A single m6A modification in U6 snRNA diversifies exon sequence at the 5’ splice site

Ishigami, Yuma; Ohira, Takayuki; Isokawa, Yui; Suzuki, Yutaka; Suzuki, Tsutomu

doi:10.1038/s41467-021-23457-6

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Published: 28 May 2021

A single m⁶A modification in U6 snRNA diversifies exon sequence at the 5’ splice site

Nature Communications volume 12, Article number: 3244 (2021) Cite this article

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Abstract

N⁶-methyladenosine (m⁶A) is a modification that plays pivotal roles in RNA metabolism and function, although its functions in spliceosomal U6 snRNA remain unknown. To elucidate its role, we conduct a large-scale transcriptome analysis of a Schizosaccharomyces pombe strain lacking this modification and found a global change of pre-mRNA splicing. The most significantly impacted introns are enriched for adenosine at the fourth position pairing the m⁶A in U6 snRNA, and exon sequences weakly recognized by U5 snRNA. This suggests cooperative recognition of 5’ splice site by U6 and U5 snRNPs, and also a role of m⁶A facilitating efficient recognition of the splice sites weakly interacting with U5 snRNA, indicating that U6 snRNA m⁶A relaxes the 5’ exon constraint and allows protein sequence diversity along with explosively increasing number of introns over the course of eukaryotic evolution.

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Introduction

Post-transcriptional modification is a characteristic structural feature of RNAs. About 150 types of chemical modifications have been found in various RNAs from all domains of life^1,2,3. Recent studies using deep-sequencing technologies successfully mapped several species of RNA modifications in eukaryotic mRNAs and non-coding RNAs in a transcriptome-wide manner^4,5,6. These findings raise the concept of the “epitranscriptome” and highlight the importance of RNA modification as a previously unrecognized layer of regulatory gene expression.

N⁶-methyladenosine (m⁶A), an abundant modification in eukaryotic mRNAs and long non-coding RNAs, plays a critical role in various biological processes, including meiosis^7,8, circadian rhythm⁹, sex determination^10,11, cell proliferation¹², differentiation¹³, reprogramming¹⁴, and stress responses¹⁵. The biogenesis and dynamics of m⁶A have been studied extensively. The METTL3/METTL14 writer complex introduces m⁶A in the RRACH motif (R = A or G; H = except G)^{16,17,18,19,20,21,22,23}. Internal m⁶As are abundant in the last exon of mRNAs, near the stop codon^20,21,24,25, and are decoded by several reader proteins, including YTH proteins, thereby contributing to the diverse fates of mRNAs^{26,27,28,29,30,31,32,33}. m⁶A also has the ability to destabilize double-stranded regions of structured RNA, thereby facilitating recognition by hnRNP C and hnRNP G^34,35. It has been proposed that m⁶A is a reversible modification that can be modified back to adenosine by erasers FTO and ALKBH5^36,37.

In addition to internal m⁶A, N⁶, 2′-O-dimethyladenosine (m⁶Am) is present at the transcription start site of mRNAs in vertebrates^38,39. Recently, our group reported a cap-specific m⁶A writer, CAPAM/PCIF1, which catalyzes N⁶-methylation of m⁶Am⁴⁰. Structural studies of CAPAM revealed the molecular basis of cap-specific m⁶Am formation, and ribosome profiling analysis revealed that CAPAM-mediated N⁶-methylation promotes translation of mRNAs starting from m⁶Am.

METTL16 is an m⁶A writer for U6 snRNA and MAT2A mRNA^41,42,43, which encodes S-adenosylmethionine (SAM) synthetase. METTL16 targets a specific nonamer sequence in structured RNA (UACAGAGAA) for m⁶A formation^41,44,45. At high SAM concentrations, METTL16 introduces m⁶A into six hairpin structures in the 3′ UTR of MAT2A mRNA, leading to degradation of the mRNA mediated by YTHDC1. At low SAM concentrations, METTL16 is bound to the hairpin structures and acts as a splicing enhancer for the 3′ terminal intron, thereby increasing production of mature MAT2A mRNA. Thus, METTL16 mediates a feedback mechanism that controls SAM homeostasis. This regulatory mechanism is critical for cell viability, as shown by the essentiality of METTL16 gene in human culture cells⁴⁶ and Mettl16 in mouse embryo⁴⁴. Mettl16 null mutations are embryonically lethal, and homozygotes die around the implantation stage (E3.5). Early embryos lacking Mettl16 contain reduced amounts of Mat2a mRNA, highlighting the importance of Mettl16 for production of this message.

U snRNAs are essential components of the spliceosome, contributing to recognition of substrate pre-mRNAs and serving as ribozyme catalysts of two consecutive transesterifications to ligate two exons concomitant with removal of an intron^{47,48,49,50,51,52,53,54}. U snRNAs contain a number of modified internal nucleosides, including pseudouridines, ribose 2′-O-methylations, and several base methylations^55,56,57,58. 2′-O-methylation and pseudouridylation of U2 snRNA are essential for snRNP biogenesis and efficient splicing^{59,60,61,62,63,64,65}. In U6 snRNA from mammals^56,66, S. pombe⁵⁷, and broad bean⁶⁷, an m⁶A is present in the middle position of the ACAGA box that recognizes the 5′ splice site (5′SS)^56,57,66,67 (Fig. 1a and Supplementary Fig. 1). In human, the m⁶A at position 43 in the ACAGA box is introduced by METTL16^41,42,68, whereas in Schizosaccharomyces pombe, the ortholog Mtl16 is responsible for the m⁶A modification at position 37 in the ACAGA box⁴¹. This m⁶A faces the fourth position of the intron when the ACAGA box base-pairs with the 5′SS in the spliceosome B complexes, implying a functional role of m⁶A in mRNA splicing⁴². However, because little change in global splicing is observed in null mutant mice embryos⁴⁴, and because alteration in splicing has not been studied in METTL16-deficient strains from other organisms, the functional significance of m⁶A in U6 snRNA remains elusive.

**Fig. 1: Phenotypes of cells lacking the m⁶A modification in U6 snRNA.**

To elucidate the functional role of m⁶A in U6 snRNA, we performed RNA-seq analyses of an S. pombe mtl16 knockout strain. We found that a subset of introns was retained in mRNAs, indicating a splicing defect caused by loss of m⁶A in U6 snRNA. The most significantly impacted introns were enriched for adenosine at the fourth position from the 5′SS, which base-pairs with m⁶A of U6 snRNA. In addition, the retained introns tended to have sequences that are weakly recognized by U5 snRNA at the 3′ termini of their 5′ exons, suggesting that U6 and U5 snRNPs cooperatively recognize the 5′SS, and that m⁶A in U6 snRNA facilitates efficient splicing of a subset of pre-mRNAs.

Results

Physiological phenotypes of S. pombe mtl16 knockout strains

To elucidate the functional roles of m⁶A in U6 snRNA (Fig. 1a and Supplementary Fig. 1), we chose S. pombe as a model organism because m⁶A was undetectable in the poly(A)⁺ RNA fraction (Supplementary Fig. 2) and the S. pombe MAT2a homolog sam1 contains no hairpin structures harboring the specific nonamer sequence targeted by METTL16⁶⁹. Consequently, U6 snRNA m⁶A is the sole target for S. pombe Mtl16. First, we verified that U6 snRNA m⁶A is introduced by S. pombe Mtl16. To this end, we isolated U6 snRNAs from wild-type (WT) and mtl16 knockout (mtl16Δ) strains by reciprocal circulating chromatography (RCC)⁷⁰. The isolated U6 snRNA was digested by RNase T₁ and subjected to capillary LC-nano-ESI-MS to analyze the RNA fragment containing m⁶A37. m⁶A was clearly present in U6 snRNA isolated from the WT strain (Fig. 1b and Supplementary Fig. 3a), and no fragment bearing A37 was detected, indicating that U6 snRNA was fully modified with m⁶A37. By contrast, m⁶A37 was completely absent in the U6 snRNA isolated from the mtl16Δ strain (Fig. 1b and Supplementary Fig. 3b). Next, we conducted an in vitro reconstitution of m⁶A in U6 snRNA using recombinantly expressed S. pombe Mtl16 in the presence or absence of SAM. m⁶A was efficiently introduced at position 37 of U6 snRNA in the presence of SAM (Fig. 1c), whereas m⁶A was slightly detected even in the absence of SAM (Fig. 1c), indicating that Mtl16 contains endogenous SAM in its active site. In addition, we probed the modified RNA fragment by collision-induced dissociation (CID) analysis to confirm the presence of m⁶A at position 37 (Supplementary Figs. 3c, d). These results demonstrated that S. pombe Mtl16 is an m⁶A writer for U6 snRNA. In a previous study⁴³, we reconstituted m⁶A formation in 3′ UTR hairpins of MAT2A mRNA using human METTL16. Similarly, in this study, we confirmed m⁶A formation in U6 snRNA by human METTL16 (Supplementary Figs. 4a, b).

We searched for physiological phenotypes of the mtl16Δ strain on media containing some inducers for cellular stress. As reported previously⁷¹, the mtl16Δ strain was sensitive to reagents that induce DNA damage, e.g., hydroxyurea (HU) and thiabendazole (TBZ) (Fig. 1d). In addition, we found that the mtl16Δ strain was sensitive to salt stress (KCl and CaCl₂); these sensitivities were exacerbated when the cells were cultured at 37 °C (Fig. 1d). The mtl16Δ strain grew more slowly than the WT strain in medium containing a nonfermentable carbon source (Fig. 1e), implying slight reduction of mitochondrial activity.

To determine whether m⁶A loss affects stability of U6 snRNA, we analyzed steady-state levels of U snRNAs by Northern blotting (Fig. 1f). We detected no significant difference in the levels of U6 and other U snRNAs between the WT and mtl16Δ strains, implying that the phenotype of the mtl16Δ strain is due to a functional consequence of the m⁶A loss from U6 snRNA.

