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. 2026 Mar 6;21(3):446–456. doi: 10.1021/acschembio.5c00656

Metabolism of Epigenetic Ribonucleosides Leads to Nucleolar Stress and Cytotoxicity

Xuemeng Sun 1, Anita Donlic 2,3, Jacob A Boyer 1,5,6, Theodore E Press 1, Minjae Kim 1, Neal K Reddy 1, Clifford P Brangwynne 2,3,4, Joshua D Rabinowitz 1,5,6, Ralph E Kleiner 1,*
PMCID: PMC13010358  PMID: 41790172

Abstract

Post-transcriptional RNA modifications are ubiquitous in biology, but the fate of epigenetic ribonucleotides after RNA turnover and the consequences of their metabolism and misincorporation into nucleic acids are largely unknown. Here, we explore epigenetic ribonucleoside metabolism in human cells by studying effects on cell growth, quantifying RNA misincorporation and identifying metabolic regulators, and exploring phenotypes associated with cytotoxicity. We find that bulky N6-modified adenosines (i.e., i6A) exhibit high levels of cytotoxicity and RNA misincorporation, whereas cells dramatically restrict the misincorporation of small N6-modified adenosines (i.e., m6A), partly through sanitization by enzymatic deamination, consistent with a recent report. Epigenetic ribopyrimidines also exhibit cytotoxicity, dependent on nucleoside kinase UCK2, but only at much higher concentrations than ribopurines. We further characterize the effects of cytotoxic ribonucleoside metabolism on nucleolar morphology and protein translation. Taken together, our work provides new insights into the metabolism of epigenetic ribonucleosides and mechanisms underlying their cytotoxicity to cells.

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Introduction

Nucleotides are essential for life, serving as the building blocks for RNA and DNA. Beyond their role in nucleic acid synthesis, nucleotides also participate in diverse signaling pathways. It is well established that imbalances in the nucleotide pool can disrupt cellular processes and contribute to various human diseases. −

Cells generate nucleotides through two primary biosynthetic routes: de novo synthesis and salvage pathways. The de novo pathway generates nucleotides from basic precursors including amino acids, sugars, and bicarbonate. Conversely, salvage pathways recycle nucleobases or nucleosides derived from RNA/DNA turnover or from external sources, such as dietary nucleic acids and cellular degradation products. , These two pathways are interconnected, allowing cells to compensate for inhibition of one pathway by upregulating the other. , Notably, the relative contributions of de novo synthesis and salvage vary significantly across tissues.

Because the salvage pathway recycles nucleotides derived from nucleic acid turnover, this pathway is exposed not only to canonical nucleotide structures but also modified epigenetic nucleotides found in RNA/DNA. In particular, over 150 distinct chemical modifications are known on cellular RNA, spanning rRNA, tRNA, mRNA and other noncoding RNAs. , These modifications are highly conserved across all kingdoms of life and are involved in regulating fundamental biological processes through modulation of RNA structure, base pairing, and protein-RNA interactions. ,, Dysregulation of RNA modifications has also been implicated in numerous human diseases, including cancer and neurological disorders. ,,

Among the diverse collection of RNA modifications, those that are restricted in their abundance or exotically structured and likely incompatible with metabolism , are unlikely to make substantial contributions to cellular nucleotide pools. In contrast, a number of modified RNA nucleotides are broadly conserved and abundant (present in concentrations ∼ 1–10% of canonical nucleotides) and structurally similar to canonical nucleotides. These include pseudouridine (Ψ), dihydrouridine (D), 5-methyluridine (m5U), 5-methylcytidine (m5C), and N6-methyladenosine (m6A), among others. Despite the prevalence of RNA modifications, the metabolic fate of modified RNA nucleotides after RNA turnover remains largely unknown. Whether specialized pathways exist to distinguish and process these structures, and what cellular consequences arise from their improper metabolism, are open questions (Figure A).

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Metabolism of epigenetic ribonucleosides. (A) Scheme of potential fates of epigenetic ribonucleosides that originate from post-transcriptionally modified RNA. (B) Structures of modified epigenetic purines and pyrimidines used in this work. (C) Cytotoxicity of epigenetic ribonucleosides (100 μM for 72 h) in WT A549 cells. Cell viability was measured by an MTS-based assay and plot represents mean normalized cell viability ± s. d. (n = 9; three independent biological replicates with three technical replicates for each). (D) Structures of modified N6-adenosine analogues used in this work. (E) Screening of N6-modified adenosine cytotoxicity in a panel of human cell lines. Heatmap shows percentage growth inhibition with 10 μM or 100 μM nucleoside treatment for 72 h, corresponding to Supplementary Figure 1C–J. Cell viability was measured by an MTS-based assay and represents mean normalized by untreated cells. (n = 9 with three technical replicates for each of the three independent biological replicates). (F) RNA modification levels in total RNA after treatment with N6-modified adenosine analogues. WT A549 cells were treated with 10 μM nucleoside for 24 h. Modified nucleotide levels were quantified by nucleoside LC-QQQ-MS. Data are mean ± s.d. (n = 3). Multiple unpaired t test were performed between control and treated. Adjusted p values for e6A: p = 0.0024; for CPA: p = 0.000001; for: i6A: p = 0.00052; for Bz6A: p = 0.000002.

