 67 (MKI67) expression at both time points, consistent with reduced proliferative activity (Figure 10). A similar pattern was observed for proliferating cell nuclear antigen (PCNA), with significant reductions in transcript levels in DLD-1 and HCT-116 cells after 24 and 72 h of treatment. In HT-29 cells, a significant decrease in PCNA expression was observed only after prolonged exposure (72 h). Analysis of cyclin-dependent kinase inhibitor 1A (CDKN1A) expression revealed a time- and cell line–dependent response. A significant increase in CDKN1A transcript levels was detected in DLD-1 and HT-29 cells after 72 h of RA treatment, whereas in HCT-116 cells, elevated expression was observed at both 24 and 72 h. Expression of cellular tumor antigen p53 (TP53) exhibited divergent patterns among the analyzed cell lines. In DLD-1 cells, RA treatment led to a significant increase in TP53 expression at both time points analyzed. In contrast, a significant decrease in TP53 transcript levels was observed in HCT-116 cells after 24 h of exposure, while in HT-29 cells, a reduction in TP53 expression was detected only after 72 h. Collectively, these gene expression changes are consistent with the observed inhibition of proliferation and alterations in cell cycle progression induced by RA.

3. Discussion

In our previous study, RA was shown to exert pronounced cytostatic and antiproliferative effects in cancer and normal cells, with selective activity toward CRC models [9]. While these findings established the growth-inhibitory potential of RA, the cellular mechanisms underlying this effect remained unresolved. The present study addressed this gap by providing a comprehensive mechanistic evaluation of RA activity in CRC cells. Importantly, our data demonstrated that RA-mediated growth inhibition occurs independently of genotoxic stress, oxidative damage, and extensive cytotoxicity at concentrations corresponding to IC50 values. Instead, RA induced coordinated alterations in cell cycle progression and mitochondrial function, with apoptotic features emerging only under conditions of prolonged exposure or elevated concentrations. These findings support the concept that RA limits CRC cell growth predominantly through cytostatic mechanisms, acting as a regulator of proliferation rather than a conventional cytotoxic or cell-killing agent.
The marked inhibition of DNA synthesis and clonogenic potential observed in our previous study prompted a more detailed investigation into the mechanisms underlying RA-mediated growth inhibition, with a particular focus on cell cycle regulation. Since both bromodeoxyuridine incorporation and colony formation assays reflect the ability of cells to proliferate and sustain long-term growth actively, these findings suggest that RA interferes with fundamental proliferative processes. Flow cytometric evaluation revealed that RA treatment led to a clear accumulation of CRC cells in the G0/G1 phase, accompanied by a reduction in the S-phase population and, in some conditions, a decrease in the G2/M fraction. This pattern indicated that RA primarily disrupts early cell cycle checkpoints, enforcing a G0/G1 arrest that restricts the G1/S transition and limits entry into DNA synthesis. In selected cell lines and following prolonged exposure, the appearance of a sub-G1 population was also observed, which may reflect early apoptotic features emerging as a secondary and time-dependent consequence of sustained cell cycle perturbation. A similar concentration- and time-dependent shift from cytostatic to cytotoxic effects has been reported for several bioactive natural compounds [3,11,12,13,14]. Consistent with our findings, Xu et al. demonstrated that RA induces a pronounced G0/G1 accumulation in CRC cells (HCT-116 and HCT-115) after 24 h of exposure, accompanied by a reduction in the S-phase population [7]. However, based primarily on metabolic viability assays, they interpreted RA as predominantly cytotoxic and reported lower IC50 values. Differences in potency estimates across studies may arise from methodological and experimental factors, including the use of metabolic viability assays that are sensitive to cytostatic growth inhibition, as well as differences in exposure conditions, such as serum deprivation, which can increase cellular susceptibility to stress and cell death [15,16].
Given the observed alterations in cell cycle distribution, we next focused on the analysis of established molecular markers for cellular proliferation [17,18]. MKI67 and PCNA are widely used indicators of proliferative status, as they reflect the proportion of actively cycling cells and the cellular capacity for DNA replication, respectively [19,20]. Ki-67 is expressed exclusively during the active phases of the cell cycle (G1, S, G2, and M) and is absent in quiescent (G0) cells, making it a sensitive marker of proliferative activity rather than cell viability [20,21,22,23]. Consistent with the observed G0/G1 arrest, RA treatment resulted in a significant reduction in MKI67 transcript levels across all analyzed CRC cell lines, as determined by RT-qPCR. This transcriptional downregulation was further corroborated at the protein level by immunofluorescence staining using anti-Ki-67 antibodies, which revealed a marked decrease in Ki-67 expression. Notably, in DLD-1 cells exposed to a sub-inhibitory concentration of RA (0.5 × IC50), a modest but statistically significant increase in Ki-67 signal was observed. This apparent biphasic response is unlikely to reflect a genuine pro-proliferative effect of RA, but rather suggests adaptive or selective processes occurring under partial growth-inhibitory pressure. At low concentrations, antiproliferative agents may fail to suppress all cycling cells uniformly, thereby favoring the transient expansion or enrichment of more proliferative or resistant subpopulations [24,25]. Importantly, at the IC50 concentration, RA induced a pronounced and statistically significant reduction in Ki-67 expression in DLD-1 cells, consistent with effective suppression of proliferative activity. In contrast, HCT-116 and HT-29 cells exhibited a significant decrease in Ki-67 expression already at 0.5 × IC50, indicating a more uniform antiproliferative response in these models. Taken together, these changes provide molecular evidence for a shift in RA-treated cells toward a non-proliferative, quiescent-like state. While MKI67 reflects the fraction of actively cycling cells, PCNA represents a complementary marker associated with DNA replication competence. At the protein level, PCNA functions as a central coordinator of DNA replication and S-phase progression, acting as a sliding clamp for DNA polymerases and participating in DNA repair processes [19,20,23,26]. The observed reduction in PCNA expression, particularly in DLD-1 and HCT-116 cells and following prolonged exposure in HT-29 cells, is consistent with impaired DNA replication competence and further complements the changes observed in proliferation markers. Given PCNA’s established role in coordinating DNA synthesis and repair, its modulation under RA treatment is compatible with an adaptive cytostatic response, in which replication-associated processes are progressively restrained rather than abruptly disabled.
To further explore the molecular basis of RA-induced G0/G1 arrest, we analyzed transcripts involved in checkpoint regulation. A consistent feature across the models was the induction of CDKN1A (P21), particularly after prolonged exposure. P21 is a canonical cyclin-dependent kinase inhibitor that restrains G1/S progression by inhibiting cyclin-dependent kinase 2 (CDK2)–cyclin E/A complexes and thereby limits commitment to DNA synthesis, functioning as a key effector of cytostatic checkpoint enforcement [27,28,29,30]. Importantly, although P21 is classically regulated by P53, its expression can also be induced through TP53-independent pathways, enabling effective checkpoint control even in tumor cells harboring TP53 mutations [29,30]. This regulatory flexibility is particularly relevant in the context of the heterogeneous TP53 status of the analyzed CRC cell lines and provides a mechanistic framework for interpreting the divergent TP53 transcriptional responses observed in this study. CDKN1A upregulation occurred rapidly and was sustained in HCT-116 cells, which retain wild-type TP53, whereas in DLD-1 and HT-29 cells, both characterized by mutant TP53 backgrounds, CDKN1A induction became apparent predominantly after prolonged exposure [31,32,33]. The lack of uniform correlation between TP53 transcript and CDKN1A induction likely reflects the predominantly post-translational regulation of P53 and cell-line specific feedback mechanisms [34,35]. Functionally, P21-mediated checkpoint activation links these transcriptional changes to the observed antiproliferative phenotype. By enforcing G0/G1 arrest, P21 reduces the proportion of actively cycling cells, as reflected by reduced MKI67 expression and decreased Ki-67 protein levels [36]. In parallel, P21 can directly interact with PCNA and restrain DNA replication processes, thereby contributing to the reduced replication competence observed in RA-treated cells. Modulation of PCNA expression, particularly in DLD-1 and HCT-116 cells and following prolonged exposure in HT-29 cells, is consistent with progressive limitation of DNA replication capacity characteristic of an adaptive cytostatic state. In this context, the previously reported ability of RA to suppress NF-κB signaling may be mechanistically relevant [9]. RA has been shown to inhibit TNF-induced phosphorylation of IκB kinase (IKK) α/β, inhibitor of κB α (IκBα), and the P65 (RelA) subunit of NF-κB, prevent IκBα degradation, and block nuclear translocation of P65, leading to reduced NF-κB–dependent gene expression [7,9]. Given the well-established antagonistic crosstalk between NF-κB and the P53/P21 pathways, suppression of NF-κB activity by RA could alleviate constraints on pro-survival and pro-proliferative signaling, thereby facilitating P21-driven G0/G1 arrest independently of TP53 transcriptional dynamics [37,38,39]. Together with our transcriptional and functional data, these observations indicate that RA interferes with cell cycle progression at multiple regulatory levels, reinforcing its role as a modulator of proliferative capacity.