Global alteration of mRNA splicing caused by m⁶A loss in U6 snRNA

Given that m⁶A37 in U6 snRNA is present in the ACAGA box, which pairs with the 5′SS of the intron in the spliceosome, we next asked whether mRNA splicing would be affected globally by loss of m⁶A in U6 snRNA. To this end, we performed RNA-seq of poly(A)⁺ RNAs from WT and mtl16Δ strains and compared the splicing properties of the two samples. Based on S. pombe genome annotations, we first took reads that mapped to each intron and its neighboring exons and classified them into five groups (Fig. 2a): 5′ exon–intron junction reads (EIJR), intron-3′ exon junction reads (IEJR), exon–exon canonical splicing junction reads (CSR), 5′ alternative splice site selection reads (A5R), and 3′ alternative splice site selection reads (A3R). We then developed algorithms to detect alteration in splicing efficiency and transcript abundance. For each of five groups (Fig. 2a), the number of reads (coverage) is returned by function cov(x). Intron Retention Score (IRS) and Proportion of Canonical Splicing (PCS) are calculated by Eqs. (1) to (3) against each intron in each sample (Supplementary Data 1). IRS is used for measuring the difference in intron retention level and PCS is used for measuring the fraction of canonically spliced transcripts.

$${\rm{cov}}\left({\rm{Total}}\right)=\frac{{\rm{cov}}\left({\rm{EIJR}}\right)+{\rm{cov}}\left({\rm{IEJR}}\right)}{2}+{\rm{cov}}\left({\rm{A}}5{\rm{R}}\right)+{\rm{cov}}\left({\rm{A}}3{\rm{R}}\right)+{\rm{cov}}\left({\rm{CSR}}\right)$$

(1)

$${\rm{IRS}}={{{\log }}}_{2}\left(\frac{{\rm{cov}}\left({\rm{EIJR}}\right)+{\rm{cov}}\left({\rm{IEJR}}\right)+0.1}{2\times {\rm{cov}}\left({\rm{CSR}}\right)+0.1}\right)$$

(2)

$${\rm{PCS}}=\frac{{\rm{cov}}\left({\rm{CSR}}\right)}{{\rm{cov}}\left({\rm{Total}}\right)}$$

(3)

A plot of IRS for mtl16Δ versus WT revealed a notable bias in the distribution; specifically, a number of introns in the mtl16Δ strain had higher IRS values (Fig. 2b). Therefore, in the mtl16Δ strain, we suggest that the pre-mRNA is dissociated from the spliceosome during the steps from the association of tri-snRNP until the first transesterification, which is at the state of B, Bact or B* complexes, leaving the intron retained. For introns with large differences in IRS, we confirmed intron retention in the mtl16Δ strain by semi-quantitative RT-PCR (Fig. 2c, Supplementary Fig. 5). Complementation of the mtl16Δ strain by ectopic expression of WT or the active-site mutant (P169A/P170A) of mtl16^41,44 confirmed that the alterations in splicing were dependent on the methyltransferase activity of Mtl16 (Fig. 2c).

**Fig. 2: Global alteration of mRNA splicing in *mtl16*Δ cells.**

To analyze the effect of mtl16 knockout on gene expression, we estimated the global alteration of transcriptome (Supplementary Fig. 6a and Supplementary Data 2). Since pre-mRNA splicing is significantly affected by the mtl16 knockout, we corrected the read counts of each transcript by multiplying PCS of introns inside the CDS, and applied them for differential expression analysis. As a result, 54 and 32 genes were up- and down-regulated, respectively, significantly over 2-fold (Supplementary Fig. 6a and Supplementary Data 2). In the up-regulated gene list, a set of genes upregulated under some stress conditions, such as heat, osmotic, oxidation, and nutrient starvation, were enriched according to the gene ontology enrichment analysis, implying that the mtl16Δ strain is suffering from several cellular stresses (Supplementary Fig. 6b). We speculate that the phenotypic features of this strain might be caused by accumulation of aberrant transcripts and proteins upon mtl16 knockout. Within the down-regulated genes in the mtl16Δ strain, mrp17 (mitochondrial ribosomal protein) and icp55 (intermediate cleaving peptidase 55) are required for mitochondrial function^72,73,74. The slow-growth phenotype of the mtl16Δ strain under nonfermentable condition (Fig. 1e) might be explained by downregulation of these proteins. In addition, downregulation of apl5 (AP-3 adaptor complex subunit)⁷⁵ and pot1 (shelterin complex subunit)⁷⁶ is associated with sensitivity against hydroxyurea (HU) and/or thiabendazole (TBZ) (Fig. 1d).

Introns bearing A at the 4^th nucleotide were sensitive to m⁶A loss from U6 snRNA

To characterize the 5′ and 3′SSs of the introns that exhibited a splicing defect, we chose a subset of introns showing large differences in IRS. To this end, we gave each intron a Z-score of a two-sample Z-test with a null hypothesis of an equal IRS, as IRS values follow normal distribution (Supplementary Fig. 7). Then, their Z-scores were sorted in descending order and split by quartiles (Fig. 2b). Sequence logos around the 5′ and 3′SSs of introns with Z-scores above the 1^st quartile and below the 3^rd quartile were generated by SeqLogo⁷⁷ (Fig. 2d), then the nucleotides enriched and depleted in the selected introns were shown by DiffLogo⁷⁸ (Fig. 2e). Notably, the fourth nucleotide of the 5′SS was significantly more likely to be A and less likely to be U (genomic T) in the group of introns with the highest Z-scores (Fig. 2e) than in the set of introns with the lowest Z-scores, in which T and A are equally likely at this position (Fig. 2e). By contrast, we observed no large difference in the sequence logo around the 3′SS (Fig. 2e). Given that m⁶A37 in U6 snRNA faces the fourth position of the intron during the catalytic steps of splicing⁷⁹, m⁶A loss might affect 5′SS recognition of a subset of introns bearing A at the fourth position (A4 introns).

To compare the splicing errors of introns with different nucleotides at the fourth position, we classified all introns with a sufficient read depth into four groups based on their fourth nucleotide: A4 (n = 2539), T4 (n = 1846), C4 (n = 514), or G4 (n = 157). We then plotted the IRS values for each group of introns in mtl16Δ versus WT strains (Fig. 3a and Supplementary Fig. 8). A large number of A4 introns exhibited a large difference in IRS implicating splicing error in the absence of m⁶A, whereas T4 introns were only slightly over the line of equal IRS, implicating little effect by m⁶A loss (Fig. 3a). Next, we calculated the difference in IRS for each group of introns upon mtl16 knockout and generated cumulative curves (Fig. 3b). As expected, the A4 and T4 introns had the largest and smallest IRS differences, respectively. G4 introns were less strongly affected than T4 introns, whereas the IRS differences of C4 introns were significantly higher than T4 and G4 introns, but lower than those of A4 introns. As observed for the A4 introns, the C4 introns are also spliced less efficiently than T4 and G4 introns in the absence of m⁶A in U6 snRNA. Thus, m⁶A of U6 snRNA plays a functional role in splicing of most introns improving their efficiencies, but to a greater extent against A4 and C4 introns.

**Fig. 3: The fourth nucleotide of the intron is the cis-element for sensitivity against m⁶A loss.**

In a previous study, it was shown that an m⁶A protruding from a double strand stabilizes the terminal base pair through base-stacking effect enforced by the hydrophobicity of the methyl group⁸⁰. To demonstrate this phenomenon in the spliceosome context, we synthesized an in vitro transcript that mimics the interaction between the ACAGA box of U6 snRNA and the 5′SS of A4 or T4 introns, and efficiently methylated it enzymatically by recombinant human METTL16 methyltransferase domain (Fig. 3c and Supplementary Fig. 9). Then, we measured the melting temperatures of the RNA hairpins with A4 or U4 in the presence or absence of m⁶A. As a result, consistent with previous studies⁸⁰, methylation of the A–A pair to m⁶A–A increased the thermostability of the double strand, and an opposite effect was given to the A–U pair when it was methylated to m⁶A–U (Fig. 3d). This result indicates that efficient recognition of A4 introns by m⁶A is due to its thermostabilizing property in the sequence context of U6 snRNA and 5′SS.

AAG consensus of 5′ exon suppresses sensitivity against m⁶A loss

In all annotated introns (Fig. 2d), AAG triplet is present as a weak consensus sequence at the 3′ terminus of the 5′ exon (positions −3 to −1). This trinucleotide is complementary to loop I of U5 snRNA (Fig. 2e) and facilitates interaction between pre-mRNA and U5 snRNP in the B^act complex^{81,82,83,84,85,86}. In the Two Sample Logo (Fig. 2e), the AAG triplet was depleted in A4 introns with splicing deficiencies in the mtl16Δ strain, but enriched in the rest of the introns. Accordingly, we classified the A4 introns (n = 2539) into two groups based on 5′ exonic sequence, AAG-A4 introns (n = 289) and non-AAG-A4 introns (n = 2250); we then plotted their IRS values in mtl16Δ versus WT strains (Fig. 4a), which shows a notable bias only in the non-AAG-A4 introns. We further classified non-AAG-A4 introns into seven groups (BAG, ABG, AAH, BBG, BAH, ABH, and BBH-A4) (Supplementary Fig. 10a) and calculated the IRS difference between the mtl16∆ and WT strains for each group (Supplementary Fig. 10b). The AAG-A4 introns have the smallest ratio, whereas the BBH-A4 introns have the largest (Supplementary Fig. 10c). Thus, the farther the 5′ exon sequence is from the AAG consensus, the larger the IRS difference the intron group will bear. These results suggest that the AAG consensus of the 5′ exon alleviates the splicing deficiency of the A4 introns caused by m⁶A loss of U6 snRNA via a stable interaction with the loop I of U5 snRNA.

**Fig. 4: Splicing defect of A4 introns was suppressed by the AAG sequence of the 5′ exon.**

To confirm this observation, we carried out a mutation study of an A4 intron using a minigene construct. We chose intron 1 of the SPAC18B11.09c gene, whose IRS values obtained by RNA-seq are −0.733 and 3.076 in the WT and mtl16Δ strains, respectively (Supplementary Data 1). The IRS values of the minigene construct obtained from semi-quantitative RT-PCR were 0.058 and 3.375 in the WT and mtl16Δ strains, respectively (Fig. 4b and Supplementary Fig. 11). When the GU at the 5′SS was mutated to GA, no splicing took place, as expected. When we mutated A at the fourth position to U, the IRS value in the WT strain increased to the same level as that in the mtl16Δ strain, indicating that the fourth nucleotide is the key cis-element for erroneous splicing in the absence of m⁶A in U6 snRNA. In addition, when the exonic ACC triplet was mutated to AAC or ACG, the splicing efficiency in both strains increased (i.e., IRS decreased). Furthermore, when the ACC was mutated to the AAG consensus sequence, splicing of this construct reached maximum efficiency even in the absence of m⁶A in U6 snRNA (Fig. 4b and Supplementary Fig. 11). This reporter experiment nicely supported our finding that m⁶A in U6 snRNA facilitates efficient splicing of A4 introns, especially when U5 snRNA is weakly associated with the 3′ terminus of the 5′ exon.

