Pyrimidine-based compounds as promising anticancer agents targeting tumor cell senescence

Abstract

The pyrimidine ring is an important structural motif present in numerous bioactive compounds, including those with antibacterial and antitumor activities. In this study, two novel 4,6-disubstituted-2-(4-morpholinyl) pyrimidines (P12 and P14) were evaluated in breast cancer and melanoma cells with different genetic backgrounds. Both compounds demonstrated antitumor activity by reducing cell proliferation in 2D and 3D models. Mechanistic studies showed that these derivatives induce cell-cycle delay, altering the transcription of key cell-cycle mediators and leading to a senescent-like phenotype accompanied by changes in the pattern of the senescence-associated secretory phenotype (SASP). In addition, both compounds appeared to induce cell death, potentially in distinct tumor cell subpopulations. Collectively, these findings highlight the potential of pyrimidine-based derivatives as lead compounds for the development of pro-senescent anticancer agents, with utility in cancers where senescence plays a critical role, such as BRAF-mutant melanoma and the ER+/HER2− breast cancer subtype investigated in this study.

Graphical abstract

Two novel 4,6-disubstituted-2-(4-morpholinyl) pyrimidines (P12, P14) reduced proliferation in breast cancer and melanoma cells. They induced cell-cycle delay, a senescent-like phenotype with SASP changes and cell death, highlighting pyrimidine derivatives as promising pro-senescent anticancer agents.

Introduction

Cancer represents a global health burden, with estimated 9.7 million deaths in 2022.¹ Recently, breast cancer has been reported as the most frequently diagnosed cancer worldwide. Of these cases, more than 70% are hormone-receptor-positive and HER2-negative.² In contrast, skin melanoma is among the most aggressive cancer types due to its high metastatic potential and associated mortality. Additionally, 40%–80% of melanoma patients carry a driver mutation in BRAF.^1,3 Over recent decades, a pivotal shift has occurred in the clinical management of cancer, moving from broad-spectrum cytotoxic agents to targeted therapies aimed at specific tumor-driving pathways.⁴ One of the most significant advances has been the development of small-molecule inhibitors that selectively interfere with oncogenic signaling pathways. For example, approved agents such as the BRAF/MEK inhibitor vemurafenib for BRAF-mutant melanoma and CDK4/6 inhibitors for ER+/HER2− breast cancer exemplify how targeted strategies can dramatically improve clinical outcomes.^5,6,7,8 Although targeted therapies have brought important advances in tumor treatment, cancer therapy still faces major challenges, including drug resistance and the need for combination approaches.^9,10,11 This underscores the importance of developing novel agents with distinct mechanisms of action. In this context, heterocyclic compounds, particularly those based on the pyrimidine ring, have shown considerable promise in cancer research.¹²

Pyrimidine is a privileged scaffold found in different biologically active molecules with antibacterial, antiviral, and antitumor properties.^13,14,15 It forms the core of several clinically used drugs, such as the statin rosuvastatin and the PI3K inhibitor NVP-BKM120 (buparlisib), which highlights its chemical versatility and pharmacological relevance.^16,17 On the other hand, therapeutic targeting of senescence has gained considerable importance in oncology in recent years. However, senescent tumor cells can act as a double-edged sword: while initially halting proliferation and tumor progression, their persistent presence and the secretion of a complex cocktail of inflammatory factors, collectively known as the senescence-associated secretory phenotype (SASP), may contribute to therapy resistance and relapse.¹⁸ Importantly, the composition of the SASP varies substantially depending on both the cellular context and the senescence trigger, resulting in distinct profiles that can either enhance immune system activation and promote the clearance of senescent cells or conversely drive immunosuppression, inflammation, and tumor progression.^19,20,21 Given the beneficial effects of senescence as an antitumor mechanism, considerable efforts have been directed toward the development of drugs that exploit this process.²² This has led to the emergence of senolytic agents, which selectively eliminate senescent cells, and senomorphic agents, which modulate the SASP to attenuate its deleterious effects while preserving the tumor-suppressive functions of senescence.^23,24,25,26

Several targeted therapies have been shown to exert their anticancer effects, at least in part, through the induction of senescence. Notably, BRAF/MEK inhibitors in BRAF-mutant melanoma and CDK4/6 inhibitors in ER+/HER2− breast cancer can trigger a therapy-induced senescent response, contributing to durable tumor control in some patients.^27,28 It is also important to note that many conventional chemotherapeutics, as well as radiotherapy, can induce senescence and are still widely used in cancer treatment.²⁹ At present, some of the most successful pro-senescence drugs are CDK4/6 inhibitors, which have demonstrated clinical benefit in ER+/HER2− breast cancer and show promise in several other cancer types.^30,31,32 Nevertheless, these targeted approaches still face important limitations, including intrinsic and acquired resistance. Combining senolytics with targeted therapies has shown promising results. For example, the senolytic agent navitoclax, a BCL-2 family inhibitor, has demonstrated efficacy in preclinical cancer models by selectively clearing senescent cells and reducing tumor regrowth.^33,34 However, toxicity remains a significant concern and has limited its clinical translation.^23,24

