A molecular brake that modulates spliceosome pausing at detained introns contributes to neurodegeneration

Dawei Meng; Qian Zheng; Xue Zhang; Li Luo; Yichang Jia

doi:10.1101/2022.01.11.475943

Abstract

Emerging evidence suggests that intron-detaining transcripts (IDTs) are a nucleus-detained and polyadenylated mRNA pool for cell to quickly and effectively respond to environmental stimuli and stress. However, the underlying mechanisms of detained intron (DI) splicing are still largely unknown. Here, we suggest that post-transcriptional DI splicing is paused at B^act state, an active spliceosome but not catalytically primed, which depends on SNIP1 (Smad Nuclear Interacting Protein 1) and RNPS1 (a serine-rich RNA binding protein) interaction. RNPS1 and B^act component preferentially dock at DIs and the RNPS1 docking is sufficient to trigger spliceosome pausing. Haploinsufficiency of Snip1 attenuates neurodegeneration and globally rescues IDT accumulation caused by a previously reported mutant U2 snRNA, a basal spliceosomal component. Snip1 conditional knockout in cerebellum decreases DI splicing efficiency and causes neurodegeneration. Therefore, we suggest that SNIP1 and RNPS1 form a molecular brake for the spliceosome pausing, and that its misregulation contributes to neurodegeneration.

Introduction

Introns are spliced from eukaryotic messenger RNA precursors (pre-mRNA) by the spliceosome via two transesterification reactions - branching and exon ligation (1). During these reactions, the spliceosome undergoes structural and compositional dynamics (2–4). Firstly, the 5’ splice site (SS), branch site (BS), and 3’SS of an intron are recognized by the U1 small nuclear RNA (snRNA), SF1, and U2AF, respectively (E complex). Then, U2 small nuclear ribonucleoprotein particles (snRNPs) are recruited by the E complex to the BS (A complex), which then binds to U4/U6.U5 tri-snRNP to form the fully assembled and pre-catalytic spliceosome (B complex). The resulting B complex is empowered by an ATPase/helicase Brr2, remodeling to the active spliceosome (B^act). Through additional remodeling by the ATPase/helicase Prp2, B^act matures into a catalytically activated spliceosome (B*), in which the branching reaction occurs. During the B-to-B^act and B^act-to-B* transitions, a number of proteins are loaded/unloaded into the spliceosome, including more than 10 proteins recruited into early B^act and release of SF3a, SF3b, and pre- mRNA REtention and Splicing (RES) complexes from B* (5–8). The resulting catalytic step I spliceosome (C complex) is remodeled by the ATPase/helicase Prp16 into a step II catalytically activated spliceosome (C* complex), in which the exon ligation reaction occurs.

In contrast to constitutive splicing, > 90% of human multiexon genes undergo alternative splicing (AS) (9, 10), which contributes to proteomic diversity (11, 12). Accuracy in the recognition of reactive splice sites must be compromised by flexibility in splice site choice during AS. As one of the major categories of AS, intron retention (IR) was originally thought to be nonproductive for protein production, because it often introduces a premature termination codon (PTC) into the transcript, which is subsequently targeted for degradation by nonsense-mediated decay (NMD), a cytoplasmic mRNA surveillance mechanism (13, 14). However, Boutz et al. identified a group of intron-detaining transcripts (IDTs) in human and mouse cells that are polyadenylated, detained in the nucleus, and immune to NMD, and termed these incompletely-spliced introns as detained introns (DIs) (15). Nucleus DIs have also been documented by others and their splicing and subsequent mRNA export to the cytoplasm for protein translation has been associated with specific stimuli and stress (16–23). Given that gene size in human is large (∼27 kb in average) but RNA transcription rate is slow (∼2-4kb/minute) (24–27), post- transcriptional DI splicing is a quick and effective way for cells to adapt environment changes or stress. However, it remains unclear whether spliceosome is indeed paused at DIs, which catalytic step the spliceosome is paused at, what the molecular mechanisms underlie the spliceosome pausing, and what are the biological consequences of misregulation of this process.