As an orthogonal approach to examine whether U6 and U5 snRNPs cooperatively recognize the 5′SS, we constructed a mutant of U5 snRNA in which UUU sequence of the loop I was replaced with AGA sequence (U5_AGA) (Fig. 4c). Then, a plasmid harboring the mutant U5_AGA was introduced in the WT and mtl16Δ strains. We measured the splicing efficiencies of several A4 introns harboring non-consensus triplet sequences at the 3′ terminus of the 5′ exon (Fig. 4c). The A4 intron of nhm1 mRNA has UCU triplet at the 5′ exonic sequence, which is weakly associated with loop I of U5 snRNA. The IRS of this intron was kept low in the WT strain, but significantly increased in the mtl16Δ strain as expected (Fig. 4c). When U5_AGA was introduced, the IRS of this intron was reduced significantly, whereas WT U5 snRNA did not change its IRS (Fig. 4c), because the UCU triplet is complementary to the AGA sequence of U5_AGA. The A4 intron of ysh1 has a UCC triplet that is recognized by the AGA in loop I of U5_AGA with two basepairs and one mismatch. The increased IRS in the mtl16Δ strain was reduced significantly upon U5_AGA expression (Fig. 4c). The A4 introns of trk2 and sec71 have CUG and CUC triplets at the 5′ exonic sequence, respectively. The AGA sequence of U5_AGA weakly recognizes these triplets with single-wobble pairing at the middle of the sequence. Curiously, but as expected, the increased IRS of these A4 introns in the mtl16Δ strain was slightly but significantly reduced by U5_AGA expression (Fig. 4c). As a negative control, the A4 intron of atg1803 has a canonical AAG triplet that is stably recognized by the endogenous U5 snRNA, but not recognized by the U5_AGA mutant (Fig. 4c). The IRS increased by U5_AGA expression (Fig. 4c), because U5_AGA competes with the endogenous U5 snRNA. These results clearly show that stable recognition of the 5′ exon triplet by loop I of U5 snRNA suppresses intron retention caused by m⁶A loss in U6 snRNA.

Discussion

Biochemical^79,81,82, genetic^87,88,89,90, and structural studies^{83,84,85,86,91,92,93,94} reveal that the ACAGA box of U6 snRNA and loop I of U5 snRNA recognize 5′SS at B or B^act complexes, and maintain this interaction until the intron lariat–spliceosome dissociation (Fig. 5a and Supplementary Fig. 12). Recognition of 5′SS by U6/U5 snRNAs contributes to definition and proofreading of pre-mRNA splicing^{88,95,96,97,98}, but little is known about the functional interaction between U5 snRNA–exon pairing and U6 snRNA–intron pairing. Here, we showed that in the WT S. pombe strain, the AAG-A4 introns are robustly recognized by U5 snRNA and U6 snRNA harboring the m⁶A modification (Fig. 5b). Even when the AAG consensus sequence is not present in the 5′ exon, the BBH-A4 introns are stably recognized by U6 snRNA with m⁶A modification (Fig. 5b) because m⁶A stabilizes the interaction between U6 snRNA and the intron via m⁶A–A pairing. In the mtl16Δ strain, the AAG-A4 introns are still recognized by U5 and U6 snRNPs in B or B^act complex, even when the m⁶A is not present in U6 snRNA (Fig. 5b). In this case, interaction between the 5′ exon and U5 snRNA plays a predominant role in efficient splicing. However, if the 5′ exon sequence deviates from the AAG consensus, the BBH-A4 introns are not stably recognized by U6 snRNA without the m⁶A modification (Fig. 5b). Collectively, these observations indicate that m⁶A in U6 snRNA facilitates efficient splicing of A4 introns only when U5 snRNA weakly recognizes the exon side of the 5′SS (Fig. 5b), indicating that U6 and U5 snRNPs interact cooperatively with pre-mRNA and stabilize the spliceosome during the splicing reaction. The cryo-EM structures of spliceosomes indicate that the interaction between pre-mRNA and U5/U6 snRNAs juxtaposes the 5′SS to the catalytic active site formed by the U snRNA backbones in the B complexes. Then, the Prp2 (S. pombe Cdc28) helicase pulls the intron to bring the branch site to the active site to form the B* complex. During these dynamic conformational changes, the 5′SS needs to be anchored to the active site, which enables the branch site 2′-OH to attack the splice site. We speculate that the stability of this anchoring interaction between pre-mRNA and U5/U6 snRNA decides the overall efficiency of the first transesterification (Fig. 5c).

**Fig. 5: Mechanistic insight into the functional role of U6 snRNA m⁶A modification on pre-mRNA splicing.**

As shown by the cryo-EM structures of the B and B^act complexes (Fig. 5a and Supplementary Fig. 12a), the 3′ terminal nucleotides of the 5′ exon are recognized by the UUmU sequence of loop I in U5 snRNA, whereas the intron sequence of 5′SS is base-paired with the ACm⁶A sequence of the ACAGA box in U6 snRNA. The m⁶A of U6 snRNA faces the fourth position of the intron. In the ILS complex (Supplementary Fig. 12b), the U6 snRNA–intron pairing is slightly different from that in the B complexes. Especially, m⁶A37 shifts out of the pairing with U4 in the intron lariat. It is possibly due to the conformational alterations of the spliceosome after the first transesterification. Tight recognition of 5′SS by U6 snRNA might not be necessary after the lariat formation. From a chemical viewpoint, the N⁶-methyl group of m⁶A has two conformations, syn and anti. Energetically, the syn conformation is favored over the anti conformation by 1.5 kcal/mol⁸⁰. However, structural studies revealed that m⁶A–U pairing in the RNA duplex assumes the high-energy anti conformation. In support of this idea, thermodynamic measurement of RNA duplexes revealed that m⁶A destabilizes A–U pairs by 0.5–1.7 kcal/mol, but stabilizes A–A pairs by 0.7 kcal/mol. In addition, when m⁶A is present at the 5′ or 3′ overhang position, it stacks with the terminal base pair and stabilizes the RNA duplex by 0.9–1.2 kcal/mol, compared to unmodified A. Also, these thermodynamic properties were shown to be consistent with the results obtained from the sequence context of the spliceosome, by measuring the melting temperatures of modified and unmodified transcripts that mimic the base pairing between U6 snRNA and the 5′ end of an intron (Figs. 3c, d). These chemical properties of m⁶A neatly explain how m⁶A in U6 snRNA facilitates 5′SS recognition of A4 introns.

The evolutionary conservation of METTL16 implies that m⁶A in U6 snRNA is functionally important. METTL16 orthologs are highly conserved in metazoans, fungi, plants, and protists, implying that this modification is a fundamental component of the splicing machinery. However, m⁶A is not present in some organisms with small numbers of introns, including several species of budding yeasts such as Saccharomyces cerevisiae and Candida albicans and the red algae Cyanidioschyzon merolae (Supplementary Fig. 13). Presumably, METTL16 and m⁶A in U6 snRNA have been lost independently from these organisms over the course of evolution. In vertebrates, including humans, more than 200,000 introns are present in the set of all genes⁹⁹, and the consensus sequence of 5′SS is GUAAGU (A4 intron). Among fungi, S. pombe has over 5000 introns and S. cerevisiae has 296; the consensus sequences of the 5′SS are GUAAGU (A4 intron) and GUAUGU (T4 intron), respectively^69,100. Given that m⁶A in U6 snRNA facilitates efficient splicing of A4 introns, S. cerevisiae may preferentially use T4 introns rather than A4 introns to achieve efficient splicing in the absence of this modification. C. albicans also has a 5′SS consensus of T4¹⁰⁰. On the other hand, C. merolae has only 27 introns, and its 5′SS consensus sequence is GUAAGU (A4 intron)¹⁰¹. Curiously, in C. merolae U6 snRNA, the ACAGA box is ACUGA, and the central U base pairs with the A4 position of introns with Watson–Crick geometry¹⁰². This is an alternative means of stabilizing 5′SS recognition by U6 snRNA without having m⁶A in the ACAGA box. Based on these cases, m⁶A in U6 snRNA has likely contributed to expansion of 5′SS sequence variation in organisms with large numbers of introns as it facilitates efficient splicing of non-AAG-A4 introns.

METTL16 is a key regulator of SAM synthetase (MAT2A), which maintains cellular SAM homeostasis^41,43. METTL16 introduces m⁶A into the hairpin structures in the 3′ UTR of MAT2A mRNA by sensing cellular SAM concentrations, thereby controlling mRNA stability. In addition, METTL16 acts as a splicing enhancer to recognize the same hairpins and produce functional MAT2A protein. In fact, Mettl16 null mice die during early embryogenesis with a reduced steady-state level of Mat2a mRNA⁴⁴, highlighting the importance of Mettl16 function in regulation of Mat2a levels. The 3′ UTR hairpins of MAT2A mRNA are conserved in vertebrates, but not in other organisms¹⁰³. In fact, S. pombe sam1 mRNA, an ortholog of MAT2A, is not affected in the mtl16Δ strain (0.96-fold change; no significant alteration). Thus, the observed phenotypes of this strain can be explained as a function of the splicing defect caused by loss of m⁶A in U6 snRNA. Nematode and plant homologs of METTL16 also have physiological functions, and strains harboring knockouts of these genes exhibit severe phenotypes. mett-10, a Caenorhabditis elegans ortholog, is required for embryonic development and germ cell differentiation¹⁰⁴. FIONA1, an Arabidopsis thaliana homolog, plays an essential role in regulating the circadian clock¹⁰⁵. These phenotypes might be explained by splicing alterations caused by loss of m⁶A from U6 snRNA. Because little change in global splicing is observed in early embryos (E3.5) of Mettl16 null mice⁴⁴, the functional significance of m⁶A in U6 snRNA in mammals remains elusive. Presumably, however, maternal U6 snRNA bearing m⁶A is still present at this stage. Further studies will be necessary to determine whether m⁶A loss of U6 snRNA affects global splicing in vertebrate systems.