Importantly, the magnitude and nature of these molecular effects appeared to be strongly dependent on RA concentration and exposure time. At concentrations corresponding to IC50 values, RA predominantly induced transcriptional modulation of proliferation-associated genes, including downregulation of MKI67 and PCNA and upregulation of CDKN1A, consistent with checkpoint activation and G0/G1 arrest. These alterations were detectable prior to the emergence of substantial apoptotic fractions, indicating that RA initially exerts a growth-regulatory effect rather than immediate cytotoxicity. In contrast, at higher concentrations (2 × IC50) and following prolonged exposure, apoptotic features became more pronounced, suggesting that increased RA burden may exceed the adaptive capacity of CRC cells and shift the cellular response toward activation of cell death pathways. Thus, the balance between cytostatic and cytotoxic outcomes appears to be determined by RA concentration and treatment duration [40,41].
Mitochondrial function is tightly linked to cellular energy homeostasis, proliferation, and survival, and its modulation may underlie both cytostatic and cytotoxic responses [42,43]. In the present study, RA induced a significant reduction in MMP, as assessed using the mitochondria-selective fluorescent probe MitoTracker Red CMXRos, within 24 h of exposure across CRC cell lines, indicating an early disturbance of mitochondrial function. Importantly, these depolarizations occurred under conditions in which extensive apoptotic or necrotic cell death was not predominant, suggesting that mitochondrial perturbation represents an early, mechanistically distinct event rather than a direct trigger of acute cell death. A decrease in ΔΨm does not inevitably signify activation of the intrinsic apoptotic pathway but may instead reflect impaired mitochondrial bioenergetics [44]. Because progression through the cell cycle and DNA synthesis are highly energy-dependent processes, even moderate disruption of mitochondrial polarization may restrict ATP availability and thereby limit the proliferative capacity of cancer cells [45,46,47]. In this context, the observed mitochondrial depolarization is fully consistent with the G0/G1 arrest, reduced S-phase entry, and downregulation of proliferation markers induced by RA, supporting a model in which mitochondrial dysfunction contributes to a predominantly cytostatic phenotype. Notably, RA-induced changes in ΔΨm were not accompanied by increased intracellular ROS generation. Given that mitochondrial dysfunction is frequently associated with oxidative stress and ROS-driven cytotoxicity, the absence of ROS accumulation indicates that RA-mediated mitochondrial alterations occur independently of overt oxidative damage. Only at higher concentrations and prolonged exposure did mitochondrial perturbation coincide with increased apoptotic fractions, suggesting that sustained mitochondrial stress may eventually exceed the adaptive capacity of CRC cells and trigger activation of cell death pathways.
Consistent with the absence of ROS generation, RA did not induce DNA strand breaks, as assessed by the alkaline comet assay, further supporting the lack of genotoxic stress. Importantly, to our knowledge, this is the first study (as of 27 January 2026) to systematically evaluate the genotoxic potential of RA in both CRC cells and non-cancerous human cell models (CCD-841 CoN and WI-38). Under the applied experimental conditions, RA did not increase DNA damage levels relative to untreated controls, while the assay’s responsiveness was confirmed using bleomycin as a positive control. The combined absence of ROS induction and DNA damage clearly distinguishes RA from many conventional anticancer agents, whose mitochondrial effects are tightly coupled to oxidative injury and genome destabilization [48,49,50]. In contrast, RA appears to modulate mitochondrial function, limiting proliferative capacity without eliciting acute genomic damage. Such a mechanistic profile distinguishes RA from classical chemotherapeutic agents that suppress tumor growth primarily through DNA damage or oxidative stress. This non-genotoxic mode of action is particularly relevant in the context of long-term or combination-based therapeutic strategies, as it implies a reduced risk of therapy-induced mutagenesis.
The emergence of a sub-G1 population in cell cycle analysis, together with the early loss of MMP, prompted further investigation into RA’s pro-apoptotic activity in CRC cells. These observations suggested that, under certain conditions, sustained cell cycle and mitochondrial perturbations might progress toward activation of cell death pathways. Assessment of apoptosis by annexin V-FITC flow cytometry revealed an increase in phosphatidylserine-positive cells, predominantly at the highest tested concentration (2 × IC50) and following prolonged exposure. In contrast, at concentrations corresponding to IC50 values, annexin V positivity was limited or absent at earlier time points. Further evaluation using dual AO/EB staining, which identifies apoptotic cells based on morphological criteria associated with later stages of cell death, revealed a markedly lower fraction of apoptotic cells compared with annexin V/FITC analysis [51]. This discrepancy reflects the different biological processes captured by these methods. Annexin V/FITC binding detects early externalization of phosphatidylserine, which may represent an initial and potentially reversible stage of apoptosis, whereas AO/EB staining identifies cells that have progressed to advanced, morphologically defined apoptotic stages [52,53]. Consequently, annexin V positivity does not necessarily indicate irreversible commitment to apoptotic execution [54,55,56]. Notably, even under prolonged exposure conditions, the necrotic fraction remained low and non-dominant across all tested cell lines, indicating that RA does not induce nonspecific membrane damage or acute cytotoxicity. Instead, these data suggest that RA primarily imposes sustained cytostatic stress, with apoptotic features emerging only when mitochondrial and cell-cycle perturbations exceed the adaptive capacity of cancer cells. In contrast to our findings, Xu et al. reported pronounced apoptosis at lower RA concentrations [7]. However, their experiments were conducted under serum-deprived conditions, which may lower the apoptotic threshold. Differences in baseline stress levels and biological characteristics of the analyzed cell lines further complicate direct comparison. These discrepancies likely reflect context-dependent variability in RA responsiveness rather than fundamental mechanistic divergence.
Taken together, the present study extends our previous observations by providing an integrated mechanistic framework linking mitochondrial perturbation, transcriptional modulation, and growth inhibition in RA-treated CRC cells. At concentrations corresponding to IC50 values, RA induced early mitochondrial depolarization accompanied by coordinated downregulation of proliferation markers and induction of CDKN1A, resulting in G0/G1 arrest and a sustained cytostatic phenotype. Importantly, these effects occurred in the absence of detectable ROS accumulation or DNA damage, indicating that RA does not act as a classical oxidative or genotoxic cytotoxic agent. A clear concentration-dependent relationship emerged: moderate RA exposure predominantly enforced checkpoint-mediated growth restraint, whereas higher concentrations and prolonged exposure were required for substantial apoptotic engagement. Thus, mitochondrial perturbation and transcriptional reprogramming appear mechanistically linked to the observed antiproliferative effect, while apoptotic cell death represents a secondary consequence of sustained proliferative and metabolic stress rather than the primary driver of growth inhibition. This distinction is particularly relevant when interpreting viability-based readouts and underscores the importance of integrating complementary functional and molecular assays when characterizing natural compounds. Collectively, these findings position RA as a non-genotoxic, non-pro-oxidative modulator of cancer cell proliferation that restricts tumor growth through regulatory interference with bioenergetic and cell cycle networks rather than through acute destructive cytotoxicity. Future studies should further delineate the upstream signaling pathways connecting mitochondrial dysfunction to checkpoint activation and explore how this growth-restrictive profile may be leveraged in rational combination-based therapeutic strategies.