In this study, two novel 4,6-disubstituted-2-(4-morpholinyl) pyrimidines (P12 and P14)³⁵ were identified as potential candidates with pro-senescence and antitumor activity. Structurally, P12 is a non-symmetrical derivative containing a 1-naphthyl group at C-4 and a trifluoromethylphenyl group at C-6, while P14 is a symmetrical pyrimidine with a 2-benzothiophene unit at C-4 and C-6. We introduce these pyrimidine-based compounds and show that, by affecting cell-cycle progression, they enhance cellular senescence, alter the pattern of the SASP, and induce cell death in two distinct tumor types. Our results demonstrate that 4,6-disubstituted-2-(4-morpholinyl) pyrimidines represent a valuable platform for the development of agents capable of arresting tumor growth and preventing tumor progression through the induction of cellular senescence and cell death.

Results

Pyrimidine-based compounds reduce the proliferation capacity of different tumor types

In line with our ongoing interest in identifying antitumoral candidates, this study focuses on evaluating symmetrical and non-symmetrical pyrimidine derivatives, specifically P12 and P14, which were previously synthesized via palladium-catalyzed cross-coupling with indium organometallics.³⁵ Additionally, we consider that pyrimidines bearing a 4-morpholinyl moiety at the C-2 position have been reported to exhibit antitumor activity.^16,17 Our study focused on targeting luminal breast cancer cell line MCF7 (ER+/HER2−) and the BRAFV600E-mutant melanoma line A375. Dose-response experiments were conducted using MCF7 and A375 cell lines to assess the activity of the pyrimidine-based compounds in cancer cells. Different concentrations of P12 and P14 were applied to confluent cell monolayers (2D) for 7 days, with drug and medium renewed every 48 h, and the effects on cell proliferation were evaluated using a colony formation assay (Figure 1A). As shown, P12 and P14 significantly reduced the colony-forming capacity of both breast and melanoma cells in a dose-dependent manner (Figures 1B and 1C). Since P14 showed a potent anti-proliferative effect even at low concentrations (5 μM) in both cell lines, we performed IC₅₀ calculations. In MCF7 cells, the IC₅₀ values were 30.22 μM (P12) and 2.018 μM (P14), in melanoma cells these were 2.915 μM (P12) and 1.887 μM (P14) (Figure 1D). Taking these data and the colony formation assay into account, the reduction in proliferation observed with P14 at high concentrations (20–50 μM) may reflect a toxic effect. To assess toxicity in non-tumoral cells, we treated the chondrocyte cell line TC28a2 with different concentrations of P12 and P14 for 7 days (Figure 1E). The results indicated that P12 did not a produce significant reduction in the number of adherent viable cells from 5 to 35 μM in non-tumoral TC28a2 cells, whereas high concentrations of P14 (up to 20 μM) potentially induced cytotoxicity (Figure 1E).

Figure 1.

Pyrimidine-based compounds reduce the proliferation capacity of cancer cells

(A) Diagram showing experiment setup for the treatment of cancer cells with P12 and P14 for 7 days. (B) Colony formation capacity of breast cancer cells (MCF7) under the treatment of different concentration of P12 (0, 5, 20, 35, 50 μM) and P14 (0, 5, 20, 35, 50 μM) for 7 days. Representative images are shown. Quantifications are shown below. (C) Representative images of a dose-response experiment using colony formation of melanoma cells (A375) after 7 days treatment with P12 (0, 5, 20, 35, 50 μM) and P14 (0, 5, 20, 35, 50 μM). Quantifications are shown below. (D) IC₅₀ calculations for P12 and P14 in MCF7 (right) and A375 (left). The table below shows the IC₅₀ of each compound. (E) Crystal violet staining to determine the effect of P12 and P14 in a non-tumoral cell line (TC28a2). The cells were treated for 7 days using the following concentrations—0, 5, 20, 35, 50 μM—for both compounds. Drugs were refreshed every 48 h. (F) Spheroids of MCF7 and A375 cells treated with 5 μM of P12 and P14 to demonstrate cell proliferation arrest in a 3D culture. Spheroids were cultured for 12 days. The graph represents the mean of six spheroids/day. Data represent the mean ± SEM of three independent experiments. Two-tailed Student’s t test and ANOVA were used to calculate the significance represented as follows: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001.