Given that the frequency of m⁶A in U6 snRNA is nearly 100% (Fig. 1b), this modification might be a structural component of spliceosome, relaxing the 5′ exon constraint and diversifying the protein sequence along with the number of introns increases over the course of eukaryotic evolution. However, because vertebrate METTL16 is a SAM-sensitive methyltransferase⁴⁵, and the m⁶A modifications in the 3′UTR of MAT2A mRNA are regulated by availability of cellular SAM⁴³, m⁶A in U6 snRNA might be regulated by intracellular SAM concentrations. U6 snRNA m⁶A might also be targeted by “m⁶A eraser” demethylases in higher eukaryotes. This idea raises the possibility that U6 snRNA m⁶A is not only a structural component of the spliceosome, but also a regulator of the splicing efficiency of a subset of mRNAs under SAM-depleted conditions.

Methods

Yeast strains and cultivation

The ED668 (h+ ade6-M216 ura4-D18 leu1–32) (WT) and SPAC27D7.08cΔ (mtl16Δ) strains were obtained from the Bioneer deletion collection⁷⁴. The deletion mutation was verified by PCR. Strains were grown at 30 °C in YE5S media [0.5% yeast extract and 3% glucose, supplemented with adenine, uracil, lysine, histidine, and leucine (225 mg/mL each)] or EMM media [EMM without dextrose (MP Biochemicals) and 2% glucose, supplemented with adenine and leucine (225 mg/mL each)]. For spot analyses, yeast cells were first grown to mid-log phase, serially diluted (1:5) from a starting sample of 10⁴ cells, and spotted onto YE5S agar plates supplemented with inducers of cellular stress: 15 μg/ml thiabendazole (TBZ), 7.5 mM hydroxyurea (HU), 1 M KCl, or 100 mM CaCl₂. The plates were photographed after 3–4 days of incubation at 30 °C or 37 °C. To measure non-fermentable growth, the yeast strains were cultured in medium containing 0.5% yeast extract, 0.1% glucose, and 3% glycerol supplemented with adenine, uracil, lysine, histidine, and leucine (225 mg/mL each). OD₆₀₀ was measured every hour for 12 h.

RNA preparation

Total RNA from yeast cells for snRNA isolation was prepared by hot phenol extraction¹⁰⁶. Cell pellets were resuspended in extraction buffer [50 mM NaOAc (pH 5.2), 10 mM EDTA (pH 8.0), and 10% SDS], mixed with an equal volume of neutral phenol, incubated at 95 °C for 15 min, and centrifuged at 8000 × g for 30 min at 4 °C. The supernatant was collected and mixed with 1/4^th volume of chloroform, followed by centrifugation at 8000 × g for 15 min at 4 °C. The supernatant was subjected to isopropanol precipitation to obtain an RNA pellet. The RNA pellet was dissolved in 10 mL buffer consisting of 20 mM HEPES-KOH (pH 7.5) and 80 mM NaCl, and then mixed with 4 mL of 2-butoxyethanol. The mixture was placed on ice for 30 min. The precipitate containing polysaccharide was removed by centrifugation at 8000 × g for 30 min at 4 °C. The supernatant was subjected to ethanol precipitation to obtain total RNA.

Nucleoside analysis of poly(A)⁺ RNA

Total RNA extracted from exponentially growing S. pombe WT cells underwent four rounds of enrichment of poly(A)⁺ RNA using Dynabeads oligo [dT]₂₅ (Invitrogen). Eluate RNAs from each round of enrichment (1 μg each) were digested into nucleosides with 0.07 units of nuclease P₁ (Fujifilm Wako Pure Chemical Corporation), 0.127 units of phosphodiesterase I (Worthington Biochemical Corporation), and 0.08 units of bacterial alkaline phosphatase (BAP from E. coli C75, Takara). The digests were subjected to LC/MS analysis using a Q Exactive Hybrid Quadrupole-Orbitrap Mass Spectrometer (Thermo Fisher Scientific) equipped with a Dionex UltiMate 3000 LC System (Thermo Fisher Scientific) and an InertSustain C18 column (5 μm, 2.1 × 250 mm, GL Science). The mass chromatogram peak area of adenosine (A, m/z 268.10), N⁶-methyladenosine (m⁶A, m/z 282.12), N⁶-dimethyladenosine (m^6,6A, m/z 296.14), and 2′-O-methyladenosine (Am, m/z 282.12) was measured. Then, the peak area of m⁶A, m^6,6A, and Am was divided by the peak area of A, and these ratios at each round of enrichment were normalized by the ratio from the first eluate and then plotted for each modification. Data were analyzed on Xcalibur 4.1 and Excel 2016, and visualized by R 4.0.3 and Canvas 15.

Isolation of U6 snRNA and mass spectrometry

Isolation of yeast U6 snRNA was performed by RCC⁷⁰ using a DNA probe (Supplementary Data 3). Mass spectrometric analysis of RNase T₁- or A-digested fragments of the isolated U6 snRNA was performed by capillary LC-nano-ESI-MS¹⁰⁷. Mongo Oligo Mass Calculator v2.08 (https://mods.rna.albany.edu/masspec/Mongo-Oligo) was used for assignment of the product ions in CID spectra¹⁰⁸.

Preparation of transcripts for in vitro reconstitution

DNA templates for in vitro transcription were prepared using a series of DNA oligos (described in Supplementary Data 3). U6 snRNAs were transcribed by T7 RNA polymerase in vitro essentially as described¹⁰⁹. Briefly, DNA template was mixed with in-house made T7 RNA polymerase, NTPs, and a transcription buffer consisting of 40 mM Tris-HCl (pH 7.5), 24 mM MgCl₂, 5 mM DTT, 2.5 mM Spermidine, and 0.01% Triton X-100, and was incubated at 37 °C overnight. The reaction was stopped by addition of an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1, v/v/v). The supernatant was desalted by PD-10. Desalted RNA was collected by ethanol precipitation. The transcript was resolved by 10% denaturing PAGE and purified by excision from the gel.

Expression and purification of recombinant proteins

To obtain recombinant S. pombe Mtl16, the mtl16 cDNA was amplified by RT-PCR of total RNA from WT S. pombe cells using the primers listed in Supplementary Data 3, which were designed based on genomic sequence retrieved from PomBase¹¹⁰. Because the original initiation codon (genomic position I:4,522,948-4,522,950) of mtl16 ORF in the database was misannotated inside an unannotated intron (genomic position I:4,522,922–4,522,964), we constructed a recombinant Mtl16 starting from an AUG codon (genomic position I:4,523,030–4,523,032) upstream of the unannotated intron. The PCR product was cloned into the NheI and XhoI sites of pET-28a(+) (Novagen) to yield pET-Spmtl16, which produces N-terminal hexahistidine-tagged Mtl16 (His₆-Mtl16). The E. coli Rosetta2 (DE3) strain was transformed with pET-Spmtl16 and cultured in LB medium containing 50 μg/mL kanamycin and 30 μg/mL chloramphenicol. When the bacteria reached OD₆₀₀ of 0.5, protein expression was induced by addition of 10 μM IPTG and the cells were grown for an additional 3 h at 37 °C. Cells were harvested and suspended in buffer A [50 mM HEPES-KOH (pH 7.5), 150 mM NaCl, and 7 mM β-mercaptoethanol] supplied with 0.5 mM PMSF, followed by sonication on ice. Cell lysates were cleared by ultracentrifugation at 100,000 × g for 60 min. The supernatant was loaded onto a nickel-charged HiTrap chelating column (GE Healthcare) and unbound proteins were washed out with buffer A. The recombinant proteins were eluted with a 50-mL linear gradient comprising 0–500 mM imidazole in buffer A. A fraction containing the recombinant protein was dialyzed overnight against dialysis buffer [50 mM HEPES-KOH (pH 7.5), 150 mM NaCl, 10% glycerol, and 7 mM β-mercaptoethanol]. The concentration of the purified protein was determined in a Bradford assay (Bio-Rad, Hercules, CA, USA) using bovine serum albumin as a standard. To obtain recombinant human METTL16, the METTL16 cDNA was amplified using the primers listed in Supplementary Data 3, which were designed based on human genomic sequence retrieved from Ensembl¹¹¹ and cloned into pET-28a(+) to yield pET-hMETTL16, which produces C-terminal hexahistidine-tagged METTL16 (METTL16-His₆). Expression and purification were carried out as described for S. pombe Mtl16. To obtain recombinant human METTL16 MTD, cDNA coding the methyltransferase domain of METTL16 was amplified using the primers listed in Supplementary Data 3 and cloned into pE-SUMO-TEV by SLiCE to yield pE-SUMO-hMETTL16MTD. Expression was carried out as described for S. pombe Mtl16. Following expression, cells were harvested and suspended in buffer A [50 mM HEPES-KOH (pH 7.5), 150 mM NaCl, and 7 mM β-mercaptoethanol] supplied with 0.5 mM PMSF, followed by sonication on ice. Cell lysates were cleared by ultracentrifugation at 100,000 × g for 60 min. The supernatant was loaded onto a nickel-charged HiTrap chelating column (GE Healthcare) and unbound proteins were washed out with buffer A. The recombinant proteins were eluted with a 50-mL linear gradient comprising 0–500 mM imidazole in buffer A. A fraction containing the recombinant protein was dialyzed overnight against dialysis buffer [50 mM HEPES-KOH (pH 7.5), 150 mM NaCl, 10% glycerol, and 7 mM β-mercaptoethanol] containing SUMO protease Ulp1. Dialyzed protein was applied to Ni Sepharose 6 Fast Flow (GE Healthcare) and the flow-through was collected. The cleaved protein was further purified on a HiScreen Capto SP ImpRes (GE Healthcare) column. The concentration of the purified protein was determined in a Pierce BCA Protein Assay (Thermo Fisher Scientific) using bovine serum albumin as a standard.

In vitro reconstitution using recombinant proteins

In vitro methylation of U6 snRNA was performed at 37 °C for 1 h in 50 µL of reaction mixture consisting of 10 mM HEPES-KOH (pH 7.5), 100 mM NaCl, 1 mM DTT, 2 μM U6 snRNA transcript, 1 μM His₆-Mtl16, and 1 mM SAM. The reaction was stopped by addition of an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1, v/v/v). RNA was collected by ethanol precipitation, digested with RNase T₁, and subjected to RNA-MS analysis as described above.