4. Materials and Methods

4.1. Chemicals

Cell culture reagents: Fetal bovine serum (FBS), phosphate-buffered saline (PBS), MEM, and RPMI-1640 culture medium were purchased from Biowest (CytoGen, Zgierz, Poland). Stabilized penicillin-streptomycin solution and trypsin-ethylenediamine tetraacetic acid (EDTA) were obtained from Merck/Sigma Aldrich Chemical Co. (Burlington, MA, USA). Black and clear-bottom 96-well plates, as well as 12-well and 6-well plates, were supplied by Thermo Fisher Scientific (Waltham, MA, USA). Comet assay reagents: normal melting point agarose, low-melting point agarose, bleomycin sulfate, sodium hydroxide solution, EDTA, disodium salt dihydrate, tris(hydroxymethyl)aminomethane (TRIS), Triton X-100, sodium chloride solution, and 4,6-diamidino-2-phenylindole (DAPI) were purchased from Merck/Sigma Aldrich Chemical Co. (Burlington, MA, USA). Apoptosis, mitochondrial function, and oxidative stress assays: MitoTracker CMX Ros was obtained from Invitrogen (Waltham, MA, USA). The Annexin V/FITC Apoptosis Detection Kit I was supplied by B.D. Biosciences (Franklin Lakes, NJ, USA). AO/BE, as well as the ROS probe H2DCFDA, were purchased from Merck/Sigma Aldrich Chemical Co. (Burlington, MA, USA). RNA isolation and gene expression analysis: Total RNA was isolated using Fenozol reagent (A&A Biotechnology, Gdańsk, Poland). Reverse transcription was performed using the TranScriba noGenome Kit (A&A Biotechnology, Gdańsk, Poland). Quantitative PCR was carried out using qPCR-HS SYBR Mix (A&A Biotechnology, Gdańsk, Poland).