Previous studies investigating the metabolism of epigenetic nucleosides have focused primarily on deoxyribonucleosides. Salvage and misincorporation of the major eukaryotic epigenetic DNA modifications, 5-methylcytosine and its oxidized derivatives, are controlled by multiple metabolic enzymes in the pyrimidine salvage pathway. Direct salvage of 5-methyl-2’-deoxycytidine, the most abundant mark, is restricted by cytidine/uridine monophosphate kinase (CMPK1) and cytidine deaminase (CDA). CMPK1 also restricts salvage of 5-hydroxymethyl-2’-deoxycytidine (5hmdC) and higher oxidation states (i.e., formyl, carboxy); however, deamination to 5-hydroxymethyl-2’-deoxyuridine (5hmdU) in cells with high CDA activity results in DNA misincorporation, DNA damage, and cytotoxicity. Cells additionally express 2’-deoxynucleoside 5′-monophosphate N-glycosidase (DNPH1), a sanitization enzyme that processes 5hmdU monophosphate (5hmdUMP) and prevents its DNA misincorporation.

An analogous sanitization pathway has been proposed for the abundant epigenetic ribonucleoside N6-methyladenosine (m6A). Studies in Arabidopsis thaliana and human cells identified N6-methyl-AMP deaminase (ADAL) as an enzyme that deaminates N6-methyl-AMP (m6AMP) and N6-methyl-2’-deoxyAMP (m6dAMP) into their corresponding inosine derivatives. These findings suggest that ADAL prevents metabolism and misincorporation of N6-methyladenosine into DNA , and RNA; however, direct evidence for a role in preventing RNA misincorporation is lacking. Additional studies of epigenetic ribonucleosides have found that extracellular N6-isopentenyladenosine (i6A) is cytotoxic to mammalian cells and can be incorporated into cellular RNA. While our manuscript was in review, Ogawa et al. reported that the epigenetic N6-modified adenosine analogs m6A, N 6, N 6-dimethyladenosine (m6 , 6A), and i6A are sequentially phosphorylated by ADK and deaminated by ADAL to mitigate their cytotoxicity, linking modified adenosine metabolism to cellular stress and disease phenotypes. Beyond N6-modified adenosines, in Arabidopsis, loss of nucleoside hydrolase 1 (NSH1) correlates with accumulation of 5-methyluridine (m5U) into mRNA and reduced seedling growth, however mammalian NSH1 homologues have yet to be identified. Overall, the metabolic pathways mediating the activation and restriction of diverse epigenetic ribonucleosides and the consequences of RNA misincorporation of these structures are poorly annotated and understood.

In this study, we investigated a panel of epigenetic pyrimidine and purine ribonucleosides in cultured human cells. We assessed nucleoside cytotoxicity and RNA misincorporation using cell growth assays and RNA nucleoside mass spectrometry. Furthermore, by employing CRISPR-Cas9 knockouts, we examined the roles of nucleotide salvage enzymes in epigenetic ribonucleoside metabolism. Finally, we evaluated the effects of exogenous ribonucleoside treatment on nucleolar structure and protein translation. We found that purine analogs were generally more toxic than pyrimidine ribonucleosides and characterized the role of ADAL in the restriction of diverse N6-modified adenosine incorporation and cytotoxicity and the role of uridine cytidine kinase 2 (UCK2) in the metabolic activation of pyrimidines. Together, our work provides new insights into the metabolic fate and cellular impact of epigenetic ribonucleosides.

Results

Screening Cytotoxicity of Epigenetic Ribonucleosides

To investigate whether epigenetic ribonucleosides can be metabolized and perturb normal biological processes in mammalian cells, we assembled a panel of 18 biologically abundant modified and unmodified ribonucleosides, comprising 8 purines and 10 pyrimidines (Figure B). We screened each nucleoside for cytotoxicity in A549 cells, a human nonsmall cell lung cancer (NSCLC) line, by adding compounds into normal growth medium at 100 μM concentration and evaluating cell proliferation after 72 h. Among the canonical ribonucleosides (A, G, C, and U), only G significantly inhibited cell proliferation, reducing cell growth by 56.2% ± 3.24% at 100 μM (Figure C), consistent with a previous report. None of the epigenetic ribopyrimidines tested showed measurable cytotoxicity at 100 μM. Among the modified ribopurines, N7-methylguanosine (m7G), N1-methyladenosine (m1A), and inosine (I) were nontoxic. In contrast, bulky N6-modified adenosines such as N6-isopentenyladenosine (i6A) and N6, N6-dimethyladenosine (m6,6A), exhibited strong cytotoxic effects (Figure C). We further evaluated the cytotoxicity of i6A and m6,6A in A549 cells and found that these compounds exhibited cytotoxicity between 1 and 10 μM concentration (Supplementary Figure 1A and 1B), aligning with previous reports in diverse human cell lines. ,, Interestingly, treatment with the structurally related ribonucleoside N6-methyladenosine (m6A) did not significantly affect the viability of A549 cells under the conditions tested (Figure C).

Bulky N6-Modified Adenosines Exhibit Cytotoxicity and RNA Incorporation

Given the differences observed among various N6-modified adenosines, we expanded our panel to include synthetic adenosine analogues with diverse N6-substituents (Figure D): N6-ethyladenosine (e6A), N6-cyclopentyladenosine (CPA), and N6-benzyladenosine (Bz6A). We then tested these analogues together with epigenetic N6-modified adenosines at 10 μM and 100 μM concentration for cytotoxicity across multiple (primarily cancer-derived) cell lines, including K562, HeLa, U2OS, HEK293T, HepG2, MCF7, A549 and MDA-MB-231. A consistent pattern emerged across most cell lines: m6,6A, i6A and Bz6A exhibited strong cytotoxicity, reducing cell viability by more than 80% at 100 μM (Figure E and Supplementary Figure 1C–J), whereas m6A, e6A, and CPA were generally much less toxic at the same concentration. We did observe some differences in nucleoside sensitivity across cell lines. For example, HeLa cells were significantly less sensitive to all assayed N6-modified adenosines (Figure E and Supplementary Figure 1C–J), whereas HepG2 and K562 were most sensitive to m6,6A, i6A, and Bz6A. In addition, m6A demonstrated modest toxicity in HEK293T and K562. These differences may stem from variation in metabolic flux or signaling pathways related to nucleic acid quality control or cellular stress across individual cell types. ,