Northern blotting

Total RNA (1 μg) from yeast cells was resolved by 10% denaturing PAGE and either stained with SYBR Gold (Invitrogen) or blotted onto a nylon membrane (Amersham Hybond N+, GE Healthcare) in 1 × TBE using a Transblot Turbo (Bio-Rad). Fluorescence was visualized on an FLA-7000 imaging analyzer (Fujifilm). The membrane was air-dried and irradiated twice with UV light (254 nm, 120 mJ/cm²; CL-1000, UVP) to cross-link the blotted RNA. Hybridization was performed using PerfectHyb (TOYOBO) at 52 °C with 4 pmol of 5′-³²P-labeled oligonucleotides specific to each U snRNA (Supplementary Data 3). The membrane was washed four times with 2 × SSC [300 mM NaCl, 30 mM trisodium citrate (pH 7.0)] containing 0.1% SDS, dried, and exposed on an imaging plate (BAS-MS2040, Fujifilm). Radioactivity was visualized on the FLA-7000. Uncropped scans are available in the Source Data file.

RNA-seq sample preparation

Yeast were harvested, frozen in liquid nitrogen, and homogenized in TriPure Isolation Reagent (Roche Life Science) using SK-100 (Funakoshi). Total RNA was extracted with chloroform, recovered from the aqueous phase by isopropanol precipitation, ethanol precipitated, and dissolved in water. The total RNAs underwent two rounds of enrichment of poly(A)⁺ RNA using Dynabeads oligo [dT]₂₅ (Invitrogen) and were sequenced on an Illumina NovaSeq sequencer (150-bp, paired-end).

RNA-seq read alignment

Raw Illumina sequencing reads were trimmed using fastp software¹¹² to remove adaptor sequences and low-quality bases. The processed reads were first aligned to S. pombe rRNA and tRNA sequences, then the unmapped reads were aligned to S. pombe genome assembly ASM294v2.34 using STAR 2.7.3a¹¹³. The following parameters were used–twopassMode Basic–outSJfilterOverhangMin 30 8 8 8–alignIntronMax 2000–outFilterMultimapNmax 30–outFilterScoreMinOverLread 0.4–outFilterMatchNminOverLread 0.44–outFilterScoreMin 10. The statistics of trimming and mapping is listed in Supplementary Data 4. Read alignment files (.bam) were processed with custom Python scripts¹¹⁴ using the libraries Biopython¹¹⁵ and Pysam¹¹⁶, and the plots were written by R 4.0.3. For each segment comprising a pair of consecutive annotated exons (exon1 and exon2), reads that mapped to the annotated SS junction (CSR), exon 1–intron junction (EIJR), intron–exon 2 junction (IEJR), any alternative 5′SS to exon 2 (A5R), and exon 1 to any alternative 3′SS (A3R), were counted.

Calculation of erroneous splicing

For each annotated segment described above, we obtained IRS by Eq. (2) to assess retention level for each annotated intron. PCS was calculated by Eq. (3) to assess the proportion of canonically spliced mRNA within a mixture of mature, immature, and aberrant mRNA. To compare the difference in IRS between WT and mtl16Δ strains, as the IRS values mostly follow normal distribution (Supplementary Fig. 7), Z-scores with a null hypothesis of zero difference in IRS for each intron were obtained by Eq. (4), where $\mu$ and $\sigma$ represent the mean and standard deviation, respectively, of the IRS values among the quadruplicates of WT and mtl16Δ strains.

$$Z=\frac{{\mu }_{{mtl}16\Delta }-{\mu }_{{WT}}}{\sqrt{{{\sigma }_{{mtl}16\Delta }}^{2}/4+{{\sigma }_{{WT}}}^{2}/4}}$$

(4)

All annotated GY-AG introns with a sufficient arbitrary number of reads in all four replicates [cov(Total) > 10] and an average CSR over 10 in either strain were applied for this calculation so that the number of introns picked up is maximized and the variance in read counts among the replicates is minimized. This process picked up 5056 GY-AG introns with sufficient read counts out of 5364 annotated introns. The 5′ and 3′SS sequences of the introns with splicing deficiencies were characterized by choosing a subset of introns with large and small IRS differences. Upon sorting the introns by Z-scores in a descending order, those over the 1^st quartile were treated as introns with large IRS differences, and those under the 3^rd quartile were treated as introns with small IRS differences. Then, sequence logos around the 5′ and 3′ SSs of introns with large or small IRS differences with sufficient read depths were generated by SeqLogo⁷⁷. Then, two logos were compared by DiffLogo⁷⁸. The raw count and result of the calculation is provided in Supplementary Data 1.

Differential expression analysis

Raw counts for each annotated transcript were determined using the htseq-count command of HTSeq version 0.6.1¹¹⁷. The splicing-adjusted read counts of each annotated gene in each sample were calculated by Eq. (5), where $i$ is the intron number and ${{\rm{PCS}}}_{k}$ is the ${\rm{PCS}}$ value of intron number $k$. The product of each ratio of canonically spliced reads for each intron inside the CDS of the gene was multiplied against the overall read count that falls inside each genomic locus of the transcript.

$${\rm{Adjusted}}\; {\rm{counts}}={\rm{Raw}}\; {\rm{counts}}\times {\prod }_{k=1}^{i}\left({{\rm{PCS}}}_{k}\right)$$

(5)

Differential expression of adjusted counts was evaluated by edgeR version 3.12^118,119, applying a statistical cutoff of FDR < 0.05 and fold-change > 2 to obtain differentially expressed genes (Supplementary Data 2). Enriched features of gene lists were searched by the AnGeLi¹²⁰ tool with protein-coding genes as a background (Supplementary Data 5).

Rescue experiment and intron-specific semi-quantitative RT-PCR

The S. pombe mtl16 gene, including untranslated regions, was PCR-amplified from the genome and cloned into pREP42-mcs+ (Addgene plasmid 52691) digested by NdeI and BamHI using the primers listed in Supplementary Data 3 and the SLiCE method¹²¹ to yield the WT plasmid p-mtl16_WT. To construct an active-site mutant, both P169 and P170 were mutated to Ala residues by site-directed mutagenesis using the primers listed in Supplementary Data 3 to yield p-mtl16_mut. S. pombe mtl16Δ strain was transformed with the WT or mutant plasmid by the lithium acetate method¹²², cultivated in EMM medium devoid of thiamine until the cell density reached OD₆₀₀ = 1.0, and harvested. Total RNA was prepared using the TriPure Isolation Reagent (Roche Life Science). One microgram of total RNA from WT, KO, and transformed strains were treated with RQ1 RNase-free DNase (Promega) and subjected to reverse transcription with random hexamer and oligo dT primers using the Transcriptor First Strand cDNA Synthesis Kit (Roche), followed by PCR amplification with intron-specific primers (Supplementary Data 3) under the following conditions: 30 cycles at 95 °C for 30 s, 53 °C for 45 s, and 68 °C for 30 s. PCR products were resolved on 8% non-denaturing PAGE, stained with EtBr, and visualized on FLA-7000.

Minigene analysis

The flanking sequence of the first intron of SPAC18B11.09c was amplified using primers bearing NdeI and BamHI sites (Supplementary Data 3). The PCR product was cloned into the corresponding sites of pREP42-mcs+ to yield the WT minigene plasmid pREP-AC18B11.09c. Mutations were introduced by site-directed mutagenesis using the primers listed in Supplementary Data 3. S. pombe WT and mtl16Δ strains were transformed with the WT or mutated plasmid using the lithium acetate method, cultivated in EMM medium devoid of thiamine, until the cell density reached OD₆₀₀ = 1.0, and harvested. Total RNA was prepared using the TriPure Isolation Reagent. One microgram of total RNA was treated with RQ1 RNase-free DNase and then subjected to reverse transcription with nmt1 promoter-specific RT primer (Supplementary Data 3) using the Transcriptor First Strand cDNA Synthesis Kit, followed by PCR amplification with minigene-specific primers (Supplementary Data 3) under the following conditions: 16–23 cycles at 95 °C for 30 s, 53 °C for 45 s, and 68 °C for 30 s. The number of cycles was adjusted so that the signals of the PCR products were not saturated. The PCR products were resolved on 8% non-denaturing PAGE, stained with EtBr, and visualized on FLA-7000. Signals corresponding to spliced or retained introns were quantified using the Multi Gauge V3.0 software (Fujifilm). Signal intensities were converted to levels of PCR product using the signals of Loading Quick 100-bp DNA Ladder (Toyobo) as a standard. IRS is the binary logarithm of the value obtained by dividing the abundance of the retained isoform by the abundance of the spliced isoform.

Intron retention-specific RT-qPCR

The LEU2+ marker of pTN69 was replaced by ura4+ using the primers listed in Supplementary Data 3 and the SLiCE method¹²¹ to yield pDblet. The S. pombe snu5 gene and its flanking region was PCR-amplified from the genome and cloned into pDblet digested by HindIII using the primers listed in Supplementary Data 3 and the SLiCE method¹²¹. Then, site-directed mutagenesis using the primers listed in Supplementary Data 3 was used to introduce an additional 15-nt stem-loop in the 3′ end of U5 snRNA to yield the plasmid pU5_WT. Site-directed mutagenesis using the primers listed in Supplementary Data 3 was utilized to introduce a TTT to AGA mutation in the snu5 gene of pU5_WT to yield pU5_AGA. Total RNA (2 μg) was extracted from WT or the mtl16Δ strains transformed with pDblet, pU5_WT or pU5_AGA grown to mid-log phase in EMM media without uracil. cDNA was synthesized by Superscript III (Thermo Fisher Scientific) using random N₆ primers, according to the manufacturer’s instructions. PCR was performed in a 10 μL mixture containing a 0.2-μL aliquot of the cDNA solution, 0.2 μM of each PCR primers, and 1 × KAPA SYBR FAST Master Mix optimized for LightCycler480 (Kapa biosystems). The thermal cycling conditions included 45 cycles of 95 °C for 10 s, 58 °C for 20 s, and 72 °C for 1 s. Amplification of DNA was monitored by a LightCycler480 (Roche), according to the manufacturer’s instructions. To write a standard curve of Cp (crossing point) value against the amount of the template, depending on each set of primers, a dilution series of the S. pombe WT genomic DNA was amplified by the primers. To measure the IRS value of a single intron, primers spanning intron and exon were used to obtain the Cp value of the retained isoform, and primers constructed inside an adjacent single exon was used to obtain the Cp value of the total of both retained and spliced isoforms. Then, the abundance of cDNA derived from each isoform was calculated based on the standard curve. IRS is the binary logarithm of the value obtained by dividing the abundance of the retained isoform by the abundance of the spliced isoform.