4.2. Cell Culture

Human CRC cell lines DLD-1 (adenocarcinoma; ATCC® CCL-221TM), HCT-116 (adenocarcinoma; ATCC® CCL-247TM), and HT-29 (carcinoma; ATCC® HTB-38TM), as well as normal cell lines WI-38 (human lung fibroblasts, ATCC® CCL-75TM) and CCD-841 CoN (human colon epithelial, ATCC® CRL-1790TM), were obtained from American Type Culture Collection (ATCC, Rockville, MD, USA). CRC cell lines were cultured in RPMI-1640 medium, whereas non-cancerous WI-38 and CCD-841 CoN cells were maintained in MEM medium, both supplemented with 10% (v/v) fetal bovine serum and 1% (v/v) penicillin–streptomycin solution. Cultures were maintained at 37 °C in a humidified atmosphere containing 5% CO2. The culture medium was replaced every 24–48 h, and cells were subcultured using 0.25% trypsin-EDTA upon reaching approximately 80–90% confluence. Mycoplasma contamination was routinely monitored monthly using the MycoBlueTM Mycoplasma Detector kit (Vazyme Biotech, Nanjing, China).

4.3. Cell Treatment

RA concentrations applied in the present study were selected based on previously published cytotoxicity and antiproliferative data obtained using MTT, crystal violet, and neutral red assays. Consensus IC50 values derived from these assays were used to establish the working concentrations for subsequent experiments. Depending on the experimental design, cells were treated with RA at 0.5 × IC50, IC50, or 2 × IC50. The exact RA concentrations used in individual assays are summarized in Table 2.

4.4. Ki-67 Immunofluorescence Assay

Ki-67 immunofluorescence staining was performed using standard procedures for the detection of nuclear proliferation markers, with modifications [54]. Cell proliferation was assessed by immunofluorescence staining of the nuclear proliferation marker Ki-67. DLD-1, HCT-116, and HT-29 cells were seeded onto glass coverslips placed in 12-well plates at a density of 3 × 104 cells/well and allowed to adhere for 24 h. Cells were subsequently treated with RA at concentrations corresponding to 0.5 × IC50 and IC50 for 48 h. Following treatment, cells were washed with PBS and fixed in 70% ethanol at −20 °C. After permeabilization with PBS containing 0.5% Triton X-100, DNA was denatured using 4 N HCl and neutralized with 0.1 M borax. Samples were then incubated for 1 h with a rabbit anti-Ki-67 primary antibody (Thermo Fisher, MA5-14520; 1:250 dilution in PBS supplemented with 1% bovine serum albumin and 0.5% Tween-20), followed by a 1 h incubation with an Alexa Fluor 488-conjugated goat anti-rabbit IgG secondary antibody (Thermo Fisher, A-11008; 1:500 dilution). Cell nuclei were counterstained with DAPI (1 µg/mL), and coverslips were mounted using Fluoromount-G. Fluorescence images were acquired using an Olympus BX60 F5 fluorescence microscope equipped with 20× and 40× objectives. For each sample, images of at least 200 cells were captured using CellSens imaging software v1.7 (Olympus Corporation, Tokyo, Japan). Quantitative analysis of Ki-67 fluorescence intensity was performed using ImageJ version 1.54f (ImageJ software, National Institutes of Health, Bethesda, MD, USA). Nuclear regions of interest were defined, and fluorescence intensity was quantified as corrected total cell fluorescence (CTCF), calculated as integrated density minus background fluorescence. The data are presented as median values and expressed relative to untreated controls.

4.5. Cell Cycle Analysis

Cell cycle distribution was analyzed by flow cytometry following propidium iodide (PI) staining. DLD-1, HCT-116, and HT-29 cells were seeded in 6-well plates at a density of 0.5 × 106 cells/mL and allowed to attach for 24 h. The assay was performed according to the previously described protocols [55,56]. Cells were subsequently treated with RA at concentrations corresponding to 0.5 × IC50 and IC50 for 24 or 48 h. After treatment, cells were harvested and centrifuged at 1400 rpm for 10 min at 4 °C. Cell pellets were washed with PBS and fixed dropwise in ice-cold 70% ethanol with continuous vortexing. Prior to flow cytometry analysis, cells were washed with PBS and incubated for 30 min at 37 °C in the dark in a staining solution containing 5 μg/mL PI and 10 U/mL RNase A in PBS. DNA content was measured using an LSR II flow cytometer (Becton Dickinson, San Jose, CA, USA) equipped with a 488 nm excitation laser and a 530 nm emission filter. A minimum of 10,000 events were collected per sample. The data were analyzed using FlowJo version 10.8 (FlowJo software, Becton, Dickinson, and Company, Ashland, OR, USA), and cell populations were quantified in the G0/G1, S, G2/M, and sub-G1 phases.