To investigate whether N6-modfied nucleosides are metabolized and incorporated into cellular RNA, we performed quantitative nucleoside LC-QQQ-MS after nucleoside feeding in A549 cells (Supplementary Figures 2 and 3). We could not detect changes in m6,6A or m6A levels after nucleoside feeding, however the high endogenous abundance of these modifications in total RNA (∼500–1000 ppm) precludes reliable measurement of low-level metabolic incorporation. In contrast, we measured clear incorporation of i6A into total RNA, rising by 830 ± 99.0 ppm, a ∼ 2-fold increase over endogenous levels (Figure F). We measured similar i6A incorporation in HeLa cells, although it required higher treatment concentration than in A549, in line with the reduced cytotoxicity of i6A in HeLa (Supplementary Figure 4A and Figure E). For synthetic adenosine analogues, we detected low-level RNA incorporation for e6A (7.3 ± 1.4 ppm) and CPA (22.7 ± 0.5 ppm), and higher RNA incorporation for Bz6A (150.9 ± 4.3 ppm) (Figure F). These results indicate that the cytotoxicity of N6-modified adenosine analogues is correlated with the extent of their incorporation into RNA. Interestingly, bulkier N6-modifications, such as those found in Bz6A and i6A, are incorporated more efficiently into RNA than N6-modified adenosines with small alkyl substituents. This stands in contrast with typical metabolic incorporation trends that favor metabolic activation of nucleotide precursors more similar in size to canonical structures, , presumably due to increased substrate compatibility with nucleotide salvage enzymes and RNA polymerases.

ADK Activates N6-Modified Adenosine Analogues

To understand the metabolic pathways mediating modified adenosine activation and RNA incorporation, we investigated genes in the adenosine salvage pathway. Adenosine nucleosides can be catabolized by adenosine deaminase (ADA) followed by purine nucleoside phosphorylase (PNPase), or phosphorylated by adenosine kinase (ADK) to adenosine monophosphate (AMP), which is then further metabolized to ATP, or alternatively deaminated by AMP deaminase (AMPD) to inosine monophosphate (IMP) (Figure A). Since ADK is the canonical enzyme responsible for phosphorylation of adenosine to AMP in mammals, , and deamination by ADA would remove the N6-substituent, we hypothesized that ADK is involved in N6-modified adenosine toxicity and RNA misincorporation. We therefore generated a population knockout (KO) of ADK in A549 cells (Supplementary Figure 5A,B) using CRISPR/Cas9 methods. Strikingly, depletion of ADK resulted in near complete reversal of cytotoxicity induced by N6-modified adenosine nucleosides, including e6A, m6,6A, i6A, and Bz6A (Figure B). Consistent with this finding, we observed a dramatic 80.9% ± 14.7% reduction of i6A and a 98.9% ± 2.0% reduction of Bz6A in RNA incorporation in ADK KO cells compared with WT control (Figure C and Supplementary Figure 4B). Further, we generated a clonal ADK KO cell line in HEK293T parent cells (Supplementary Figure 5C) and also observed resistance to the cytotoxic effects of N6-modified adenosines, which could be reversed by re-expression of WT ADK via plasmid transfection (Supplementary Figure 5D). Together, these data demonstrate that ADK is required for RNA incorporation and associated cytotoxicity of N6-modified adenosine analogues, likely by mediating their direct phosphorylation to N6-modified AMP nucleotides. This metabolic activation likely underlies their cytotoxic effects, further supporting a direct link between nucleoside metabolism, RNA incorporation, and cytotoxicity.

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Metabolism of N6-modified adenosine analogues. (A) Enzymatic pathways involved in adenosine nucleoside metabolism. (B) Quantification of cell growth for A549 WT and ADK KO cells after treatment with N6-modified adenosines (100 μM for 72 h). Cell growth was measured using an MTS-based assay. Plot represents mean normalized cell viability ± s. d. (n = 9; three independent biological replicates with three technical replicates for each). Multiple unpaired t test were performed between WT and ADK KO. Adjusted p values for m6A: p < 0.000001; for e6A: p = 0.12; for m6,6A: p < 0.000001; for i6A: p < 0.000001; for Bz6A: p < 0.000001. (C) RNA incorporation for i6A and Bz6A in A549 WT and ADK KO cells. Cells were treated with 10 μM nucleoside for 24 h and RNA modification levels in total RNA were analyzed by nucleoside LC-QQQ-MS. Data are mean ± s.d. (n = 3). The unnormalized data is shown in Supplementary Figure 4B. Multiple unpaired t test were performed between WT and ADK KO. Adjusted p values for i6A: 0.001651; for Bz6A: 0.000014. (D) Quantification of cell growth for HEK293T WT and ADAL KO2/KO6 cells after treatment with N6-modified adenosine analogues (10 μM for 72 h for all analogues except m6A, which was performed at 50 μM for 72 h). Cell growth was measured using an MTS-based assay. Plot represents mean normalized cell viability ± s. d. (n = 9; three independent biological replicates with three technical replicates for each). Multiple unpaired t test were performed between WT and ADAL KOs. Adjusted p values for m6A: WT vs KO2 and WT vs KO6: p < 0.000001; for e6A: WT vs KO2 p < 0.000001; WT vs KO6 p < 0.000001; for m6,6A: WT vs KO2 p < 0.000001; WT vs KO6 p < 0.000001; for i6A: WT vs KO2 p = 0.000008; WT vs KO6 p = 0.000003; for Bz6A: WT vs KO2 p < 0.000001; WT vs KO6 p < 0.000001. (E) RNA incorporation for N6-modified adenosine analogues in HEK293T WT and ADAL KO6. Cells were treated with nucleosides (10 μM for 24 h) and modified nucleotide levels in total RNA were analyzed by nucleoside LC-QQQ-MS. Data are mean ± s.d. (n = 3). The unnormalized data is shown in Supplementary Figure 4C. Multiple unpaired t test were performed between WT and ADAL KO6. Adjusted p values for m6A-D3: p = 0.000008; for e6A: p = 0.000003; for i6A: p = 0.000018; for Bz6A: p = 0.000117. (F) RNA incorporation of m6A-D3 and i6A in HEK293T WT and ADAL KO6 with or without ADAi treatment (10 μM pentostatin). ADAi treated cells were pretreated with 10 μM pentostatin for 6 h and then exposed to both nucleosides (10 μM) and pentostatin overnight. Cells without ADAi treatment were only incubated with 10 μM nucleosides overnight and harvested together with the treated samples. Modified nucleotide levels in total RNA were quantified by nucleoside LC-QQQ-MS. Data are mean ± s.d. Three independent biological replicates were assayed for i6A treated cells. For m6A-D3 feeding, WT without ADAi: n = 6; ADAL KO6 without ADAi: n = 5; WT or ADAL KO6 with ADAi: n = 3. The unnormalized data is shown in Supplementary Figure 8D,E. Multiple unpaired t test were performed between ADAi treated and untreated samples. Adjusted p-values for m6A-D3: p = 0.00056; for i6A: p = 0.03. (G) Scheme that summarizes the trend of cytotoxicity and RNA incorporation with modification group size at N6 position.