Melting temperature observation of the transcript

In vitro transcripts U6A4, U6U4, and U6guide were synthesized from templates U6modelA4_T7R, U6modelU4_T7R, and U6modelGuide_T7R, respectively (Supplementary Data 3) as described. U6A4 and U6U4 were annealed with U6guide by incubating at 95 °C for 5 min and ramped at −0.1 °C/sec to 10 °C in a mixture consisting of 100 mM NaCl, 20 mM HEPES-KOH (pH 7.5), 75 μM U6A4 or U6U4 RNA, and 75 μM U6guide RNA. The U6guide RNA was designed to form a secondary structure with U6A4 or U6U4 RNAs that emulates human U6 snRNA secondary structure recognized by METTL16⁶⁸. The annealed mixture was applied for methylation by incubating at 37 °C overnight in a mixture consisting of 200 mM NaCl, 50 mM HEPES-KOH (pH 7.5), 1 mM DTT, 2.5 mM SAM, 0.2 units/uL SUPERase-In RNase inhibitor (Thermo Fisher Scientific), 30 μM human recombinant METTL16 MTD, and 30 μM RNA. The reaction was stopped by mixing with an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1, v/v/v). The RNA was collected by ethanol precipitation, then U6A4 RNA or U6U4 RNA was resolved by 10% denaturing PAGE and excised from the gel piece. The methylation efficiency was verified by LC/MS as described. The methylated or unmodified transcripts were dissolved at a final concentration of 3 μM in degassed buffer consisting of 20 mM sodium phosphate (pH 7.0) and 1 M NaCl, followed by incubation at 95 °C for 5 min and ramped at −0.1 °C/sec to 10 °C for annealing. A UV–Vis spectrophotometer (V-630, JASCO) equipped with an 8 multiquartz microcell array (path length: 10 mm, JASCO) was used to monitor the hyperchromicity by monitoring absorbance at 260 nm. The temperature gradient was as follows: 25 °C for 30 sec, ramped at 5 °C/min to 40 °C and held for 5 min, and after that ramped at 0.5 °C/min to 105 °C. The measurement was performed in triplicate for each sample.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

The sequence data from this study have been submitted to the DDBJ Sequence Read Archive under accession number DRA009909. Structural data was retrieved from PDB ID: 5O9Z [https://doi.org/10.2210/pdb5o9z/pdb], 5GM6 [https://doi.org/10.2210/pdb5gm6/pdb], and 3JB9 [https://doi.org/10.2210/pdb3jb9/pdb]. Source data are provided with this paper.

Code availability

Python scripts written for RNA-seq data analysis are available from a public GitHub repository https://github.com/YumaIshigami/irscalc¹¹⁴.