Here, we disclose that more than one third of cerebellum-expressed genes transcribe IDTs and ∼90% of them only contain 1-2 intron(s). Using mouse forward genetics and gene knockout (KO), we demonstrate that haploinsufficiency of Snip1 (Smad Nuclear Interacting Protein 1) rescues IDT accumulation and neurodegeneration caused by a previously reported mutant U2 snRNA (28). SNIP1 interacts with protein components found in B^act spliceosome and protein components in peripheral exon junction complex (EJC), like RNPS1 (RNA binding protein with serine rich domain 1). Like Snip1, knockdown of Rnps1 rescue the DI accumulation while its overexpression is sufficient to trigger spliceosome pausing at DIs. B^act component and RNPS1 preferentially deposit at DIs and their surrounding sequences. Both RNPS1 docking at DIs and interaction between SNIP1 and RNPS1 are required for spliceosome pausing at DIs. Snip1 conditional KO in cerebellum reduces DI splicing efficiency and leads to IDT accumulation and neurodegeneration. Therefore, we suggest that SNIP1 and RNPS1 function as a molecular brake for spliceosome pausing at highly regulated DIs, and that misregulation of this process contributes to the pathogenesis of neurodegeneration.

Results

A mouse forward genetic screening identifies Snip1 as a modifier for NMF291 phenotypes

To understand how the previously reported mutant U2 (28) leads to global RNA splicing abnormalities and massive cerebellar granule cell loss, we established an ENU-induced mutagenesis screening for dominant modifier(s) that rescue the NMF291^-/- phenotypes (Fig. S1). A modifier (Snip1^M/+) partially rescued NMF291^-/- ataxia in a dominant manner (Fig. 1A and Movie 1). One of the modifier candidates in the family was a G to A substitution that alters the 5’ splice site (5’SS) GT of Snip1 exon 2 to AT (Fig. 1B), which completely segregated with the rescue in the NMF291^-/- mice (Table S1). Because the ENU mutation disrupts the 5’SS, we generated a Snip1 KO mouse line by Crispr- Cas9 to examine whether Snip1^M/+ rescues NMF291^-/- phenotypes through a loss-of-function mechanism (Fig. 1C). Heterozygous Snip1 KO (Snip1^-/+) rescued NMF291^-/- ataxia and significantly extended NMF291^-/- life span, comparable to the extent of Snip1^M/+ (Fig. 1D). In addition, Snip1^-/+ partially rescued neuron loss in the NMF291^-/- cerebellum (Fig. 1E), although Snip1^-/+ itself did not show cerebellar neuron loss. We failed to harvest a homozygous Snip1 mutant mouse for both ENU-induced mutation and Crispr-Cas9- generated KO (Table S2), indicating that Snip1 is an essential gene and ENU- induced mutation is possibly a null allele. Therefore, we conclude that haploinsufficiency of Snip1 rescues neurodegenerative phenotypes shown in the NMF291^-/- mutant mouse.

Figure 1. A forward genetic screening identifies Snip1 as a modifier for NMF291 phenotypes.

(A) A NMF291 modifier family pedigree. An ENU-induced mutant G1 male (NMF291^-/-;M/+) carrying less ataxia phenotype (rescued, in green) was bred to an ENU-untreated NMF291^-/+ (in white) female. The resulting NMF291^-/- mice were included for our phenotyping. The modifier is inherited in a dominant manner.

(B) The ENU-induced modifier candidate (M/+) hits the 5’SS (from GT to AT) of Snip1 exon2 (ENSMUST00000052183.6).

(C) Generation of Snip1 KO mouse (Snip1^-/+) by Crispr/Cas9. Snip1 contains 4 protein-coding exons and two functional domains. Coiled-coils, 157-184 a.a.; FHA (forkhead-associated domain), 268-331 a.a.. The Cas9 editing site was labeled as a red bar. A nucleotide insertion (+T) in Snip1 exon3 causes out-of-frame KO. PAM site was labeled in red and gRNA sequences were underlined. Codons were labeled with gray rectangles.