Sanitization of N6-Modified Adenosines by ADAL and ADA

Since the metabolic incorporation of N6-modified adenosines followed an unexpected structural trend (i.e., larger derivatives were incorporated more readily than smaller derivatives), we explored whether selective sanitization pathways may exist for smaller N6-modified adenosine nucleosides that closely resemble epigenetic nucleosides (i.e., m6A). An active sanitization pathway for N6-modified adenosines that preferentially removes smaller nucleotides from the nucleotide pool would explain the inefficient metabolism and RNA incorporation of these compounds. We focused our efforts on adenosine deaminase-like protein (ADAL) (Figure A), as previous studies have shown that ADAL can deaminate m6AMP and m6dAMP and prevent misincorporation of N6-methyladenine into DNA; , whether ADAL prevents the misincorporation of N6-modified adenosines into RNA is poorly understood. We investigated the role of ADAL in N6-modified adenosine sanitization by generating clonal ADAL KO cell lines in HEK293T and HeLa (Supplementary Figure 6A-C, Supplementary Figure 7A,B). ADAL KO cells showed no growth defects (Supplementary Figure 6D) and no disruption in endogenous levels of m6A, m6,6A, or i6A (Supplementary Figure 6E). However, deletion of ADAL sensitized cells to all five N6-modified adenosine analogues tested, as observed in two independent HEK293T ADAL KO lines and two independent HeLa KO lines (Figure D and Supplementary Figure 7C). These effects could be rescued by re-expression of WT ADAL enzyme by plasmid transfection (Supplementary Figure 7D). As the effects were more pronounced in HEK293T, we further focused on these cell lines. Consistent with increased cytotoxicity, ADAL KO also led to increased RNA incorporation of all five analogues (Figure E, Supplementary Figure 4C,D) with 7-fold or more increase over WT for m6A-D3 (+17.5 ± 0.8 ppm) and e6A (+14.0 ± 0.5 ppm), and ∼ 3-fold increase over WT for i6A (+782 ± 47.6 ppm) and Bz6A (+420 ± 48.6 ppm). We also found that m6A-D3 incorporation in poly­(A) RNA was comparable to that in total RNA (primarily rRNA/tRNA) in ADAL KO cells (Supplementary Figure 4E), suggesting that it can be incorporated into RNA by different polymerases.

Consistent with previous reports showing that ADAL prevents misincorporation of ribonucleotide m6A into genomic DNA, , we measured elevated levels of m6dA in our ADAL KO cells that were not fed exogenous m6A ribonucleoside (Supplementary Figure 6F). Interestingly, we were not able to measure increased m6dA levels in the genome of WT or ADAL KO cells upon feeding exogenous isotopically labeled m6A-D3 at 50 μM for 2 days (data not shown). We speculate that longer-term feeding or higher nucleoside concentrations would be required to see perturbation of genomic m6dA levels; indeed previous studies used prolonged incubation with m6A-D3 to generate detectable levels in genomic DNA.

Whereas ADAL KO sensitized cells to diverse N6-modified adenosines and concomitantly increased their incorporation into RNA, incorporation of small alkyl-substituted m6A and e6A into RNA still proceeded ∼ 30–70-fold less efficiently than larger i6A and Bz6A derivatives in the ADAL KO lines. We therefore explored whether sanitization pathways upstream of ADAL, such as deamination by nucleoside deaminases (i.e., ADA1, ADA2), may prevent the metabolism of small N6-modified nucleosides. Notably, ADA1 shares a high degree of structurally similarity with ADAL, with a root-mean-square-deviation (RMSD) of only 1.56 Å (Supplementary Figure 8A–C)). To inhibit the activity of ADA proteins, we treated WT and ADAL KO cells with pentostatin, an established and potent small molecule inhibitor of ADA1/2 enzyme, , and measured RNA incorporation of m6A-D3 and i6A nucleosides. Pentostatin treatment modestly increased RNA incorporation of m6A-D3 in ADAL KO cells (82% increase), but not in WT cells, and had minimal effect on incorporation of i6A (19% increase in KO) (Figure F and Supplementary Figure 8D,E). Collectively, our results indicate that diverse N6-modified adenosine analogues are activated by ADK and incorporated into RNA. Bulky N6-modified adenosines exhibit the highest levels of cytoxicity and RNA incorporation, and cells can mitigate misincorporation and cytotoxicity of N6-modified adenosine analogues through sanitization pathways involving nucleotide and nucleoside deaminases, ADAL and ADA (Figure G); however, even in the absence of ADAL/ADA-mediated sanitization pathways, m6A and structurally related derivatives show limited RNA misincorporation, suggesting that other mechanisms exist to prevent stochastic RNA incorporation of epigenetic N6-modified adenosines.