4.6. Annexin V-FITC Binding Assay

Apoptotic cell death was evaluated in DLD-1, HCT-116, and HT-29 cells using the FITC Annexin V Apoptosis Detection Kit I, based on the detection of phosphatidylserine externalization on the outer leaflet of the plasma membrane [52,53]. The combined use of annexin V–FITC and PI enabled discrimination between viable, early apoptotic, late apoptotic, and necrotic cells. The assay was performed according to the previously described protocols [55,56]. Cells were cultured in 6-well plates at densities of 7 × 105 cells/mL or 3.5 × 105 cells/mL and allowed to attach for 24 h. Subsequently, cells were treated with RA at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50 for 24 or 72 h. After treatment, cells were harvested, washed with PBS, and centrifuged at 1400 rpm for 10 min at 4 °C. Cell pellets were resuspended in Annexin V binding buffer and stained with Annexin V-FITC and PI for 15 min in the dark at room temperature. Samples were analyzed using an LSR II flow cytometer with fluorescence detection at 530 nm. A minimum of 10,000 events were acquired per sample. Data were analyzed using FlowJo version 10.8.

4.7. Dual Acridine Orange/Ethidium Bromide Fluorescent Staining

Dual staining with AO and EB, combined with morphological assessment, was used to distinguish viable, necrotic, and apoptotic cells [57,58]. AO penetrates intact cell membranes and intercalates into nucleic acids, emitting green fluorescence in viable cells, whereas EB is membrane-impermeable and stains only cells with compromised membrane integrity, emitting red fluorescence. The assay was performed according to the previously described protocols [56,59]. DLD-1, HCT-116, and HT-29 cells were seeded in 12-well plates at densities of 1.5 × 105 cells/mL (for 24 h treatment) or 7.5 × 104 cells/mL (for 72 h treatment) and allowed to attach for 24 h. Cells were then treated with RA at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50 for 24 and 72 h. After treatment, cells were harvested, centrifuged at 1400 rpm for 10 min at 4 °C, and resuspended in 25 μL of PBS. AO/EB staining solution (1 μL; 100 μg/mL of each dye) was added to the cell suspension, and 10 μL of the mixture was placed on glass microscope slides. Samples were examined immediately under a fluorescence microscope (Olympus BX60 F5, Olympus Optical Co., Ltd., Nagano, Japan). At least 200 cells per condition were scored and classified as viable, necrotic, or apoptotic based on differential dye uptake and characteristic morphological features.

4.8. Mitochondrial Membrane Potential Assay

MMP was assessed using the fluorescent probe MitoTracker Red CMXRos, which selectively accumulates in active mitochondria. The assay was performed according to the previously described protocols [56,59]. DLD-1, HCT-116, and HT-29 cells were seeded in black 96-well plates at a density of 1.2 × 104 cells/mL and allowed to attach for 24 h. Cells were then treated with RA at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50 for 24 h. Following treatment, cultures were washed with PBS and incubated with MitoTracker Red CMXRos (0.1 µM) for 40 min in the dark. After staining, fluorescence intensity was measured using a SpectraMax® i3x Multi-Mode Detection Platform (Molecular Devices, San Jose, CA, USA) at excitation and emission wavelengths of 579 and 599 nm, respectively. Results were expressed as mean fluorescence intensity compared with untreated control cells (normalized to 100%).

4.9. Determination of Intracellular Reactive Oxygen Species Level

Intracellular ROS levels were determined using the fluorescent probe H2DCFDA according to the protocol described by Ruiz-Leal and George [60]. DLD-1, HCT-116, and HT-29 cells were seeded in black 96-well plates at a density of 1.2 × 104 cells/mL and allowed to attach for 24 h. Cells were then loaded with H2DCFDA (20 µM) for 20 min in the dark, washed with PBS, and treated with RA at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50. H2O2 (50 µM) was used as a positive control. Fluorescence intensity was recorded at excitation and emission wavelengths of 485 and 535 nm, respectively, using a SpectraMax® i3x Multi-Mode Detection Platform over 2 h at 15 min intervals.

4.10. Alkaline Comet Assay

The alkaline comet assay was performed to evaluate DNA strand breaks in CRC cell lines according to the protocol described by Kciuk et al. [61]. DLD-1, HCT-116, and HT-29 cells, as well as non-cancerous WI-38 and CCD-841 CoN cells, were seeded in 12-well plates at a density of 1.2 × 105 cells/mL and allowed to attach for 24 h. Cells were then treated with RA at concentrations corresponding to 0.5 × IC50 and IC50 for an additional 24 h. The 20 µM bleomycin was chosen as a positive control. Following treatment, cells were processed for the comet assay as previously described [61]. Briefly, cells were embedded in agarose on microscope slides, lysed, and subjected to electrophoresis under alkaline conditions. After electrophoresis, DNA was stained with DAPI (1 μg/mL) and visualized using a fluorescence microscope (Olympus) equipped with CellSens software at an excitation wavelength of 360 nm. For each experimental condition, at least 200 cells were analyzed. DNA damage was quantified as the mean percentage of DNA content in comet tails with the interquartile range, minimum, and maximum.

4.11. Gene Expression

4.11.1. RNA Isolation and cDNA Synthesis

DLD-1, HCT-116, and HT-29 cells were seeded in 6-well plates at densities of 5 × 105 cells/well (24 h incubation) or 2 × 105 cells/well (72 h incubation) and treated with RA at the IC50 concentrations or left untreated as controls. After 24 or 72 h, total RNA was isolated using the phenol–chloroform extraction method with Fenozol Plus reagent (cat. no. 203-100P; A&A Biotechnology, Gdańsk, Poland) according to the manufacturer’s instructions. RNA concentration and purity were determined spectrophotometrically using a BioDrop µLite + system (Biochrom Ltd., Cambridge, UK), based on absorbance ratios at 260/280 nm and 260/230 nm. Complementary DNA was synthesized from 250 ng of total RNA using the TranScriba noGenome Kit (cat. no. 4000NG-100; A&A Biotechnology) according to the manufacturer’s protocol.