References

Boccaletto, P. et al. MODOMICS: a database of RNA modification pathways. 2017 update. Nucleic Acids Res 46, D303–D307 (2018).
Article CAS PubMed Google Scholar
Frye, M., Harada, B. T., Behm, M. & He, C. RNA modifications modulate gene expression during development. Science 361, 1346–1349 (2018).
Article ADS CAS PubMed PubMed Central Google Scholar
Suzuki, T. The expanding world of tRNA modifications and disease relevance. Nat. Rev. Mol. Cell Biol. https://doi.org/10.1038/s41580-021-00342-0 (2020).
Harcourt, E. M., Kietrys, A. M. & Kool, E. T. Chemical and structural effects of base modifications in messenger RNA. Nature 541, 339–346 (2017).
Article ADS CAS PubMed PubMed Central Google Scholar
Roundtree, I. A., Evans, M. E., Pan, T. & He, C. Dynamic RNA modifications in gene expression regulation. Cell 169, 1187–1200 (2017).
Article CAS PubMed PubMed Central Google Scholar
Frye, M., Jaffrey, S. R., Pan, T., Rechavi, G. & Suzuki, T. RNA modifications: what have we learned and where are we headed?. Nat. Rev. Genet. 17, 365–372 (2016).
Article CAS PubMed Google Scholar
Schwartz, S. et al. High-resolution mapping reveals a conserved, widespread, dynamic mRNA methylation program in yeast meiosis. Cell 155, 1409–1421 (2013).
Article CAS PubMed PubMed Central Google Scholar
Clancy, M. J., Shambaugh, M. E., Timpte, C. S. & Bokar, J. A. Induction of sporulation in Saccharomyces cerevisiae leads to the formation of N6-methyladenosine in mRNA: a potential mechanism for the activity of the IME4 gene. Nucleic Acids Res. 30, 4509–4518 (2002).
Article CAS PubMed PubMed Central Google Scholar
Fustin, J. M. et al. RNA-methylation-dependent RNA processing controls the speed of the circadian clock. Cell 155, 793–806 (2013).
Article CAS PubMed Google Scholar
Lence, T. et al. m(6)A modulates neuronal functions and sex determination in Drosophila. Nature 540, 242–247 (2016).
Article ADS CAS PubMed Google Scholar
Haussmann, I. U. et al. m(6)A potentiates Sxl alternative pre-mRNA splicing for robust Drosophila sex determination. Nature 540, 301–304 (2016).
Article ADS CAS PubMed Google Scholar
Wang, Y. et al. N6-methyladenosine modification destabilizes developmental regulators in embryonic stem cells. Nat. Cell Biol. 16, 191–198 (2014).
Article CAS PubMed PubMed Central Google Scholar
Batista, P. J. et al. m(6)A RNA modification controls cell fate transition in mammalian embryonic stem cells. Cell Stem Cell 15, 707–719 (2014).
Article CAS PubMed PubMed Central Google Scholar
Geula, S. et al. Stem cells. m6A mRNA methylation facilitates resolution of naive pluripotency toward differentiation. Science 347, 1002–1006 (2015).
Article ADS CAS PubMed Google Scholar
Zhou, J. et al. Dynamic m(6)A mRNA methylation directs translational control of heat shock response. Nature 526, 591–594 (2015).
Article ADS CAS PubMed PubMed Central Google Scholar
Schibler, U., Kelley, D. E. & Perry, R. P. Comparison of methylated sequences in messenger RNA and heterogeneous nuclear RNA from mouse L cells. J. Mol. Biol. 115, 695–714 (1977).
Article CAS PubMed Google Scholar
Csepany, T., Lin, A., Baldick, C. J. Jr. & Beemon, K. Sequence specificity of mRNA N6-adenosine methyltransferase. J. Biol. Chem. 265, 20117–20122 (1990).
Article CAS PubMed Google Scholar
Harper, J. E., Miceli, S. M., Roberts, R. J. & Manley, J. L. Sequence specificity of the human mRNA N6-adenosine methylase in vitro. Nucleic Acids Res. 18, 5735–5741 (1990).
Article CAS PubMed PubMed Central Google Scholar
Rottman, F. M., Bokar, J. A., Narayan, P., Shambaugh, M. E. & Ludwiczak, R. N6-adenosine methylation in mRNA: substrate specificity and enzyme complexity. Biochimie 76, 1109–1114 (1994).
Article CAS PubMed Google Scholar
Dominissini, D. et al. Topology of the human and mouse m6A RNA methylomes revealed by m6A-seq. Nature 485, 201–206 (2012).
Article ADS CAS PubMed Google Scholar
Meyer, K. D. et al. Comprehensive analysis of mRNA methylation reveals enrichment in 3’ UTRs and near stop codons. Cell 149, 1635–1646 (2012).
Article CAS PubMed PubMed Central Google Scholar
Linder, B. et al. Single-nucleotide-resolution mapping of m6A and m6Am throughout the transcriptome. Nat. Methods 12, 767–772 (2015).
Article CAS PubMed PubMed Central Google Scholar
Liu, J. et al. A METTL3-METTL14 complex mediates mammalian nuclear RNA N6-adenosine methylation. Nat. Chem. Biol. 10, 93–95 (2014).
Article CAS PubMed Google Scholar
Ke, S. et al. A majority of m6A residues are in the last exons, allowing the potential for 3’ UTR regulation. Genes Dev. 29, 2037–2053 (2015).
Article CAS PubMed PubMed Central Google Scholar
Huang, H. et al. Histone H3 trimethylation at lysine 36 guides m(6)A RNA modification co-transcriptionally. Nature 567, 414–419 (2019).
Article ADS CAS PubMed PubMed Central Google Scholar
Wang, X. et al. N6-methyladenosine-dependent regulation of messenger RNA stability. Nature 505, 117–120 (2014).
Article ADS PubMed CAS Google Scholar
Alarcon, C. R. et al. HNRNPA2B1 is a mediator of m(6)A-dependent nuclear RNA processing events. Cell 162, 1299–1308 (2015).
Article CAS PubMed PubMed Central Google Scholar
Wang, X. et al. N(6)-methyladenosine modulates messenger RNA translation efficiency. Cell 161, 1388–1399 (2015).
Article CAS PubMed PubMed Central Google Scholar
Xiao, W. et al. Nuclear m(6)A reader YTHDC1 regulates mRNA splicing. Mol. Cell 61, 507–519 (2016).
Article CAS PubMed Google Scholar
Patil, D. P. et al. m(6)A RNA methylation promotes XIST-mediated transcriptional repression. Nature 537, 369–373 (2016).
Article ADS CAS PubMed PubMed Central Google Scholar
Roundtree, I. A. & et al. YTHDC1 mediates nuclear export of N(6)-methyladenosine methylated mRNAs. eLife 6, e31311 (2017).
Article PubMed PubMed Central Google Scholar
Shi, H. et al. YTHDF3 facilitates translation and decay of N(6)-methyladenosine-modified RNA. Cell Res. 27, 315–328 (2017).
Article CAS PubMed PubMed Central Google Scholar
Li, A. et al. Cytoplasmic m(6)A reader YTHDF3 promotes mRNA translation. Cell Res. 27, 444–447 (2017).
Article CAS PubMed PubMed Central Google Scholar
Liu, N. et al. N(6)-methyladenosine-dependent RNA structural switches regulate RNA-protein interactions. Nature 518, 560–564 (2015).
Article ADS CAS PubMed PubMed Central Google Scholar
Liu, N. et al. N6-methyladenosine alters RNA structure to regulate binding of a low-complexity protein. Nucleic Acids Res. 45, 6051–6063 (2017).
Article CAS PubMed PubMed Central Google Scholar
Jia, G. et al. N6-methyladenosine in nuclear RNA is a major substrate of the obesity-associated FTO. Nat. Chem. Biol. 7, 885–887 (2011).
Article CAS PubMed PubMed Central Google Scholar
Zheng, G. et al. ALKBH5 is a mammalian RNA demethylase that impacts RNA metabolism and mouse fertility. Mol. Cell 49, 18–29 (2013).
Article CAS PubMed Google Scholar
Keith, J. M., Ensinger, M. J. & Moss, B. HeLa cell RNA (2’-O-methyladenosine-N6-)-methyltransferase specific for the capped 5’-end of messenger RNA. J. Biol. Chem. 253, 5033–5039 (1978).
Article CAS PubMed Google Scholar
Banerjee, A. K. 5’-terminal cap structure in eucaryotic messenger ribonucleic acids. Microbiol. Rev. 44, 175–205 (1980).
Article CAS PubMed PubMed Central Google Scholar
Akichika, S. & et al. Cap-specific terminal N(6)-methylation of RNA by an RNA polymerase II-associated methyltransferase. Science 363, aav0080 (2019).
Article CAS Google Scholar
Pendleton, K. E. et al. The U6 snRNA m(6)A methyltransferase METTL16 regulates SAM synthetase intron retention. Cell 169, 824–835 e14 (2017).
Article CAS PubMed PubMed Central Google Scholar
Warda, A. S. et al. Human METTL16 is a N(6)-methyladenosine (m(6)A) methyltransferase that targets pre-mRNAs and various non-coding RNAs. EMBO Rep. 18, 2004–2014 (2017).
Article CAS PubMed PubMed Central Google Scholar
Shima, H. et al. S-Adenosylmethionine synthesis is regulated by selective N(6)-adenosine methylation and mRNA degradation involving METTL16 and YTHDC1. Cell Rep. 21, 3354–3363 (2017).
Article CAS PubMed Google Scholar
Mendel, M. et al. Methylation of structured RNA by the m(6)A writer METTL16 Is essential for mouse embryonic development. Mol. Cell 71, 986–1000 e11 (2018).
Article CAS PubMed PubMed Central Google Scholar
Doxtader, K. A. et al. Structural basis for regulation of METTL16, an S-adenosylmethionine homeostasis factor. Mol. Cell 71, 1001–1011 e4 (2018).
Article CAS PubMed PubMed Central Google Scholar
Blomen, V. A. et al. Gene essentiality and synthetic lethality in haploid human cells. Science 350, 1092–1096 (2015).
Article ADS CAS PubMed Google Scholar
Lerner, M. R., Boyle, J. A., Mount, S. M., Wolin, S. L. & Steitz, J. A. Are snRNPs involved in splicing? Nature 283, 220–224 (1980).
Article ADS CAS PubMed Google Scholar
Steitz, T. A. & Steitz, J. A. A general two-metal-ion mechanism for catalytic RNA. Proc. Natl Acad. Sci. USA 90, 6498–6502 (1993).
Article ADS CAS PubMed MATH PubMed Central Google Scholar
Yean, S. L., Wuenschell, G., Termini, J. & Lin, R. J. Metal-ion coordination by U6 small nuclear RNA contributes to catalysis in the spliceosome. Nature 408, 881–884 (2000).
Article ADS CAS PubMed PubMed Central Google Scholar
Fica, S. M. et al. RNA catalyses nuclear pre-mRNA splicing. Nature 503, 229–234 (2013).
Article ADS CAS PubMed PubMed Central Google Scholar
Staley, J. P. & Guthrie, C. Mechanical devices of the spliceosome: motors, clocks, springs, and things. Cell 92, 315–326 (1998).
Article CAS PubMed Google Scholar
Wahl, M. C., Will, C. L. & Luhrmann, R. The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 (2009).
Article CAS PubMed Google Scholar
Fica, S. M. & Nagai, K. Cryo-electron microscopy snapshots of the spliceosome: structural insights into a dynamic ribonucleoprotein machine. Nat. Struct. Mol. Biol. 24, 791–799 (2017).
Article CAS PubMed PubMed Central Google Scholar
Yan, C., Wan, R. & Shi, Y. Molecular Mechanisms of pre-mRNA Splicing through Structural Biology of the Spliceosome. Cold Spring Harb. Perspect. Biol. 11, a032409 (2019).
Article CAS PubMed PubMed Central Google Scholar
Karijolich, J. & Yu, Y. T. Spliceosomal snRNA modifications and their function. RNA Biol. 7, 192–204 (2010).
Article CAS PubMed Google Scholar
Epstein, P., Reddy, R., Henning, D. & Busch, H. The nucleotide sequence of nuclear U6 (4.7 S) RNA. J. Biol. Chem. 255, 8901–8906 (1980).
Article CAS PubMed Google Scholar
Gu, J., Patton, J. R., Shimba, S. & Reddy, R. Localization of modified nucleotides in Schizosaccharomyces pombe spliceosomal small nuclear RNAs: modified nucleotides are clustered in functionally important regions. RNA 2, 909–918 (1996).
CAS PubMed PubMed Central Google Scholar
Zhou, H., Qu, L., Du, Y. & Zhou, W. Identification of a novel methylated nucleoside inSchizosac-charomyces pombe U6 snRNA. Chin. Sci. Bull. 45, 1304–1308 (2000).
Article CAS Google Scholar
Yu, Y. T., Shu, M. D. & Steitz, J. A. Modifications of U2 snRNA are required for snRNP assembly and pre-mRNA splicing. EMBO J. 17, 5783–5795 (1998).
Article CAS PubMed PubMed Central Google Scholar
Newby, M. I. & Greenbaum, N. L. Sculpting of the spliceosomal branch site recognition motif by a conserved pseudouridine. Nat. Struct. Biol. 9, 958–965 (2002).
Article CAS PubMed Google Scholar
Ma, X., Zhao, X. & Yu, Y. T. Pseudouridylation (Psi) of U2 snRNA in S. cerevisiae is catalyzed by an RNA-independent mechanism. EMBO J. 22, 1889–1897 (2003).
Article CAS PubMed PubMed Central Google Scholar
Donmez, G., Hartmuth, K. & Luhrmann, R. Modified nucleotides at the 5’ end of human U2 snRNA are required for spliceosomal E-complex formation. RNA 10, 1925–1933 (2004).
Article PubMed PubMed Central CAS Google Scholar
Ma, X. et al. Pseudouridylation of yeast U2 snRNA is catalyzed by either an RNA-guided or RNA-independent mechanism. EMBO J. 24, 2403–2413 (2005).
Article CAS PubMed PubMed Central Google Scholar
Wu, G., Xiao, M., Yang, C. & Yu, Y. T. U2 snRNA is inducibly pseudouridylated at novel sites by Pus7p and snR81 RNP. EMBO J. 30, 79–89 (2011).
Article PubMed CAS Google Scholar
Wu, G. et al. Pseudouridines in U2 snRNA stimulate the ATPase activity of Prp5 during spliceosome assembly. EMBO J. 35, 654–667 (2016).
Article CAS PubMed PubMed Central Google Scholar
Shimba, S., Bokar, J. A., Rottman, F. & Reddy, R. Accurate and efficient N-6-adenosine methylation in spliceosomal U6 small nuclear RNA by HeLa cell extract in vitro. Nucleic Acids Res. 23, 2421–2426 (1995).
Article CAS PubMed PubMed Central Google Scholar
Kiss, T., Antal, M. & Solymosy, F. Plant small nuclear RNAs. II. U6 RNA and a 4.5SI-like RNA are present in plant nuclei. Nucleic Acids Res. 15, 543–560 (1987).
Article CAS PubMed PubMed Central Google Scholar
Aoyama, T., Yamashita, S. & Tomita, K. Mechanistic insights into m6A modification of U6 snRNA by human METTL16. Nucleic Acids Res. 48, 5157–5168 (2020).
Article CAS PubMed PubMed Central Google Scholar
Wood, V. et al. The genome sequence of Schizosaccharomyces pombe. Nature 415, 871–880 (2002).
Article ADS CAS PubMed Google Scholar
Miyauchi, K., Ohara, T. & Suzuki, T. Automated parallel isolation of multiple species of non-coding RNAs by the reciprocal circulating chromatography method. Nucleic Acids Res. 35, e24 (2007).
Article PubMed PubMed Central CAS Google Scholar
Pan, X. et al. Identification of novel genes involved in DNA damage response by screening a genome-wide Schizosaccharomyces pombe deletion library. BMC Genomics 13, 662 (2012).
Article CAS PubMed PubMed Central Google Scholar
Naamati, A., Regev-Rudzki, N., Galperin, S., Lill, R. & Pines, O. Dual targeting of Nfs1 and discovery of its novel processing enzyme, Icp55. J. Biol. Chem. 284, 30200–30208 (2009).
Article CAS PubMed PubMed Central Google Scholar
Vogtle, F. N. et al. Global analysis of the mitochondrial N-proteome identifies a processing peptidase critical for protein stability. Cell 139, 428–439 (2009).
Article PubMed CAS Google Scholar
Kim, D. U. et al. Analysis of a genome-wide set of gene deletions in the fission yeast Schizosaccharomyces pombe. Nat. Biotechnol. 28, 617–623 (2010).
Article CAS PubMed PubMed Central Google Scholar
Nguyen, T. T. et al. Predicting chemotherapeutic drug combinations through gene network profiling. Sci. Rep. 6, 18658 (2016).
Article ADS CAS PubMed PubMed Central Google Scholar
Tadeo, X. et al. Elimination of shelterin components bypasses RNAi for pericentric heterochromatin assembly. Genes Dev. 27, 2489–2499 (2013).
Article CAS PubMed PubMed Central Google Scholar
Bembom, O. & Ivanek, R. seqLogo: Sequence logos for DNA sequence alignments. R package version 1.56.0. (2020).
Nettling, M. et al. DiffLogo: a comparative visualization of sequence motifs. BMC Bioinforma. 16, 387 (2015).
Article CAS Google Scholar
Wassarman, D. A. & Steitz, J. A. Interactions of small nuclear RNA’s with precursor messenger RNA during in vitro splicing. Science 257, 1918–1925 (1992).
Article ADS CAS PubMed Google Scholar
Roost, C. et al. Structure and thermodynamics of N6-methyladenosine in RNA: a spring-loaded base modification. J. Am. Chem. Soc. 137, 2107–2115 (2015).
Article CAS PubMed PubMed Central Google Scholar
Sontheimer, E. J. & Steitz, J. A. The U5 and U6 small nuclear RNAs as active site components of the spliceosome. Science 262, 1989–1996 (1993).
Article ADS CAS PubMed Google Scholar
Newman, A. J., Teigelkamp, S. & Beggs, J. D. snRNA interactions at 5’ and 3’ splice sites monitored by photoactivated crosslinking in yeast spliceosomes. RNA 1, 968–980 (1995).
CAS PubMed PubMed Central Google Scholar
Yan, C., Wan, R., Bai, R., Huang, G. & Shi, Y. Structure of a yeast activated spliceosome at 3.5 A resolution. Science 353, 904–911 (2016).
Article ADS CAS PubMed Google Scholar
Rauhut, R. et al. Molecular architecture of the Saccharomyces cerevisiae activated spliceosome. Science 353, 1399–1405 (2016).
Article ADS CAS PubMed Google Scholar
Haselbach, D. et al. Structure and conformational dynamics of the human Spliceosomal B(act) complex. Cell 172, 454–464 e11 (2018).
Article CAS PubMed Google Scholar
Zhang, X. et al. Structure of the human activated spliceosome in three conformational states. Cell Res. 28, 307–322 (2018).
Article ADS CAS PubMed PubMed Central Google Scholar
Madhani, H. D., Bordonne, R. & Guthrie, C. Multiple roles for U6 snRNA in the splicing pathway. Genes Dev. 4, 2264–2277 (1990).
Article CAS PubMed Google Scholar
Newman, A. & Norman, C. Mutations in yeast U5 snRNA alter the specificity of 5’ splice-site cleavage. Cell 65, 115–123 (1991).
Article CAS PubMed Google Scholar
Newman, A. J. & Norman, C. U5 snRNA interacts with exon sequences at 5’ and 3’ splice sites. Cell 68, 743–754 (1992).
Article CAS PubMed Google Scholar
Lesser, C. F. & Guthrie, C. Mutations in U6 snRNA that alter splice site specificity: implications for the active site. Science 262, 1982–1988 (1993).
Article ADS CAS PubMed Google Scholar
Plaschka, C., Lin, P. C. & Nagai, K. Structure of a pre-catalytic spliceosome. Nature 546, 617–621 (2017).
Article ADS CAS PubMed PubMed Central Google Scholar
Bertram, K. et al. Cryo-EM structure of a pre-catalytic human spliceosome primed for activation. Cell 170, 701–713 e11 (2017).
Article CAS PubMed Google Scholar
Yan, C. et al. Structure of a yeast spliceosome at 3.6-angstrom resolution. Science 349, 1182–1191 (2015).
Article ADS CAS PubMed Google Scholar
Wan, R., Yan, C., Bai, R., Lei, J. & Shi, Y. Structure of an intron lariat spliceosome from Saccharomyces cerevisiae. Cell 171, 120–132 e12 (2017).
Article CAS PubMed Google Scholar
Koodathingal, P., Novak, T., Piccirilli, J. A. & Staley, J. P. The DEAH box ATPases Prp16 and Prp43 cooperate to proofread 5’ splice site cleavage during pre-mRNA splicing. Mol. Cell 39, 385–395 (2010).
Article CAS PubMed PubMed Central Google Scholar
Yang, F. et al. Splicing proofreading at 5’ splice sites by ATPase Prp28p. Nucleic Acids Res. 41, 4660–4670 (2013).
Article CAS PubMed PubMed Central Google Scholar
O’Keefe, R. T. Mutations in U5 snRNA loop 1 influence the splicing of different genes in vivo. Nucleic Acids Res. 30, 5476–5484 (2002).
Article PubMed PubMed Central Google Scholar
Kershaw, C. J., Barrass, J. D., Beggs, J. D. & O’Keefe, R. T. Mutations in the U5 snRNA result in altered splicing of subsets of pre-mRNAs and reduced stability of Prp8. RNA 15, 1292–1304 (2009).
Article CAS PubMed PubMed Central Google Scholar
Chen, W. & Moore, M. J. Spliceosomes. Curr. Biol. 25, R181–R183 (2015).
Article CAS PubMed Google Scholar
Neuveglise, C., Marck, C. & Gaillardin, C. The intronome of budding yeasts. C. R. Biol. 334, 662–670 (2011).
Article CAS PubMed Google Scholar
Matsuzaki, M. et al. Genome sequence of the ultrasmall unicellular red alga Cyanidioschyzon merolae 10D. Nature 428, 653–657 (2004).
Article ADS CAS PubMed Google Scholar
Stark, M. R. et al. Dramatically reduced spliceosome in Cyanidioschyzon merolae. Proc. Natl Acad. Sci. USA 112, E1191–E1200 (2015).
Article CAS PubMed PubMed Central Google Scholar
Parker, B. J. et al. New families of human regulatory RNA structures identified by comparative analysis of vertebrate genomes. Genome Res. 21, 1929–1943 (2011).
Article CAS PubMed PubMed Central Google Scholar
Dorsett, M., Westlund, B. & Schedl, T. METT-10, a putative methyltransferase, inhibits germ cell proliferative fate in Caenorhabditis elegans. Genetics 183, 233–247 (2009).
Article CAS PubMed PubMed Central Google Scholar
Kim, J., Kim, Y., Yeom, M., Kim, J. H. & Nam, H. G. FIONA1 is essential for regulating period length in the Arabidopsis circadian clock. Plant Cell 20, 307–319 (2008).
Article CAS PubMed PubMed Central Google Scholar
Collart, M. A. & Oliviero, S. Preparation of yeast RNA. Curr. Protoc. Mol. Biol. Chapter 13, Unit13 12 (2001).
PubMed Google Scholar
Suzuki, T. & Suzuki, T. A complete landscape of post-transcriptional modifications in mammalian mitochondrial tRNAs. Nucleic Acids Res. 42, 7346–7357 (2014).
Article CAS PubMed PubMed Central Google Scholar
Gao, Y., Yang, J., Cancilla, M. T., Meng, F. & McLuckey, S. A. Top-down interrogation of chemically modified oligonucleotides by negative electron transfer and collision induced dissociation. Anal. Chem. 85, 4713–4720 (2013).
Article CAS PubMed Google Scholar
Sampson, J. R. & Uhlenbeck, O. C. Biochemical and physical characterization of an unmodified yeast phenylalanine transfer RNA transcribed in vitro. Proc. Natl Acad. Sci. USA 85, 1033–1037 (1988).
Article ADS CAS PubMed PubMed Central Google Scholar
Lock, A. et al. PomBase 2018: user-driven reimplementation of the fission yeast database provides rapid and intuitive access to diverse, interconnected information. Nucleic Acids Res. 47, D821–D827 (2019).
Article CAS PubMed Google Scholar
Aken, B. L. et al. Ensembl 2017. Nucleic Acids Res. 45, D635–D642 (2017).
Article CAS PubMed Google Scholar
Chen, S., Zhou, Y., Chen, Y. & Gu, J. fastp: an ultra-fast all-in-one FASTQ preprocessor. Bioinformatics 34, i884–i890 (2018).
Article PubMed PubMed Central CAS Google Scholar
Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
Article CAS PubMed Google Scholar
Ishigami, Y. YumaIshigami/irscalc: First Release. https://doi.org/10.5281/zenodo.4673906] (2021).
Cock, P. J. et al. Biopython: freely available Python tools for computational molecular biology and bioinformatics. Bioinformatics 25, 1422–1423 (2009).
Article CAS PubMed PubMed Central Google Scholar
Li, H. et al. The sequence alignment/map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).
Article PubMed PubMed Central CAS Google Scholar
Anders, S., Pyl, P. T. & Huber, W. HTSeq–a Python framework to work with high-throughput sequencing data. Bioinformatics 31, 166–169 (2015).
Article CAS PubMed Google Scholar
Robinson, M. D., McCarthy, D. J. & Smyth, G. K. edgeR: a Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).
Article CAS PubMed Google Scholar
McCarthy, D. J., Chen, Y. & Smyth, G. K. Differential expression analysis of multifactor RNA-Seq experiments with respect to biological variation. Nucleic Acids Res. 40, 4288–4297 (2012).
Article CAS PubMed PubMed Central Google Scholar
Bitton, D. A. et al. AnGeLi: a tool for the analysis of gene lists from fission yeast. Front. Genet. 6, 330 (2015).
Article PubMed PubMed Central CAS Google Scholar
Zhang, Y., Werling, U. & Edelmann, W. Seamless ligation cloning extract (SLiCE) cloning method. Methods Mol. Biol. 1116, 235–244 (2014).
Article CAS PubMed PubMed Central Google Scholar
Suga, M. & Hatakeyama, T. A rapid and simple procedure for high-efficiency lithium acetate transformation of cryopreserved Schizosaccharomyces pombe cells. Yeast 22, 799–804 (2005).
Article CAS PubMed Google Scholar