Cytotoxicity of Epigenetic Pyrimidine Ribonucleosides

Although epigenetic ribopyrimidines did not show cytotoxic effects in our initial screen at 100 μM concentration in A549 cells (Figure C), we repeated the screen at higher concentrations and expanded to additional human cell lines. In A549, we began to see cytotoxic effects at 5 mM concentration (Figure B and Supplementary Figure 9A). Among the compounds tested, dihydrouridine (D) inhibited cell growth by 83.7% ± 1.3%, 5-methyluridine (m5U) by 50.3% ± 4.0%, 5-methylcytidine (m5C) by over 90%, and 2’-O-methylcytidine (Cm) by 63.8% ± 2.6% (Figure B). These results indicate that epigenetic ribopyrimidines can be cytotoxic, but only at high (millimolar) concentrations. We then extended the assay to a panel of cell lines. At 1 mM treatment, m5C consistently exhibited the strongest cytotoxicity, inhibiting growth by over 90% in HeLa and HEK293T cells (Figure C and Supplementary Figure 9C–H). Interestingly, pseudouridine (Ψ) showed cell-type specific effects, with more than 90% inhibition in HeLa cells, but minimal toxicity in A549 and MCF7. Moderate inhibition (∼30%) was observed in HepG2, U2OS and HEK293T cells. When the treatment concentration was further increased to 5 mM, the cytotoxicity patterns become more pronounced: D, m5C, and m5U were broadly toxic across most cell lines, while Ψ remained selectively toxic in specific cell types (Supplementary Figure 9B–H).

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Metabolism of modified ribopyrimidine analogues. (A) Metabolic pathways involved in ribopyrimidine metabolism. (B) Cytotoxicity of epigenetic ribonucleosides (5 mM for 72 h) in WT A549 cells. Cell viability was measured by MTS assay and plot represents mean normalized cell viability ± s. d. (n = 9; three independent biological replicates with three technical replicates for each). (C) Ribopyrimidine nucleoside cytotoxicity in a panel of human cell lines. Heatmap shows percentage growth inhibition (1 mM nucleoside for 72 h), corresponding to Supplementary Figure 9A and C–H. Cell viability was measured by an MTS-based assay and represents mean normalized by untreated cells. (n = 9 with three technical replicates for each of the three independent biological replicates). (D) Quantification of cell growth for A549 WT and UCK2 KO cells with m5C, m5U or D treatment. Cell growth was measured by MTS assay after nucleoside treatment (5 mM for 72 h). Plot represents mean normalized cell viability ± s. d. (n = 9; three independent biological replicates with three technical replicates for each). Multiple unpaired t test were performed between WT and UCK2 KO. Adjusted p values for m5C: p < 0.000001; for D: p < 0.000001. (E) RNA incorporation levels for m5C, m5U, or D. A549 WT or UCK2 KO cells were treated with 1 mM nucleoside for 24 h and RNA modification levels in total RNA were quantified by nucleoside LC-QQQ-MS. Data are mean ± s.d. (n = 3). The unnormalized data is shown in Supplementary Figure 10E. Multiple unpaired t test were performed between WT and UCK2 KO. Adjusted p value for m5C: p = 0.000004. (F) Quantification of cell growth for HeLa WT and UCK2 KO cells after treatment with m5C, m5U or Ψ (1 mM for 72 h). Cell growth was measured by MTS assay. Plot represents mean normalized cell viability ± s. d. (n = 9; three independent biological replicates with three technical replicates for each). Multiple unpaired t test were performed between WT and UCK2 KO. Adjusted p values for m5C: p < 0.000001; for m5U: p = 0.000009; for Ψ p < 0.000001. (G) RNA incorporation levels for m5C, m5U or Ψ in HeLa WT or UCK2 KO cells. Modified nucleotide levels were quantified in total RNA by nucleoside LC-QQQ-MS after m5C/ Ψ (0.5 mM) or m5U (1.5 mM) treatment for 24 h. Data are mean ± s.d. (n = 3). Unnormalized data is shown in Supplementary Figure 11D. Multiple unpaired t test were performed between WT and UCK2 KO. Adjusted p values for m5C: p < 0.000001; for m5U: p = 0.00045; for Ψ: p = 0.018.

UCK2 Mediates the Incorporation and Toxicity of Epigenetic Ribopyrimidines

Our group − and others − have previously established that uridine-cytidine kinase 2 (UCK2), which phosphorylates uridine/cytidine nucleosides to NMPs, is rate-limiting for uptake and RNA incorporation of modified pyrimidine ribonucleosides (Figure A). To explore its role in metabolism of epigenetic ribopyrimidines, we generated UCK2 KO cells in A549 (Supplementary Figure 10A) and HeLa (Supplementary Figure 11A) and assayed sensitivity to D, m5C and m5U. In A549, UCK2 depletion markedly rescued the cytotoxicity induced by m5C, while no significant rescue was observed for D or m5U treatments (Figure D and Supplementary Figure 10B,C). Through dose–response titrations, we observed >7-fold increase in the IC50 of m5C in A549 cells with UCK2 depletion. In contrast, the IC50 values for m5U and D remained nearly unchanged between UCK2 KO and WT cells (Supplementary Figure 10D). Using nucleoside LC-MS/MS, we quantified RNA incorporation (Supplementary Figure 12) and detected 3.65% ± 0.07% incorporation of m5C in WT A549 cells (a 4-fold increase over endogenous m5C levels), compared to only 0.38% ± 0.096% incorporation in UCK2 KO cells (Figure E and Supplementary Figure 10E). In contrast, D and m5U levels did not significantly change between UCK2 KO and WT (Figure E and Supplementary Figure 10E), consistent with similar cytotoxicity profiles (Figure D).