(D) The survival curves for the indicated genotypes (***p<0.001, N.S., no statistical significance, Log-rank test). Both the ENU-induced mutation (M/+) and Crispr/Cas9-generated Snip1 KO (Snip1^-/+) extended the NMF291^-/- mutant lifespans.

(E) Hematoxylin and eosin-stained cerebellar sagittal sections of the indicated genotypes at 4 months (4 Mos.) of age. Lower panels, the coresponding high magnification images (lobule II). Scale bar, upper 500 μm; lower, 50 μm.

See also Figure S1, Movie 1, and Table S1 and Table S2.

Haploinsufficiency of Snip1 globally rescues IRs in NMF291 mutant cerebellum

One of pathological features in NMF291 mutant cerebellum is severe IRs (28). To quantitatively and globally interrogate whether Snip1^-/+ is able to rescue the IRs, we performed cerebellar RNA-seq with poly(A) selection in wildtype (+/+), Snip1^-/+, NMF291^-/-, and NMF291^-/-;Snip1^-/+ mice at one month of age, when the expression of mutant U2 snRNAs starts to be upregulated and IRs become severe (28). We employed intron retention index (IRI) to represent the IR levels, which was calculated by the ratio of intronic reads normalized to flanking exonic reads (28–30). Consistent with a previous report (28), the IRI ratio of NMF291^-/- to +/+ showed more IRs in the NMF291^-/- cerebella (Figs. 2A and S2A-S2C), regardless of whether considering high (FC > 1.2 and padj < 0.1) or low (FC >1.2 and padj ≥ 0.1) confidence IR events. As reported before (28), about half of high confidence IRs were small introns (intron length < 150bps). The comparative IRI ratio of Snip1^-/+ to +/+ showed a slight depletion of IRs in Snip1^-/+ cerebella (Fig. 2B). Pairwise comparison between NMF291^-/- and NMF291^-/-;Snip1^-/+ indicated that the majority of IRs was rescued by Snip1^-/+ (Figs. 2C and S2A-S2C). Indeed, the IRI ratio of NMF291^-/-;Snip1^-/+ to +/+ revealed even fewer IRs shown in NMF291^-/-;Snip1^-/+ cerebella than that of +/+ (Fig. 2D). Among these high confident but not rescued IRs (493 shown in Fig. 2D), 74.0% are small introns with a much higher IRI ratio (median = 8.2) than that of the rest (median = 4.5), suggesting that these IRs are insensitive to haploinsufficiency of Snip1.

Figure 2. Haploinsufficiency of Snip1 partially rescues the NMF291 cerebellar IRs and the majority of them also exist in wildtype cerebellum.

(A-D) Pairwise comparisons of cerebellar IRI between the indicated genotypes. We set two separate cutoffs colored by red and gray, respectively. Red dots: events with FDR adjusted p-value (padj) < 0.1 and the IRI fold-change (FC) > 1.2; gray dots: padj ≥ 0.1 and the IRI FC > 1.2. Only events with IRI > 0.1 were included for the comparison. The retained introns less than 150bps in length were highlighted with rectangles and their percentage of all the IRs (red rots) was labeled. Mice, male, 1 month of age, n=3.

(E) Heatmap represents IRI changes across the indicated genotypes. Z-score was used to normalize IRIs in each row. The numbers of IRs and their corresponding genes (Genes) are shown.

(F and G) Cerebellar IRs (IRI > 0.1, F) of the indicated genotypes and ages and their overlaps (G).

(H) GO analysis of gene clusters shown in (E). The values in X axis are -Lg adjusted p-values. Gene numbers in each term were labeled. Some of gene names were labeled in the plot.