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Acknowledgements

We thank members of Tom Suzuki’s laboratory, especially Yuriko Sakaguchi, for RNA-MS analysis, Kenjyo Miyauchi for RNA isolation with RCC, and Shunpei Okada for NGS data analysis. We thank Terumi Horiuchi and Kiyomi Imamura of Yutaka Suzuki laboratory for sample preparation, data generation, and data registration of NGS. Some computations were performed on the NIG supercomputer at the ROIS National Institute of Genetics. This work was supported by Grants-in-Aid for Scientific Research on Priority Areas from the Ministry of Education, Culture, Sports, Science, and Technology of Japan (MEXT), Japan Society for the Promotion of Science (JSPS) [20292782 and 18H05272 to T.S., 16H06279 (PAGS) to Y.S., and 18J13582 to Y.Ish.], and Exploratory Research for Advanced Technology (ERATO, JPMJER2002 to T.S.) from Japan Science and Technology Agency (JST).

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Department of Chemistry and Biotechnology, Graduate School of Engineering, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, Japan
Yuma Ishigami, Takayuki Ohira, Yui Isokawa & Tsutomu Suzuki
Department of Computational Biology and Medical Sciences, Graduate School of Frontier Sciences, The University of Tokyo, 5-1-5 Kashiwanoha, Kashiwa-shi, Chiba, Japan
Yutaka Suzuki

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Yuma Ishigami
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Y.Ish. performed biochemical, genetic, and bioinformatic works assisted by T.O. Y.Iso performed the poly(A)⁺ purification-nucleoside analysis experiment. Y.Ish. and T.S. wrote this paper. Y.S. performed NGS. All authors discussed the results and revised this paper. T.S. designed and supervised all work.

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Correspondence to Tsutomu Suzuki.

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Ishigami, Y., Ohira, T., Isokawa, Y. et al. A single m⁶A modification in U6 snRNA diversifies exon sequence at the 5’ splice site. Nat Commun 12, 3244 (2021). https://doi.org/10.1038/s41467-021-23457-6

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Received: 26 November 2019
Accepted: 29 April 2021
Published: 28 May 2021
DOI: https://doi.org/10.1038/s41467-021-23457-6

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