Similar trends were observed in HeLa cells. UCK2 KO rescued m5C-induced toxicity but not m5U toxicity (Figure F and Supplementary Figure 11B), suggesting m5U is a poor substrate for UCK2. As noted above, HeLa cells were selectively sensitive to Ψ, an effect that we found to be dependent on UCK2 status (Figure F and Supplementary Figure 11B,C). This effect was less pronounced but still evident in A549 cells (Supplementary Figure 10C). IC50 values for m5C and Ψ in HeLa UCK2 KO cells increased ∼4–5-fold as compared to WT cells (Supplementary Figure 11C). In WT HeLa cells, LC-QQQ-MS quantification revealed UCK2-dependent incorporation of m5C at 3.83% ± 0.07% and Ψ at 2.72% ± 0.49% following 0.5 mM treatment in WT HeLa (Figure G and Supplementary Figure 11D). Surprisingly, although m5U did not show UCK2-dependent toxicity in HeLa, we detected UCK2-dependent RNA incorporation: m5U levels increased by ∼ 14.5-fold in WT (2.41% ± 0.20% incorporation) and by only 5-fold in UCK2 KO (0.86% ± 0.08% incorporation) (Figure G and Supplementary Figure 11D). While UCK2 depletion did not rescue m5U toxicity, the underlying mechanism remains unclear. It is possible that metabolism-independent processes or alternative UCK2-independent metabolic pathways contribute to m5U cytotoxicity. Further, we demonstrated rescue of pyrimidine cytotoxicity in UCK2 KO cells by re-expression of WT UCK2 by plasmid transfection (Supplementary Figure 11E). Taken together, these results demonstrate that UCK2 is the key enzyme responsible for m5C and Ψ toxicity via its role in promoting their RNA incorporation. In contrast, m5U and D show minimal UCK2 dependence, suggesting distinct metabolism or cytotoxicity stemming from the unprocessed nucleosides.

Pyrimidine Ribonucleoside Cytotoxicity Induces DNA Damage but Is Not Associated with DNA Incorporation

We next investigated the mechanisms by which modified ribonucleosides cause cell death. Given the high concentrations of epigenetic pyrimidine ribonucleosides needed for cytotoxicity, and associated high levels of RNA incorporation, we questioned whether modified pyrimidines fed in this way may also be metabolized to modified deoxynucleotides and possibly incorporated into DNA, thereby inducing DNA damage or replication stress/inhibition. First, we investigated whether modified ribonucleoside treatment induced DNA damage in A549 or HeLa cells. In HeLa cells, treatment with m5C, Ψ, or m5U for 48h at concentrations that induced comparable RNA incorporation (∼3%) resulted in DNA damage as measured by γH2AX Western blot, whereas i6A treatment did not increase γH2AX signal in either cell line (Supplementary Figure 13A,B). In A549, γH2AX induction was more modest; however, m5C treatment did result in a slight increase in DNA damage-associated signal. We next focused on quantifying DNA incorporation and deoxynucleotide metabolites derived from m5C because it was the most toxic epigenetic ribopyrimidine we assayed, it induced DNA damage in HeLa and A549 cells, and corresponding modified deoxyribonucleoside standards for LC-MS analysis were available. We extracted genomic DNA from treated and untreated cells and analyzed 5-methyl-2’-deoxycytidine (5mdC) levels by nucleoside LC-QQQ-MS. Upon m5C feeding, we did not observe significant differences in 5mdC levels in either A549 or HeLa cells (Supplementary Figure 13C,D). However, considering the high endogenous abundance of 5mdC in genomic DNA, it is possible that subtle changes might be masked in bulk LC-QQQ-MS analysis. Further, we examined the nucleotide pools and found no detectable formation of m5dCTP after 2 or 24 h m5C treatments (Supplementary Figure 14). Instead, we observed a marked accumulation of m5C monophosphate (m5CMP) and m5C triphosphate (m5CTP), as well as the corresponding deamination products, m5U monophosphate (m5UMP) and m5U triphosphate (m5UTP) (Supplementary Figure 14), aligning with increases in m5C in RNA (Figure G and Supplementary Figure 11D). The abundance of m5C and m5U-derived nucleotide was higher 2 h postfeeding than 24 h, indicating that cells were actively metabolizing these molecules. In contrast, whereas m5C nucleoside levels also declined from 2 to 24 h, m5U nucleoside and 5-methyluracil (thymine) accumulated in the cell from 2 to 24 h. Taken together, our findings show that epigenetic ribopyrimidine-induced cytotoxicity correlates with DNA damage but appears to be independent of metabolism to modified deoxynucleotides or DNA incorporation.

i6A and Ψ Induce Nucleolar Stress through Distinct Mechanisms

Active rDNA transcription is essential for nucleolar organization, and nucleolar function is tightly linked to its structure as a multiphase condensate. − The mammalian nucleolus consists of three subcompartments: the fibrillar center (FC), dense fibrillar component (DFC) and granular component (GC). rDNA is transcribed at the FC-DFC interface, pre-rRNA processing occurs in the DFC, and ribosomal subunit assembly takes place in the GC. − Inhibition of rDNA transcription triggers nucleolar stress, characterized by redistribution of nucleolar proteins to the periphery, leading to a rounded, condensed appearance. − This can be visualized using markers specific to different nucleolar compartments. ,