Intron-containing transcripts overrepresented in mutant U2 cerebellum are IDTs featured by nuclear-localized, incompletely spliced, and polyadenylated

Intron-containing transcripts often contain PTCs, triggering NMD for degradation (14, 31). However, in NMF291 mutant cerebellum, intron- containing transcripts are abundant and stable (28) (Figs. 2 and S2), which prompted us to examine whether these transcripts are targeted by NMD. To this end, we cultured Neuro2a (N2a) cells, a mouse neuroblastoma cell line, and transfected the cells with wildtype (WT-U2) and mutant U2 (Mu-U2) snRNA expressing plasmids. Compared to WT-U2, expression of Mu-U2 increased IRs at several endogenous sites, including Ptbp1 intron5 (Fig. S3A). The IRs were not sensitive to inhibition of NMD, either by addition of cycloheximide (CHX) in the culture medium or knockdown of Upf1 (Figs. S3B-S3E), suggesting that they are stable and detained in nucleus (14, 15). To test this possibility, we performed nuclear and cytosolic fractionation and examined IRs (Fig. S3F). Irrespective of WT- or Mu-U2 expression, IRs were enriched in the nuclear fraction. Therefore, we suggest that these incompletely spliced introns are DIs, previously featured by polyadenylated, detained in the nucleus, and immune to NMD (15).

To globally examine the nuclear enrichment of IDTs, we performed RNA- seq with poly(A) selection in isolated nuclear and cytosolic fractions in N2a cells (Figs. 3A and 3B). The IRI ratio of the nucleus to the cytosol indicated a global nuclear enrichment of polyadenylated IDTs in these cells regardless of whether WT-U2 or Mu-U2 was expressed. Consistent with what we observed in the NMF291 mutant cerebellum, Mu-U2 expression in N2a cells increased the number of DIs (Figs. 3C). Genes transcribing these IDTs are functionally involved in RNA metabolism/processing and cellular response to stress (Fig. 3D), similar to what we observed in vivo (Fig. 2H). In fact, a majority of the DIs (56.6%) in the NMF291 mutant cerebellum also appear in N2a cells expressing Mu-U2 (Fig. 3E).

Overrepresented IDTs in NMF291 mutant cerebellum (Figs. 2 and S2) allowed us to examine whether these polyadenylated IDTs produce protein or not with high confidence. Ptbp1 transcripts with intron 5 were a minor form in the wildtype cerebellum but became dominant in the mutant (Figs. S2C and S4A). Using a PTBP1 N-terminal antibody, we detected comparable amounts of PTBP1 (∼50kD) in wildtype and mutant cerebella but failed to detect the corresponding truncated protein supposedly produced from Ptbp1 intron 5- containing transcripts (Figs. S4B and S4C), supporting the idea that the IDTs are detained in nucleus with limited accessibility to cytoplasmic protein translation machinery. To globally examine the protein products derived from DIs, we generated a customized peptide database, including peptides encoded by the DIs found in wildtype and NMF291 mutant cerebellum and their upstream exons (Fig. S4D). Although we retrieved thousands of peptides coded by the upstream exons, we failed to retrieve any peptide encoded by the DIs from three biological replicates of the wildtype and NMF291 mutant cerebellar protein lysates (Fig. S4E). Therefore, we suggest that the majority of intron- containing transcripts over-represented by expression of mutant U2 are IDTs featured by nuclear-localized, incompletely spliced, and polyadenylated.

More than 1/3 of cerebellum-expressed genes transcribe IDTs that are highly regulated and ∼90% of these IDTs only contain 1-2 intron(s)