4.11.2. Quantitative Real-Time PCR (RT-qPCR)

Quantitative real-time PCR was performed with SYBR Green chemistry with qPCR-HS Mix SYBR (cat. no. 2008HS-100; A&A Biotechnology) on a real-time PCR thermocycler (RotorGene Q, QIAGEN, Hilden, Germany). Primer sequences used for RT-qPCR analysis are provided in Supplementary Table S1. Thermal cycling conditions were as follows: initial denaturation at 95 °C for 3 min, followed by 40 cycles of denaturation at 95 °C for 15 s, annealing at 54–56 °C (primer-dependent) for 30 s, and extension at 72 °C for 30 s.
Relative gene expression was calculated using the ΔCt method. Threshold cycle (Ct) values were obtained using Rotor-Gene Q Series software v2.3.5. For each sample, Ct values of target genes were normalized to the reference gene GAPDH according to the following formula:
ΔCt = Cttarget gene − Ctref gene.
Due to the non-normal distribution of RT-qPCR data, statistical analyses were performed on ΔCt values. For graphical presentation, gene expression data are shown as −ΔCt values to facilitate visualization of relative changes in transcript levels. All reactions were performed in duplicate, and mean Ct values were used for further analyses.

4.12. Statistical Analysis

Statistical analyses were performed using GraphPad Prism version 8.0 (GraphPad Software, San Diego, CA, USA) and Statistica version 13.3 (TIBCO Software Inc., Palo Alto, CA, USA). Data normality was assessed using the Shapiro–Wilk test. Data obtained from annexin V binding assay, MMP measurements, intracellular ROS determination, AO/EB double staining assays, and cell cycle analysis are presented as mean ± SD. Results from Ki-67 immunofluorescence analysis, the alkaline comet assay, and RT-qPCR are presented as medians with interquartile ranges because of non-normal distributions. For comparisons involving more than two groups, one-way analysis of variance (ANOVA), followed by Dunnett’s or Tukey’s post hoc tests, was applied for normally distributed data. Cell cycle distribution and intracellular ROS measurements were additionally analyzed using two-way ANOVA, followed by Dunnett’s or Tukey’s multiple comparisons tests as appropriate. For non-normally distributed data, the Kruskal–Wallis test followed by Dunn’s post hoc test was used. Statistical significance was defined as p < 0.05. For the alkaline comet assay, at least 200 cells were analyzed per experimental condition. The comet assay, MMP assay, annexin V binding assay, and intracellular ROS determination were performed in triplicate. The AO/EB double staining assay was conducted in two independent experiments. RT-qPCR reactions were performed in duplicate for each sample.

5. Conclusions

RA does not exhibit genotoxic activity in CRC cells, indicating that its antiproliferative effects are not mediated by DNA damage. RA induces an early reduction in mitochondrial membrane potential without concomitant ROS generation or oxidative stress, distinguishing its mode of action from that of many conventional cytotoxic anticancer agents. Under conditions corresponding to IC50 values, RA primarily exerts cytostatic effects, manifested by cell cycle arrest, suppression of proliferation markers, and limited induction of cell death. Apoptotic features become apparent mainly at the highest concentrations (2 × IC50) or following prolonged exposure and are highly dependent on cell line–specific and experimental conditions, suggesting that apoptosis represents a secondary consequence of sustained cellular stress rather than a primary mechanism of action. Collectively, these findings indicate that RA limits CRC cell growth predominantly through cytostatic mechanisms of the cell cycle and mitochondrial function, rather than through direct cytotoxicity, oxidative stress, or genotoxic injury. This non-genotoxic, non-oxidant profile highlights RA as a biologically active compound with a distinct mechanism of action and potential relevance for long-term or combination-based anticancer strategies. In particular, its growth-modulatory rather than DNA-damaging activity suggests possible utility as an adjuvant agent aimed at restraining tumor proliferation while minimizing therapy-associated mutagenic risk. Future investigations should therefore include in vivo validation, pharmacokinetic assessment, and evaluation of RA in rational combination regimens with standard chemotherapeutics or targeted agents. Furthermore, additional studies are warranted to elucidate the precise signaling pathways involved, including those related to cell cycle control, mitochondrial regulation, and adaptive stress responses such as autophagy or senescence.

Supplementary Materials

The supporting information can be downloaded at https://www.mdpi.com/article/10.3390/ijms27052478/s1.

Author Contributions

Conceptualization, A.G., M.K., R.G., S.W. and R.K.; collected literature, A.G.; wrote the manuscript, A.G.; supervised project and proofread the paper, A.G., M.K., R.G., S.W. and R.K. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available in the main text of this article/Supplementary Materials of this article, or on request from the corresponding author.

Acknowledgments

The authors acknowledge the support of the Laboratory of Microscopic Imaging and Specialized Biological Techniques at the University of Lodz, which provided access to specialized microscopy and biological research equipment.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

AO/EBAcridine orange/ethidium bromide
CAIICarbonic anhydrase II
CDKN1ACyclin-dependent kinase inhibitor 1A
CES1Carboxyl esterase 1
CRCColorectal cancer
CtThreshold cycle
CTCFCorrected total cell fluorescence
DAPI4,6-diamidino-2-pheny-lindole
EDTAEthylenediamine tetraacetic acid
EGFREpidermal growth factor receptor
FBSFetal bovine serum
FITCFluorescein isothiocyanate
GAPDHGlyceraldehyde-3-phosphate dehydrogenase
H2DCFDA2′,7′-dichlorodihydrofluoresceindiacetate
H2O2Hydrogen peroxide
IKKIκB kinase
IκBαInhibitor of κB α
MKI67Marker of proliferation Kiel 67
MMPMitochondrial membrane potential
NF-κBNuclear factor kappa B
PBSPhosphate-buffered saline
PCNAProliferating cell nuclear antigen
PIPropidium iodide
PLA2G2APhospholipase A2
RARoburic acid
ROSReactive oxygen species
RT-qPCRQuantitative real-time PCR
TNF-αTumor necrosis factor-alpha
TP53Tumor antigen p53
TRISTris(hydroxymethyl)aminomethane