Subsequently, we aimed to confirm the results in a more physiologically relevant setting by using 3D spheroid cultures. Both pyrimidines, P12 and P14, were tested at 5 μM in this 3D tumor model (Figure 1F). Spheroid-like structures of breast and melanoma cells were treated with P12 and P14 for 12 days, with drug and medium renewed every 48 h. The spheroids were monitored every 2 days, and their area was measured. In ER+/HER2− breast cancer cells (MCF7), P12 and P14 significantly reduced spheroid area compared with the untreated control, but only after 8 and 12 days of treatment. This suggests a role in cell-cycle delay or cellular senescence rather than cell death, as cells remained viable for at least 4 days in culture in the presence of both pyrimidines (Figure 1F, left). In contrast, BRAF-mutated melanoma cells (A375) grown in 3D structures showed different response. P12 had a more pronounced effect on melanoma spheroids, significantly reducing their size at a concentration as low as 5 μM and after only 4 days in culture (Figure 1F, right). P14 also showed an inhibitory effect, although the reduction in spheroid size was less marked than with P12, but still statistically significant (Figure 1F). Taken together, these results show that both pyrimidine-based compounds exert anti-proliferative effects in at least two distinct molecular contexts in cancer cells. However, the underlying biological effects and mechanisms may differ between models.

Induction of G1/S cell-cycle arrest by pyrimidine-based compounds

Based on the abovementioned results, we next investigated the potential effects of P12 on the cell cycle and its possible mechanism of action. To this end, cancer cells were treated with P12 and P14 for 7 days, followed by analysis of cell-cycle distribution by flow cytometry (Figure 2A). As shown in Figure 2A, treatment with P12 and P14 suggested a trend toward cell-cycle alterations, as evidenced by a reduction in the proportion of cells progressing to the G2 phase in both cell lines (MCF7: 15.65% with P12 and 13.10% with P14; A375: 1.94% with P12 and 0.52% with P14) compared with the untreated controls (MCF7: 26.25%; A375: 5.44%). The effects of the pyrimidine compounds differed between tumor cell types, probably due to their distinct genetic backgrounds. A375 cells showed a higher proportion of cells in G0/G1 and a lower proportion in G2/M compared with untreated MCF7 cells. After treatment with both pyrimidines, MCF7 cells exhibited significantly higher proportions of cells in G2 phase compared with A375 cells, indicating that a larger fraction of MCF7 cells continued to progress through the cell cycle and prepare for mitosis. In contrast, the markedly lower G2 percentages observed in A375 cells suggest either a more rapid transition through this phase, cell-cycle arrest at an earlier stage, or reduced proliferative activity. Quantitative PCR (qPCR) was performed to evaluate the expression levels of key cell-cycle-related genes (Figure 2B). The results indicated that P12 and P14 exert their antiproliferative effects through modulation of cell-cycle regulators. In melanoma cells, treatment with P12 and P14 produced a stronger antiproliferative effect, as evidenced by significant downregulation of key regulators of the G1/S transition and S-phase entry, including CCND1, CDK4, CDK6, and CCNE1. In contrast, breast cancer cells exhibited a weaker response, with these genes showing only moderate changes in expression compared with treated melanoma cells, indicating a less pronounced inhibitory effect. Overall, these findings highlight a cell-type-specific sensitivity to treatments. In melanoma cells, particularly with P12, there was a significant reduction in the expression of cell-cycle-related genes involved in the G1/S transition, S-phase entry, and G2 phase. In contrast, in MCF7 breast cancer cells, treatment with P12 and P14 produced a different biological effect, with a significant upregulation of genes associated with G1/S transition and S-phase entry (Figure 2B).

Figure 2.

Treatment with pyrimidine-based compounds induces phenotypic hallmarks of senescence in cancer cells