To examine whether epigenetic ribonucleoside cytotoxicity and RNA incorporation correlate with perturbation of nucleolar structure and function, which could lead to defects in ribosome biogenesis and protein translation and arrest cell cycle progression, we examined nucleolar morphology by staining for nucleophosmin (NPM1), a GC marker, and RPA-194 (RNA Polymerase I Subunit A), an FC marker. In HeLa cells treated with i6A treatment, we observed nucleolar caps forming around the GC (Figure A), a hallmark of nucleolar segregation caused by Pol I inhibition. , Quantification of nucleolar eccentricity showed a 27% decrease (Figure B), reflecting nucleolar rounding in the images (Figure A) and consistent with Pol I inhibition. , In addition, NPM1 partitioning was significantly altered by i6A treatment (Figure C). While Ψ treatment did not induce nucleolar caps, it did cause nucleolar rounding, as evidenced by a 14% decrease in eccentricity (Figure A,B). These morphological changes suggest that Ψ also induces nucleolar stress, albeit likely through a mechanism distinct from Pol I inhibition. In contrast, m5C treatment had no obvious effect on nucleolar morphology (Figure A,B).

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Epigenetic nucleosides induce nucleolar stress and global translation inhibition. (A) Confocal micrographs of HeLa cells treated with 100 μM i6A, 500 μM m5C or 500 μM Ψ for 24h. Cells were stained with antibodies for NPM1 and RPA-194 and a nuclear dye. Scale bar represents 10 μm. Representative images from two biological replicates are shown. (B) Quantification of nucleolus morphology (eccentricity). Boxes in the plot represent 25th to 75th percentiles and whiskers show min to max. Two tailed unpaired t test was performed between treated and control samples and p values for i6A: p < 0.000001; Ψ (0.5 mM): p = 0.000011; Ψ (0.3 mM): p = 0.047; 40–120 cells were analyzed per condition. (C) Quantification of NPM1 partitioning. Boxes in the plot represent 25th to 75th percentiles and whiskers show min to max. Two tailed unpaired t test was performed between treated and control samples and p values for i6A: p < 0.000001; Ψ (0.5 mM): p = 0.0062; m5C: p < 0.000001. Between 40 and 300 cells were analyzed per condition. (D) Global translation levels upon nucleoside feeding. Quantification of OP-puro incorporation (Cy3 signal) from HeLa cells treated with 100 μM i6A, 500 μM m5C, or 500 μM Ψ for 24h. Boxes in the plot represent 25th to 75th percentiles and whiskers show min to max. Two tailed unpaired t test were performed between treated and control samples and p values are for i6A: p < 0.000001; for m5C: p < 0.000001; for Ψ: p < 0.000001.

We observed similar nucleolar perturbation with i6A treatment in A549 cells (Supplementary Figure 15A), though changes in eccentricity and NPM1 partitioning were less pronounced (Supplementary Figure 15B,C). These differences likely reflect cell-type-specific responses. Nonetheless, the consistent formation of nucleolar caps upon i6A treatment in both HeLa and A549 cells supports its role in Pol I inhibition. In contrast, m5C treatment did not induce noticeable alterations in A549 cells (Supplementary Figure 15), consistent with observations in HeLa cells (Figure A). Together, these findings suggest that i6A inhibits cell growth by inhibiting Pol I and inducing nucleolar stress, presumably by first misincorporating into rRNA. Ψ also induces nucleolar stress, possibly by interfering with ribosome biogenesis at the level of pre-rRNA processing or modification, given the role of pseudouridylation in these processes. The absence of these effects with m5C suggests its cytotoxicity operates through mechanisms unrelated to nucleolar function.

Epigenetic Nucleosides Inhibit Global Translation

Finally, we employed O-propargyl-puromycin (OP-puro), which incorporates into nascent polypeptides, to assess global protein synthesis after epigenetic nucleoside treatment. We observed a marked reduction in OP-puro signal following epigenetic nucleoside treatments: 27.7% ± 1.4% (mean ± SEM) decrease for m5C, 22.9% ± 1.2% (mean ± SEM) decrease for Ψ, and 8.6% ± 1.4% (mean ± SEM) decrease for i6A (Figure D and Supplementary Figure 16). These results indicated that all three epigenetic nucleosides disrupt global translation to varying extents.

Discussion

In this study, we show that a subset of epigenetic ribonucleoside can be metabolized by cells and incorporated into RNA, with associated cytotoxicity. We found that N6-modified adenosines were the most toxic of those tested and further showed that their metabolic activation relies upon ADK and is opposed by ADAL- and ADA-mediated sanitization pathways. Using a broader panel of epigenetic and synthetic N6-modified adenosines, we found a strong restriction against the misincorporation of adenosine analogues with small N6-alkyl substitutions such as the abundant epigenetic ribonucleoside m6A. Compared to purine analogues, pyrimidines were less toxic but demonstrated measurable UCK2-dependent and UCK2-independent cytotoxicity at high concentrations in multiple cell lines. Furthermore, we show that i6A and Ψ misincorporation correlates with nucleolar stress and global translation inhibition.