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Figure 1. Chemical structure of roburic acid (RA). Image adapted from PubChem (CID: 5317800).
Figure 1. Chemical structure of roburic acid (RA). Image adapted from PubChem (CID: 5317800).
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Figure 2. Effect of roburic acid (RA) on Ki-67 expression in colorectal cancer (CRC) cell lines. Ki-67 immunofluorescence staining in DLD-1, HCT-116, and HT-29 following 48 h of exposure to RA at concentrations corresponding to 0.5 × IC50 and IC50. Quantitative data are presented as medians with interquartile ranges, minimums, and maximums. Statistical analysis was performed using the Kruskal–Wallis test, followed by Dunn’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
Figure 2. Effect of roburic acid (RA) on Ki-67 expression in colorectal cancer (CRC) cell lines. Ki-67 immunofluorescence staining in DLD-1, HCT-116, and HT-29 following 48 h of exposure to RA at concentrations corresponding to 0.5 × IC50 and IC50. Quantitative data are presented as medians with interquartile ranges, minimums, and maximums. Statistical analysis was performed using the Kruskal–Wallis test, followed by Dunn’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
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Figure 3. Cell cycle distribution of DLD-1, HCT-116, and HT-29 colorectal cancer (CRC) cells following (A) 24 h and (B) 48 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50 and IC50. The data are presented as the mean percentages of cells [%] ± SD in G0/G1, S, and G2 phases. Statistical analysis was performed using a two-way ANOVA, followed by Dunnett’s multiple comparisons test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
Figure 3. Cell cycle distribution of DLD-1, HCT-116, and HT-29 colorectal cancer (CRC) cells following (A) 24 h and (B) 48 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50 and IC50. The data are presented as the mean percentages of cells [%] ± SD in G0/G1, S, and G2 phases. Statistical analysis was performed using a two-way ANOVA, followed by Dunnett’s multiple comparisons test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
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Figure 4. Apoptotic cell fraction in DLD-1, HCT-116, and HT-29 colorectal cancer (CRC) cell lines following (A) 24 h and (B) 72 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50. Apoptosis was assessed by annexin V-FITC binding. The data are presented as the mean percentages [%] ± SD of annexin V-positive cells. Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05), N > 1 × 104.
Figure 4. Apoptotic cell fraction in DLD-1, HCT-116, and HT-29 colorectal cancer (CRC) cell lines following (A) 24 h and (B) 72 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50. Apoptosis was assessed by annexin V-FITC binding. The data are presented as the mean percentages [%] ± SD of annexin V-positive cells. Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05), N > 1 × 104.
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Figure 5. Percentage of apoptotic cells in colorectal cancer (CRC) cell lines DLD-1, HCT-116, and HT-29 following (A) 24 h and (B) 72 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50, assessed by dual acridine orange/ethidium bromide (AO/EB) staining. The data are shown as mean percentages [%] ± SD of AO/EB-identified apoptotic cells. Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05), N > 1 × 104.
Figure 5. Percentage of apoptotic cells in colorectal cancer (CRC) cell lines DLD-1, HCT-116, and HT-29 following (A) 24 h and (B) 72 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50, assessed by dual acridine orange/ethidium bromide (AO/EB) staining. The data are shown as mean percentages [%] ± SD of AO/EB-identified apoptotic cells. Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05), N > 1 × 104.
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Figure 6. Changes in mitochondrial membrane potential (MMP; ∆Ψm) in DLD-1, HCT-116, and HT-29 cells following 24 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50, assessed using MitoTracker Red CMX Ros. The data are presented as mean fluorescence intensity ± SD relative to untreated control (normalized to 100%). Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
Figure 6. Changes in mitochondrial membrane potential (MMP; ∆Ψm) in DLD-1, HCT-116, and HT-29 cells following 24 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50, assessed using MitoTracker Red CMX Ros. The data are presented as mean fluorescence intensity ± SD relative to untreated control (normalized to 100%). Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
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Figure 7. Intracellular reactive oxygen species (ROS) levels in colorectal cancer (CRC) cell lines (A) DLD-1, (B) HCT 116, and (C) HT-29 following exposure to RA at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50, assessed using the 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA) probe. The data are presented as means relative fluorescence intensity ± SD over 2 h. Hydrogen peroxide (H2O2) was used as a positive control. Statistical analysis was performed using two-way ANOVA followed by Tukey’s multiple comparisons test (p < 0.05).
Figure 7. Intracellular reactive oxygen species (ROS) levels in colorectal cancer (CRC) cell lines (A) DLD-1, (B) HCT 116, and (C) HT-29 following exposure to RA at concentrations corresponding to 0.5 × IC50, IC50, and 2 × IC50, assessed using the 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA) probe. The data are presented as means relative fluorescence intensity ± SD over 2 h. Hydrogen peroxide (H2O2) was used as a positive control. Statistical analysis was performed using two-way ANOVA followed by Tukey’s multiple comparisons test (p < 0.05).