(A) Cell cycle was performed after treatment with 5 μM of P12 and P14. Cells were cultured until they reached approximately 80% confluency. At that point, 10 μM of 5-ethynyl-2′-deoxyuridine (EdU) was added to the medium to label cells actively synthesizing DNA during the S phase. Following a 2-h incubation at 37°C to enable EdU incorporation, cells were collected and fixed. This was done by suspending the cells in PBS, centrifuging at 500 × g for 5 min, and then resuspending the pellet in 500 μL of 70% ethanol for fixation. MCF7 UT (G0/G1: 11.65%, S: 61.1%, G2/M: 26.25%); MCF7 P12 (G0/G1: 20.35%, S: 63.8%, G2/M: 15.65%); MCF7 P14 (G0/G1: 15.9%, S: 68.15%, G2/M: 13.1%); A375 UT (G0/G1: 34.45%, S: 57.45%, G2/M: 5.44%); A375 P12 (G0/G1: 25.4%, S: 72.9%, G2/M: 1.94%); A375 P14 (G0/G1: 33.2%, S: 67.3%, G2/M: 0.52%). (B) Heatmap showing the mRNA levels of different cell-cycle regulators in MCF7 and A375 cells after treatment with 5 μM of P12 and P14 for 7 days. (C) qPCR analysis showing the expression levels of the senescent markers p21, p53, and p53-related genes IGFBP3 and GDF15 in MCF7 cells (left) and A375 (right) after treatment with 5 μM of P12 and P14 for 7 days. The mean of three independent experiments is shown. (D) Representative images of the SA-β-galactosidase activity in cancer cells after treatment with P12 and P14. Quantifications are shown in the right. (E) Heatmap showing the mRNA levels of SASP factors (IL-6, IL-8, MMP3, and MMP9) in MCF7 and A375 cells after treatment with 5 μM of P12 and P14 for 7 days. Data represent the mean ± SEM of three independent experiments. (F) Flow cytometry in MCF7 and A375 after treatment with 5 μM of P12 and P14 for 7 days. Cells were stained using a dual labeling approach with PI at 2 μg/mL and YO-PRO-1 at 150 nM. Cells negative for both PI and YO-PRO-1 were classified as viable, while apoptotic stages were determined according to the levels of YO-PRO-1 or PI incorporation. Viable cells were identified using forward scatter (FSC) and side scatter (SSC) parameters to exclude cellular debris. Additionally, single cells were separated from aggregates based on FSC-H and FSC-A gating (n = 3). (G) Graphical abstract illustrating the effects of pyrimidine compounds in the different cellular models. Data are presented as mean ± SEM. Two-tailed Student's t test and one-way ANOVA were used to calculate the significance represented as follows: ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.0001.

Treatment with pyrimidine-based compounds induces phenotypic hallmarks of senescence in cancer cells

As the effects of both compounds required more than 96 h (4 days) of treatment, we next investigated whether the pyrimidines P12 and P14 could induce a senescent-like phenotype, which is typically characterized by stable cell-cycle arrest.^36,37 qPCR analysis of genes implicated in cellular senescence showed a notable increase in the expression of senescence-associated factors, including p21-, p53-, and p53-responsive genes involved in growth arrest, senescence, or apoptosis (IGFBP3 and GDF15), in both ER+/HER2− and BRAF-mutant cells treated with P12 and P14. These results indicate that the pyrimidine compounds may induce a senescent-like phenotype (Figure 2C). However, no significant changes were observed in β-galactosidase activity, a well-established marker of senescence (Figure 2D). This discrepancy between gene expression and β-galactosidase activity suggests that P12 and P14 may induce partial senescence or trigger senescent-like changes that do not fully conform to classical senescence markers.

To further explore the potential senescence-like state induced by P12 and P14, we next examined their ability to promote the secretion of SASP factors. Cytokine profiling by qPCR revealed a significant upregulation of classical SASP components, including the pro-inflammatory cytokines interleukin (IL)-6 and IL-8, as well as metalloproteinases involved in tissue remodeling, following treatment with P12 and P14 in ER+/HER2− breast cancer cells (Figure 2E, upper panel). However, in BRAF-mutant melanoma cells, treatment with P12 and P14 for 7 days did not induce SASP factor expression; instead, a trend toward reduced synthesis was observed, suggesting an alteration in the SASP profile (Figure 2E, lower panel). These results are consistent with those obtained for cell-cycle regulators, where differences were observed depending on the tumor genetic background, although both models exhibited proliferation arrest. In melanoma cells, P12 and P14 likely suppress proliferation primarily through direct cell-cycle inhibition (Figures 2A and 2B). By contrast, in breast cancer cells, their effects may be partially mediated through additional regulatory pathways, with transcriptional changes influencing factors such as SASP components that can indirectly, or directly, affect cell-cycle entry (Figures 2A–2E). In both cancer models, however, P12 and P14 induced a significant upregulation of p21 and p53, central effectors of senescence (Figure 2C), although the mechanisms driving this induction may differ between tumor types.