Our findings on the role of ADAL in deaminating N6-methyladenosine nucleotides align with previous work in Arabidopsis and human cells, where ADAL enzymes were shown to serve a similar protective function. Additionally, we show that ADAL has broad substrate specificity and can process native and synthetic N6-modified adenosines including i6A, Bz6A, and e6A, and prevent their misincorporation into RNA. Similarly, ADK possesses broad substrate specificity and can activate diverse functionalized N6-modified adenosines. While this manuscript was in preparation, similar findings were reported by Ogawa et al. By measuring RNA misincorporation rates of different N6-modified adenosines, we found that larger N6-substituents, such as isopentenyl and benzyl, exhibited incorporation at >100-fold higher levels than small alkyl N6-substitutions (i.e., methyl and ethyl) (Figure E). This is in stark contrast to typical metabolic incorporation trends for modified nucleosides that favor activation and incorporation of nucleosides that more closely resemble canonical nucleoside structures. While the precise mechanism underlying the preference for misincorporation of larger N6-modified adenosines remains to be determined, we speculate that it likely results from a combination of factors including the ability of larger analogues to evade catabolic adenosine pathways as well as additional layers of selectivity which may include nucleoside transporters, RNA polymerases, and sanitization pathways functioning downstream of RNA synthesis. One possible explanation is that ADAL more efficiently deaminates small N6-modified adenosines, as was suggested by an in vitro report. However, even in ADAL KO cells larger N6-modified adenosines still incorporate into RNA at >100-fold higher levels than m6A or e6A. This suggests that ADAL is not the only (and perhaps not even the major) restriction mechanism for preventing m6A misincorporation. We show that adenosine deaminase (ADA) also serves a role (albeit a minor one) in sanitizing exogenous m6A nucleosides. Additional studies will be required to disentangle the relative contributions of these enzymes and pathways to modified adenosine metabolism and RNA misincorporation.

In contrast to purine ribonucleosides, pyrimidines (both modified and unmodified) did not exhibit cytotoxicity except at millimolar concentrations. At these high concentrations, cytotoxicity of epigenetic pyrimidine ribonucleosides largely depended upon UCK2, which aligns with previous studies implicating UCK2 as the rate-limiting enzyme for salvage of cytotoxic pyrimidine ribonucleoside analogues. − UCK2 is known to have stringent substrate selectivity , and therefore this kinase (and NMP kinase CMPK1) may serve as a filter to prevent metabolism and subsequent RNA misincorporation of epigenetic pyrimidines. Accordingly, pyrimidine toxicity may only emerge when this filter capacity is exceeded at supraphysiological concentrations, allowing sufficient nucleotide accumulation and RNA misincorporation to perturb RNA function. Whereas we measured high levels of RNA misincorporation correlating with cytotoxicity, it is unknown whether cells tolerate lower concentrations of these nucleosides due to inefficient metabolism (or efficient sanitization) or lack of major perturbation to RNA structure/function upon misincorporation of pyrimidine modifications. Catabolic enzymes for Ψ (the most abundant RNA modification) have been reported in bacteria and in plants, but homologous pathways in mammals are not known.

We evaluated the cytotoxicity mechanisms of epigenetic ribonucleosides by characterizing nucleolar morphology and measuring protein translation. Several synthetic nucleosides and nucleobases are known to induce nucleolar stress, including 5-fluorouracil, 5-fluorouridine, and 4-thiouridine. Additionally, DHODH inhibitors have been shown to disrupt nucleotide balance and trigger nucleolar rounding. In our studies, modified epigenetic ribonucleosides can similarly perturb nucleolar function but through diverse mechanisms. Treatment with i6A induces nucleolar caps, consistent with Pol I inhibition. Yakita et al. also demonstrated that i6A inhibits Pol I transcription by RT-qPCR analysis of rRNA transcripts. Since i6A misincorporates into RNA, we speculate that it may serve as a transcription terminator. In contrast, Ψ induces nucleolar stress without significant nucleolar cap formation, suggesting that Ψ misincorporation may perturb later steps of rRNA processing and ribosome biogenesis, rather than inhibiting Pol I transcription. Consistent with this idea, studies in E. coli have shown that excessive Ψ in rRNA can impair ribosome assembly. Future experiments measuring the accumulation of pre-rRNA intermediates will be required to directly test this mechanism in mammalian cells. Whereas m5C did not induce apparent nucleolar stress based on FC and GC morphology, it did inhibit protein translation (as did i6A and Ψ). Therefore, m5C may act at later stages of ribosome biogenesis, or translation inhibition may be due to mRNA misincorporation; indeed, m5C modifications in synthetic mRNA are known to perturb translation efficiency. It is important to note that whereas our study demonstrates a clear correlation between RNA misincorporation and cytotoxicity, we also cannot exclude the possibility that a modified nucleotide metabolite (rather than improperly modified RNA) is the cytotoxic species in cells. Pinning down this connection will require additional mechanistic studies in the future.

In summary, our study provides new insights into the cellular metabolism of epigenetic ribonucleosides and highlights the role of nucleotide salvage and sanitization pathways in managing their incorporation into RNA. These findings deepen our understanding of the metabolic fate of RNA modifications following RNA turnover and offer valuable insights into the cellular handling of nucleoside-based drugs with similar structures.

Supplementary Material

cb5c00656_si_001.pdf (8.1MB, pdf)

Acknowledgments

We thank John Eng, Venu Vendavasi, and Christina J. DeCoste at the Princeton Mass Spectrometry, Biophysics and Flow Cytometry core facilities, respectively, for providing support for nucleoside LC-MS, plate reader-based assays, and flow cytometry. R.E.K. acknowledges support from the Ludwig Princeton Branch, NIH (R01 GM132189) and NSF (MCB-1942565). J.D.R. and J.A.B. acknowledge support from the Ludwig Princeton Branch. X.S. and T.E.P. were supported by a generous gift from the Edward C. Taylor Third Year Graduate Fellowship in Chemistry.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acschembio.5c00656.

  • Experimental materials and methods, nucleoside and nucleotide LC-MS data, KO cell line validation, cell growth assays, DNA damage assays, protein translation assays, nucleolar microscopy (PDF)

The authors declare no competing financial interest.

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