To test whether DIs are regulated during cerebellum development and aging, we analyzed cerebellar DIs (IRI > 0.1) at P5, 10, 30, and 2.5-year by using RNA-Seq and grouped them into four clusters (Fig. 3F). Every developmental age point has their unique IDTs transcribed by genes involved in several cellular processes, including RNA splicing, cellular response to stress, histone modification, and ribosome biogenesis (Fig. 3G). To examine how much percentage of cerebellum-expressed genes have DIs, we extracted 8917 genes with reasonable expression (the DI surrounding exon reads > 20) in P5, 10, 30, and 2.5-year cerebella (Fig. 3H). Among them, 34.2 - 43.2% have reliable DIs (IRI > 0.1) with higher percentages in P5 and 2.5-year cerebella. The IRI ratio of 2.5-year to P30 wildtype indicated an enrichment of IDTs in aged cerebellum (Fig. 3I). However, compared to the DIs overrepresented in mutant cerebellum, less detention appears in aged cerebellum, with smaller mean value of log2 IRI ratio (0.32 for aged, 0.48 for NMF291^-/- cerebellum, both compared to that of P30 +/+) (Fig. 3J). To understand the details of DIs at single transcript level, we employed nanopore sequencing, a long-read sequencing technology (32), to detect the DI features in P30 and 2.5-year wildtype and NMF291^-/- cerebella. To call for full length IDTs, we only included the nanopore reads containing both 5’UTR and 3’UTR with IRI > 0.05. The majority of these DIs (73%) shown in the aged cerebellum also appear in that of NMF291 mutant cerebellum (Fig. S5A). In addition, higher percentage of IDTs of all full length transcripts we examined showed in both aged (16.0%) and NMF291 mutant (17.5%) cerebella (Fig. S5B), compared to that of P30 wildtype. These indicate that the DIs are primarily affected during aging process, presumably when the function of spliceosome declines.

Next, we asked whether these DIs are evolutionarily conserved between mouse and human. We categorized the efficiently-spliced introns (IRIs < 0.02) and the DIs (IRIs > 0.1) in the NMF291^-/- mutant cerebella. Compared to the efficiently-spliced introns, the DIs are significantly more conserved between human and mouse (p < 2.2 x 10^-16) (Fig. 3K). For intron length of less than 150 bps, DIs also showed significantly more conserved (p < 1.7 x 10^-7) than that of efficiently-spliced introns.

To further learn the DI features at full length transcript level, we retrieved 177002 IDTs for wildtype (+/+) and 334999 for NMF291 mutant (NMF291^-/-) cerebella by using nanopore sequencing. Consistent with our 2^nd generation RNA-seq results, nanopore-reads showed more IDTs in mutant cerebellum than that of wildtype (Fig. 3L). However, when we categorized the IDTs in terms of their intron number distribution, most IDTs contained 1-2 intron(s) in wildtype (91%) as well as in mutant (90%) cerebella (Fig. 3M). In addition, the percentage of intron number distribution between the two genotypes is almost identical with ∼70% IDTs containing only one intron in both genotypes. Taken together, our findings reveal that 1) over 1/3 of cerebellum-expressed genes transcribe IDTs and their DI splicing is highly regulated during cerebellum development and aging; 2) evolutionarily these DIs are more conserved than that of efficiently-spliced; and 3) ∼90% of IDTs only contain 1-2 intron(s), indicating that DIs only comprise a small proportion of the total introns transcribed in cerebellum.

Nuclear-localized SNIP1 binds to cerebellar polyadenylated IDTs

Previous studies showed that SNIP1 interacts with the TGF-β family Smad proteins, NF-κB transcription factor p65, and transcriptional coactivators p300/CBP, resulting in regulation of TGF-β and NF-κB signaling (33, 34). Post- transcriptionally, SNIP1 also regulates Cyclin D1 RNA stability (35). However, how SNIP1 regulates pre-mRNA splicing per se are not studied in mammal. If DI splicing is regulated by SNIP1, we speculated that SNIP1 must bind to these transcripts. To this end, we inserted 3 × Flag tag at the C-terminal end of Snip1 by Crispr-Cas9-mediated homologous recombination (Figs. 4A, S6A, and S6B). SNIP1-Flag appeared in the knockin (KI) cerebellum at the expected molecular weight, and was absent in the wildtype control (Fig. 4A). The cerebellar expression level of SNIP1-Flag at P5 and P10 was 5.0 and 3.6 times higher than that of P30 (Fig. 4A). Ubiquitous expression of SNIP1 was documented in various adult mouse tissues (Fig. S6C).

Figure 4. Nucleus-localized SNIP1 binds to cerebellar IDTs.

(A) The expression level of SNIP1 was documented with a Flag antibody at various age points in Snip1-Flag KI animals. GAPDH, loading control; f/+, mice heterozygous for Snip1-Flag KI; +/+, negative control for Flag immunoblot. A summary of the expression level was inserted.