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Figure 8. DNA damage in DLD-1, HCT-116, and HT-29 colorectal cancer (CRC) cells was assessed by the alkaline comet assay after 24 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50 and IC50. The data are presented as mean percentage of DNA in comet tails with interquartile range, minimum, and maximum. Statistical analysis was performed using the Kruskal–Wallis test, followed by Dunn’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05), N > 200.
Figure 8. DNA damage in DLD-1, HCT-116, and HT-29 colorectal cancer (CRC) cells was assessed by the alkaline comet assay after 24 h of exposure to roburic acid (RA) at concentrations corresponding to 0.5 × IC50 and IC50. The data are presented as mean percentage of DNA in comet tails with interquartile range, minimum, and maximum. Statistical analysis was performed using the Kruskal–Wallis test, followed by Dunn’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05), N > 200.
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Figure 9. Representative images from the alkaline comet assay. DLD-1: (A) control, (B) RA IC50; HCT-116: (C) control, (D) RA IC50; HT-29: (E) control, (F) RA IC50; and (G) bleomycin (20 μM)-treated cells used as a positive control. The images are shown at identical magnification (40×).
Figure 9. Representative images from the alkaline comet assay. DLD-1: (A) control, (B) RA IC50; HCT-116: (C) control, (D) RA IC50; HT-29: (E) control, (F) RA IC50; and (G) bleomycin (20 μM)-treated cells used as a positive control. The images are shown at identical magnification (40×).
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Figure 10. Transcript levels of MKI67, PCNA, CDKN1A, and TP53 in colorectal cancer (CRC) cell lines DLD-1, HCT-116, and HT-29 following treatment with roburic acid (RA) at the concentrations corresponding to IC50 for 24 and 72 h. Gene expression was analyzed by quantitative real-time PCR (RT-qPCR) and normalized to the housekeeping gene GAPDH. The data are presented as −ΔCt values (medians with interquartile ranges). Statistical analysis was performed using the Kruskal–Wallis test, followed by Dunn’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
Figure 10. Transcript levels of MKI67, PCNA, CDKN1A, and TP53 in colorectal cancer (CRC) cell lines DLD-1, HCT-116, and HT-29 following treatment with roburic acid (RA) at the concentrations corresponding to IC50 for 24 and 72 h. Gene expression was analyzed by quantitative real-time PCR (RT-qPCR) and normalized to the housekeeping gene GAPDH. The data are presented as −ΔCt values (medians with interquartile ranges). Statistical analysis was performed using the Kruskal–Wallis test, followed by Dunn’s post hoc test; an asterisk indicates significant differences relative to untreated controls (*) (p < 0.05).
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Table 1. Changes in the mitochondrial membrane potential (MMP; ∆Ψm) in DLD-1, HCT-116, and HT-29 cells following 24 h of exposure to roburic acid (RA). Results are expressed as mean relative fluorescence ± SD compared to untreated controls (normalized to 100%). Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test (p < 0.05; significant values indicated in bold).
Table 1. Changes in the mitochondrial membrane potential (MMP; ∆Ψm) in DLD-1, HCT-116, and HT-29 cells following 24 h of exposure to roburic acid (RA). Results are expressed as mean relative fluorescence ± SD compared to untreated controls (normalized to 100%). Statistical analysis was performed using one-way ANOVA, followed by Dunnett’s post hoc test (p < 0.05; significant values indicated in bold).
Cell LineResults of 24 h Incubation of Cancer Cells with RA
[%] Fluorescence Intensity Compared to Negative Control ± SD *
0.5 × IC50IC502 × IC50
DLD-184.67 ± 6.00
(p = 0.005)		86.00 ± 6.33
(p = 0.005)		82.00 ± 6.33
(p = 0.0012)
HCT-11691.61 ± 6.67
(p = 0.0501)		91.23 ± 2.33
(p = 0.0404)		84.87 ± 3.33
(p = 0.0013)
HT-2988.46 ± 2.00
(p = 0.0022)		81.90 ± 4.67
(p < 0.0001)		75.09 ± 2.67
(p < 0.0001)
(*) The asterisk indicates significant differences relative to untreated controls (p < 0.05).
Table 2. Roburic acid (RA) concentrations were applied in the individual experimental assays.
Table 2. Roburic acid (RA) concentrations were applied in the individual experimental assays.
TimeConcentrationCell Line
DLD-1HCT-116HT-29CCD-841 CoNWI-38
24 h0.5 × IC508.1210.1711.217.6317.61
IC5016.2320.3322.3935.2535.21
2 × IC5032.4740.6644.7870.570.42
72 h0.5 × IC504.616.035.38.757.66
IC509.2112.0610.617.515.32
2 × IC5018.4224.1221.1935.030.64
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Gielecińska, A.; Kciuk, M.; Gruszka, R.; Wawrocki, S.; Kontek, R. Roburic Acid as a Therapeutic Candidate: Antiproliferative Activity and Secondary Cell Death Response in Colorectal Cancer Cells. Int. J. Mol. Sci. 2026, 27, 2478. https://doi.org/10.3390/ijms27052478

AMA Style

Gielecińska A, Kciuk M, Gruszka R, Wawrocki S, Kontek R. Roburic Acid as a Therapeutic Candidate: Antiproliferative Activity and Secondary Cell Death Response in Colorectal Cancer Cells. International Journal of Molecular Sciences. 2026; 27(5):2478. https://doi.org/10.3390/ijms27052478

Chicago/Turabian Style

Gielecińska, Adrianna, Mateusz Kciuk, Renata Gruszka, Sebastian Wawrocki, and Renata Kontek. 2026. "Roburic Acid as a Therapeutic Candidate: Antiproliferative Activity and Secondary Cell Death Response in Colorectal Cancer Cells" International Journal of Molecular Sciences 27, no. 5: 2478. https://doi.org/10.3390/ijms27052478

APA Style

Gielecińska, A., Kciuk, M., Gruszka, R., Wawrocki, S., & Kontek, R. (2026). Roburic Acid as a Therapeutic Candidate: Antiproliferative Activity and Secondary Cell Death Response in Colorectal Cancer Cells. International Journal of Molecular Sciences, 27(5), 2478. https://doi.org/10.3390/ijms27052478

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