Since we cannot exclude the activation of cell death during treatment, we next explored whether P12 and P14 could induce tumor cell death by apoptosis. Flow cytometry was used to assess early apoptotic, apoptotic, and necrotic events in treated cancer cells. The results showed a tendency to increase cell death and mainly in the proportion of cells undergoing early apoptosis following treatment with P12 and P14 in both melanoma and breast cancer cell lines (Figure 2F). These findings suggest that, in addition to their effects on cell-cycle entry and progression, P12 and P14 may also trigger an apoptotic response. Altogether, our data provide evidence that P12 and P14 exert distinct biological activities, inducing a senescence-like state while also potentially enhancing cell death through apoptosis. These dual effects highlight their potential as promising antitumor agents.

Discussion

The present study investigates the antitumor effects and mechanisms of action of 4,6-disubstituted-2-(4-morpholinyl) pyrimidines, specifically compounds P12 and P14, in ER+/HER2− breast cancer and BRAF-mutant melanoma cell lines. Our findings suggest that P12 and P14 exert their anticancer activity through distinct biological effects, including the alteration of cell-cycle progression, possibly via the induction of a senescence-like phenotype, as well as the promotion of apoptosis. Moreover, at the selected concentrations, both compounds displayed selective activity against cancer cells, with minimal effects on non-tumor cells.

It has been demonstrated that pyrimidine analogues, such as 5-fluorouracil, significantly reduce tumor cell growth by impairing DNA synthesis.^38,39 Similarly, other reports have shown that pyrimidine-based compounds targeting the thymidylate synthase enzyme inhibit the proliferation of breast and colon cancer cells.^40,41 Our findings on two previously untested pyrimidine-based compounds suggest that P12 and P14 act through different mechanism, potentially by directly affecting cell-cycle progression. Specifically, they appear to arrest proliferation at the G1/S transition in BRAF-mutant cells, while delaying cell-cycle entry in ER+/HER2− cells. Our study revealed that these compounds act through complex mechanisms that ultimately lead to anti-proliferative effects, but in a context-dependent manner. Their activity varies according to the type of cancer and the specific molecular characteristics of the tumor, as evidenced by differences in cell-cycle distribution and the induction of senescence following P12 and P14 treatment in both genetic contexts analyzed (BRAF-mutant melanoma and ER+/HER2− breast cancer). Although both compounds interfere with cell-cycle progression, they also appear to exert effects at the transcriptional level, as evidenced by their differential impact on the expression of cell-cycle-related genes and the synthesis of SASP factors. In BRAF-mutant cells, treatment with P12 and P14 led to a pronounced reduction in proliferation, accompanied by a marked downregulation of key regulators of the G1/S transition and S-phase progression, including CCND1, CDK4, CDK6, and CCNE1. Our results also indicate that the antiproliferative effects of P12 and P14 are more moderate in breast cancer cells, as ER+/HER2^- cells displayed a comparatively weaker response to treatment. The overall inhibitory effect was less pronounced, and in some cases, the expression of cell-cycle-related genes was even upregulated. Collectively, these data highlight a differential sensitivity to P12 and P14 that depends on cell type and molecular context, with BRAF-mutant cells being more responsive to their cell-cycle-inhibiting properties. However, in both cell types we detected a significant increase in the key senescence regulators p21 and p53. Together with the observed changes in the expression of SASP factors, these findings suggest that both compounds may induce a senescence-like phenotype, albeit with distinct characteristics depending on the cellular context.

Senescence has been recognized as a critical anti-cancer mechanism, and recent studies have demonstrated that the induction of senescence in cancer cells can prevent further tumor growth and trigger immune surveillance.^42,43 Specifically, SASP factors, including pro-inflammatory cytokines such as IL-6 and IL-8, can activate immune cells and enhance tumor recognition and clearance.^44,45 Our cytokine profiling data revealed increased expression of IL-6 and IL-8 in breast cancer cells treated with P12 and P14, supporting the notion that these compounds may induce a senescence-like phenotype capable of modulating antitumor activity and the tumor microenvironment in our breast cancer model. Interestingly, our results showed that although P12 and P14 promoted the expression of senescence-associated genes, no significant changes in β-galactosidase activity were detected. This discrepancy suggests that P12 and P14 may trigger an incomplete senescence response or promote the elimination of senescent cells, potentially through apoptosis. This possibility is particularly relevant given that several pyrimidine-based compounds, such as dasatinib, a tyrosine kinase inhibitor with a pyrimidine scaffold, have been shown to selectively eliminate senescent cells by triggering apoptosis.⁴⁶ Therefore, if P12 and P14 contain pyrimidine-like structures or modulate pathways associated with cell survival, it is plausible that their antiproliferative effects somehow might extend to senolytic activity as well. Similar findings have been reported where certain therapeutic compounds, including pyrimidine analogues, can induce senescence without fully activating all canonical senescence markers, such as β-galactosidase.^47,48 Furthermore, while SASP factor induction was observed in breast cancer cells, no significant increase—only slight changes—in these specific SASP factors was detected in melanoma cells. This discrepancy may reflect the tumor-specific nature of SASP factor secretion, as the molecular context of each tumor type can influence how cells respond to senescence induction. In fact, previous reports have shown that different cancer types can exhibit varying levels of SASP factor secretion depending on their genetic and epigenetic backgrounds, which may explain the limited effects on SASP observed in melanoma cells treated with P12 and P14.⁴⁹