(B) Flag-immunostaining was performed in Snip1-Flag KI mouse cerebellum at one month and P7 of ages. NeuN, a neuronal marker; +/+, negative control for Flag immunostaining. Scale bar for one month, 30 μm; for P7, 20 μm.

(C) A diagram for native Flag-RIP-seq. The RNA/protein complex containing Flag-SNIP1 was eluted by competition with free Flag peptides. The resulting Flag-SNIP1-bound RNAs were applied for Smart-seq2 to amplify full-length polyadenylated mRNAs. M2, M2-beads conjugated with Flag-antibody.

(D and E) RIP was performed with cerebellar protein lysates from NMF291^-/+;f/f or NMF291^-/+ mice at one month of age. Enrichment of DIs was detected by RT-PCR in RIP products from NMF291^-/+;f/f mice but not from that of NMF291^-/+ controls. Global examination of the enrichment by Flag-RIP-seq from three biological replicates (E).

(F-G). Two representative SNIP1-bound IDTs visualized by IGV.

(H and I) Global examination of enrichment of SNIP1-bound DIs in P7 Snip1^f^/f cerebellum by Flag-RIP-seq (n = 3). Wildtype (+/+) animals served as negative control for Flag-RIP-seq. GO analysis of the genes transcribing SNIP1-bound IDTs shown in (I).

In A, the values are presented as mean ± SEM (mice, n ≥ 3 for each age point). *p<0.05, **p<0.01, ANOVA, SPSS.

DI splicing is paused prior to the first catalytic step

To understand how SNIP1 regulates DI splicing, we examined SNIP1 protein binding partners in N2a cells, a cellular model that carries endogenous DIs (Fig. S3). In N2a cells stably expressing SNIP1-Flag, we performed Flag-co- immunoprecipitation (co-IP) followed by mass spectrometry (co-IP/MS). High confidence hits from three biological replicates were protein components found in spliceosome and peripheral EJC, or proteins involved in RNA export (Fig. 5A and Table S3). These SNIP1-interacting protein partners include RNA helicases (BRR2 and SNU114) and SR/SR-like proteins (SFRS16, SRM300, ACIN1, RNPS1, and SRSF7). We noticed that protein components found in spliceosome are that of B^act, the activated spliceosome, but not that of B*, the catalytically activated spliceosome (7, 37, 38) (Fig. 5B). Given that SNIP1 binds to IDTs (Fig. 4) and recent resolved human spliceosome structure also support the presence of SNIP1 in B^act spliceosome but not B and C complex (7, 37, 39–41), we proposed that DI splicing is paused at B^act.

Figure 5. DI splicing is likely paused at B^act.

(A) Potential SNIP1 interacting proteins identified by Flag-IP coupled with mass spectrometry (co-IP/MS) in N2a cells expressing Flag-SNIP1. Many of these protein partners are involved in pre-mRNA splicing and RNA metabolism (also see Table S3).

(B) The potential SNIP1 interacting partners are found in B^act but not B*. Factors loaded into B* are labeled in gray, which were not hit by our co-IP/MS. (C and D) Interactions between SNIP1 and protein components found in B^act component SF3a120 (C) but not those in B* component YJU2 (D) were validated in N2a cells expressing the indicated tagged proteins.

(E) RIP was performed in N2a cells expressing Flag-SF3a120, Flag-YJU2, or EGFP. DIs were measured by RT-PCR using Smart-amplified cDNA.

(F-I) Flag-RIP-seq was performed in N2a cells expressing Flag-SF3a120 (F) and Flag-YJU2 (G). N2a cells expressing EGFP, a negative control for our Flag-RIP-seq. Two representative SF3a120-bound DIs visualized by IGV (H and I).