As mentioned, flow cytometry analysis showed that P12 and P14 may also enhance cell death in both melanoma and breast cancer cells. Although the effect can be considered modest, the induction of cell death may represent an important mechanism. Based on our results, we interpreted that P12 has a stronger effect in the melanoma model in both 2D and 3D assays (Figures 1C–1F). The effects on cell-cycle arrest and senescence are also stronger in melanoma cells under P12 treatment (Figures 2A–2E). However, when we analyzed cell death, the results cannot be directly compared because MCF7 cells show higher basal apoptosis than A375 cells under untreated conditions, and MCF7 appears to be more prone to activate apoptosis than A375 under treatments. The detected effects on cell death could also be diluted by the high heterogeneity of cancer cell populations, where the compounds may exert different actions on distinct subpopulations. Based on our results, apoptosis appears to be a secondary but complementary route to the senescence-like effects induced by these pyrimidine-based compounds and in both tumor cell models.

Altogether, our data demonstrate that the pyrimidine-based compounds P12 and P14 exert strong antiproliferative effects in breast cancer and melanoma models by inducing cell-cycle arrest and senescence-like phenotypes, while also activating cell death by apoptosis. Importantly, the divergent molecular responses observed between tumor types emphasize the context-dependent nature of their activity and the need to evaluate their efficacy across different cancer models (Figure 2G). We propose P12 and P14 as promising antitumor candidates, but further studies, including in vivo validation, are required to determine their therapeutic potential and impact within the tumor microenvironment.

Materials and methods

Cell lines

Human cell lines ER+/HER2− MCF7 (breast cancer) and BRAF-mutant A375 (melanoma) and healthy TC28a2 (chondrocytes) were grown in Dulbecco’s modified Eagle’s medium (DMEM) (Lonza) with the addition of 10% fetal bovine serum (FBS) (Gibco, Thermo Fisher Scientific) and antibiotics (100 U/mL penicillin and 100 mg/mL streptomycin) (Gibco, Thermo Fisher Scientific) following standard procedures.^27,50,51 All cells were maintained at 37°C in a humidified incubator with 5% CO₂ (SANYO CO₂). The culture medium was refreshed every two days.

Pyrimidines treatment

Pyrimidine-12 (P12) and pyrimidine-14 (P14) were synthesized and provided by the research group Synthetic and Catalytic Methodologies at CICA (Universidade da Coruña).¹⁵ The molecules were resuspended and used from a stock solution in DMSO at 10 mM. Cells were treated with concentrations between 5 and 50 μM for 7 or 12 days.

Colony formation assay

Colony formation assays were conducted as previously described.⁵⁰ Briefly, 5,000 to 50,000 cells per well in 6- to 12-well plates were seeded and allowed to grow for 7 to 15 days. The medium or treatments were refreshed every 48 h. Cells were washed with warm saline solution and fixed with cold 4% paraformaldehyde (PFA) for 15 min at room temperature. After washing with PBS, the cells were stained with 0.05% crystal violet (Sigma-Aldrich) for 15 min at room temperature. The cells were then rinsed with distilled water and air-dried at room temperature. Colonies were quantified by dissolving the crystal violet in 30% acetic acid. From this solution, 100 μL were taken to measure absorbance at 570 nm using a NanoQuant microplate reader Infinite M200 (TECAN).

IC50 calculation

The 50% inhibitory concentrations (IC₅₀) were determined using cell cultures treated with various concentrations of the treatments. Cells were stained with crystal violet, and cell numbers were assessed by measuring absorbance at 570 nm with a NanoQuant microplate reader Infinite M200 (TECAN), according to established protocols by or lab.²⁷

Spheroid formation and analysis

Single-cell suspensions (200 μL) containing 1,500 cells per well were seeded into ultra-low attachment, round-bottom 96-well plates (Corning) using the appropriate culture medium for each cell type. The plates were centrifuged at 1,300 rpm for 5 min to promote cell aggregation and then placed under standard cell culture conditions. Cells were left undisturbed for at least 48 h to allow spheroid formation. Starting from day 2, spheroid development was regularly assessed by optical microscopy. Half of the culture medium was replaced every 2 days, and spheroid areas were quantified using ImageJ software.

Gene expression

Total RNA was isolated from cells using TRI Reagent RT (Probiotek) according to the manufacturer’s instructions, as previously described.²⁷ Cells were harvested and lysed in 1 mL of TRIzol reagent. Following this, 200 μL of chloroform was added, and the mixture was vortexed. Samples were incubated for 5 min at 4°C and then centrifuged at 14,000 rpm for 15 min at 4°C. The upper aqueous phase, which contains the RNA, was transferred to clean tubes containing 500 μL of isopropanol. The samples were vortexed, incubated for 30 min at −20°C, and centrifuged again at 14,000 rpm for 15 min at 4°C. The supernatant was discarded, and the RNA pellets were washed with 500 μL of 70% ethanol, then centrifuged for 5 min at 14,000 rpm at 4°C. After discarding the supernatant, the tubes were air-dried. RNA was resuspended in 20 μL of DNase/RNase/Protease-free water (Sigma-Aldrich). The RNA samples were quantified using a Nanodrop ND-1000 (Thermo Fisher Scientific). One microgram of each sample was used to synthesize complementary DNA (cDNA) with Superscript IV VILO Master Mix (Invitrogen, Thermo Fisher Scientific) following the manufacturer’s protocol. The cDNA samples were then resuspended in DNase/RNase/Protease-free water (Sigma-Aldrich). One microgram of cDNA was combined with 0.5 μL of primer mix, which included 10 μM of each primer (Table 1), 10 μL of Applied Biosystems PowerUP SYBR Green Master Mix (Applied Biosystems, Thermo Fisher Scientific), and completed with dH₂O to a total volume of 20 μL per well on a LightCycler 480 System (Roche). The program used consisted of an initial denaturing cycle of 10 min at 95°C, followed by 50 amplification cycles of 10 s at 95°C, 30 s at 60°C for annealing, and 12 s at 72°C for extension.²⁷

Table 1.

Sequences of primers used for gene expression analysis

Gene name	Forward sequence (5′-3′)	Reverse sequence (5′-3′)
CDKN1A (p21)	GACTCTCAGGGTCGAAAACG	GCCAGGGTATGTACATGAGGA
IL6	TGTAGCCGCCCCACACA	GGATGTACCGAATTTGTTTGTA
IL8	TTGGCAGCCTTCCTGATTTC	TCTTTAGCACTCCTTGGCAAAAC
MMP9	TTCATCTTCCAAGGCCAATC	GGGCTGAACCTGGTAGACAG
CDND1	TCCTCTCCAAAATGCCAGAG	AGCGTGTGAGGCGGTAGTAG
CDND2	CACTTGTGATGCCCTGACTG	AACTGGCATCCTCACAGGTC
CDK4	CAGATGGCACTTACACCCGT	CAGCCCAATCAGGTCAAAGA
CDK6	CGTGGTCAGGTTGTTTGATGT	CGCTGTGAAAGAAAGTCC
ACTB	GCCCTGAGGCACTCTTCCA	CGGATGTCCACGTCACACTTC
MMP3	TGCTTTGTCCTTTGATGCTG	AGCCTGTGCCTTCAAAAATG
CCNE2	TCTCCTGGCTAAATCTCTTTCTCC	ACTGTCCCACTCCAAACCTG
CDK2	AAGTTGACGGGAGAGGTGGT	TGATGAGGGGAAGAGGAATG
E2F1	AGGGGCAAGGAGCTTTTAAC	CCAAAACCAGAACCCTCTCC
CCNB2	CGACCCTTGCCACTACACTT	TGACTTCCAATACTTCATTCTCTG
CCND3	GGCCGGGGACCGAAACT	CAGTGGCGAAGTGTTTACAAAGT
CDCA3	ACACTACGACAGGGTAAGCGG	AAGGGGTGAGAGGACACAGAT

β-Galactosidase activity

Cells were washed with PBS and fixed with 4% PFA for 15 min at room temperature. After a second wash with PBS, the cells were incubated overnight at 37°C with 5-bromo-4-chloro-3-indolyl-beta-D-galacto-pyranoside (X-gal) solution (Senescence Cells Histochemical Staining Kit, Ref. CX0030-1KT; Sigma) as previously described.²⁷ At least four representative images of each condition were captured using a light microscope at 20X magnification. Quantification was performed using ImageJ software.

Click-it assay

Cell proliferation by tracking new DNA synthesis was performed using the Alexa Fluor 647 Click-IT Assays kit (Thermo Fisher Scientific). Cells were grown to 80% confluence, after which 10 μM 5-ethynyl-2′-deoxyuridine (EdU) was added to the culture media. Cells were incubated at 37°C for 2 h to allow EdU incorporation into DNA during the S phase. Cells were then harvested and fixed by resuspending them in PBS, centrifuging at 500 g for 5 min, and resuspending them in 500 μL of 70% ethanol. For cell permeabilization, cells were washed with 10 mL of 1% BSA (bovine serum albumin) in PBS and resuspended in 200 μL of 0.25% Triton X-100 in PBS. They were incubated for 10 min at room temperature and washed again with BSA.

For intracellular staining, EdU-labeled cells were resuspended in Click-IT reaction mix (200 μL per sample), which tags cells with Alexa 647-Azide, and incubated for 30 min at room temperature. The cells were then washed twice with 1% BSA, resuspended in 500 μL of PI (propidium iodide) supplemented with 0.02 mg/mL RNase A, and incubated for 30 min at room temperature. Finally, the cells were resuspended in 300 μL of FBS. EdU incorporation was analyzed by flow cytometry using FACSCalibur and CytoFLEX S (Beckman Coulter), with 20,000 events acquired per triplicate. Data analysis was performed using FlowJo (v.10.0) software.

Cell death assays

Cells were harvested, and supernatants containing unattached cells were aspirated. Both attached and unattached cells were combined, counted, and 1 million cells were resuspended in 1 mL of flow cytometry buffer. Membrane viability analysis was conducted using double staining with PI/YO-PRO-1. For this, 150 nM YO-PRO-1 (Thermo Fisher Scientific) was added to the cells and incubated for 10 min at 4°C in the dark. Subsequently, 2 μg/mL of PI were added and incubated for 5 min at 4°C in the dark. Samples were analyzed using a FACSCalibur cytometer and CytoFLEX S (Beckman Coulter), with 20,000 events acquired per triplicate. Green fluorescence emitted by YO-PRO-1 was detected at 515 nm, and red fluorescence emitted by PI was detected at 610 nm. Both forward scatter (FSC) and side scatter (SSC) parameters were used to gate live cells and discriminate cell debris, while PI/YO-PRO double-negative cells were considered alive.²⁷ Data analysis was performed using FlowJo (v.10.0) software.

Statistical analysis

Statistical analysis was conducted using GraphPad Prism software (v.8.0.2). Data are presented as mean ± SEM (standard error of the mean). Differences between two groups were assessed using two-tailed unpaired Student’s t test. For comparisons involving more than two groups, one-way ANOVA with Tukey’s correction was employed to evaluate differences among means. Each experiment was replicated independently at least three times to ensure statistical reliability. A p value less than 0.05 was considered statistically significant, denoted as ∗ for p < 0.05, ∗∗ for p < 0.01, and ∗∗∗ for p < 0.001.

Data and code availability

The data that support the findings of this study are available from the corresponding author upon request. Source data are provided with this paper.

Acknowledgments

This work was supported in part through funding from a grant from Ministerio de Ciencia, Innovación y Universidades (MICIU/AEI/10.13039/501100011033): PID2022-137027OB-I00 ERDF/EU, HORIZON-MSCA-2023-SE-01 (EVEREST-101183034), and from EU HORIZON-CSA (TWINFLAG-101079489) (to M.D.M.). P.C.-F. was funded with post-doctoral fellowship (INB606B 2017/014 and IN606C 2021/006) from Xunta de Galicia. A.G.-C. was funded with a predoctoral fellowship (FIS20/00310) from ISCIII. Ministerio de Ciencia, Innovación y Universidades (Spain, PID2021-122335NB-I00, MCIN/AEI/10.13039/50110-0011033/FEDER, UE), Xunta de Galicia (GRC2022/039), and ERDF funds for the financial support (to J.P.S.). We thank members of the CellCOM group for helpful technical suggestions.

Author contributions

P.C.-F., together with A.C.-F., R.M., A.G.-Y. and M.B.-G., designed and performed experiments, analyzed data, or prepared figures. A.G.-C. assisted and performed the flow cytometry experiments. M.M.M., L.A.S., C.P.-C. and J.P.S. designed and synthetized the pyrimidine compounds used in this study. M.D.M. and J.P.S. conceived, directed, and supervised the study. P.C.-F., J.P.S., and M.D.M. wrote the manuscript with input from all co-authors. All authors reviewed the manuscript.

Declaration of interests

The authors declare no competing interests.