SNIP1 and RNPS1 function as molecular brake for spliceosome pausing at DIs

As we documented above, haploinsufficiency of Snip1 partially rescues DIs from accumulating in NMF291 mutant cerebellum (Fig. 2). In N2a cells, knockdown of Snip1 by shRNA (sh-Snip1) also reduced the levels of several endogenous intron detention events that were amplified by the expression of Mu-U2 (Figs. S8A and S8B). In addition to protein components found in B^act, protein components of the peripheral EJC (ACIN1, PNN, and RNPS1) were also identified as potential SNIP1 interacting partners (Table S3). Like Snip1, knockdown of Pnn and Rnps1 significantly reduced the intron detention amplified by Mu-U2 (Figs. 6A and 6B and S8C). Interaction of SNIP1, PNN, and RNPS1 was confirmed by co-IP in N2a cells simultaneously expressing tagged PNN, SNIP1, and RNPS1 (Fig. S8D), suggesting that these proteins work together to regulate spliceosome pausing. Although splicing factor SRm300 was found by our co-IP/MS (Table S3), SRm300 knockdown did not significantly influence the intron detention we examined (Figs. 6A and 6B).

Figure 6. SNIP1 and RNPS1 function as a molecular brake for spliceosome pausing at DIs.

(A and B) Effect of the knockdowns of genes encoding peripheral EJC components (Acin1, Pnn, and Rnps1) and a splicing factor (SRm300) on the DIs (A) amplified by expression of mutant U2 (Mu-U2) in N2a cells and the data summary (B).

(C) Annotated domains of SNIP1 and RNPS1 (upper). Co-IP was performed in N2a cells expressing the Flag-tagged full-length or FHA domain truncated SNIP1 (ΔFHA) together with RNPS1-Myc (lower).

(D) N2a cells were infected with scrambled shRNA or shSnip1 and transfected with the indicated expression plasmids.

(E) Flag-RIP was performed with protein lysates from N2a cells infected with Flag-tagged full-length or ΔFHA SNIP1 and transfected with Mu-U2 expression plasmid. RT, reverse transcriptase.

(F) Co-IP was performed in HEK293 cells expressing HA-SNIP1 and/or Flag- tagged RNPS1, including full length and various domain-truncated RNPS1.

(G) Flag-RIP was performed with protein lysates from N2a cells expressing the indicated proteins. Smart-amplified cDNA was analyzed by RT-PCR.

(H and I) Global examination of the RNPS1-bound DIs in N2a cells expressing Flag-RNPS1. Control, naïve N2a cells. Two representative RNPS1-bound DIs visualized by IGV (I).

(J and K) HEK293 cells were transfected with previously described splicing minigene reporters derived from L1cam detained intron 27 (int27) and constitutively spliced intron 28 (int28) (PMID: 22265417). The data summary was shown in (K).

(L and M) CLIP-seq read density plot for RNPS1 (L) and SF3a60 (M). Read density plot with DIs (IRI > 0.2) shown in red; read density plot with efficiently- spliced introns (IRI < 0.05) shown in blue; read density plot regardless of IRI shown in gray. RNPS1 iCLIP data from ArrayExpress (PMID: 27475226) and SF3a60 eCLIP data from ENCODE (PMID: 27018577).

(N and O) Read density tracks along two representative genes RPL10A (N) and HSP90B1 (O) viewed by IGV and scaled by RPM (reads per million usable). The most significant clusters called by CLIPper and illustrated by the red lines. In A, D, E, G, and J, DIs were measured by RT-PCR. Unspl., unspliced transcripts; spl., spliced transcripts. In B and K, the values are presented as mean ± SEM, n=3. *p<0.05, **p<0.01, ***p<0.001, N.S., no statistical significance, ANOVA, SPSS.

RNPS1 and SF3a60 dock at different positions of DIs

EJC core proteins and peripheral EJC component RNPS1 have been documented to bind to mRNA and involved in post-transcriptional mRNA processing (46–51). To examine the binding sites of RNPS1 and SF3a complex at DIs, we reanalyzed previous reported RNPS1- and SF3a60- cross-linking and immunoprecipitation (CLIP) data (52–54). Globally, RNPS1 CLIP-seq reads piled up at 5’-exons (with a binding peak at -14 nts of 5’SS) and 5’-introns (between 0 to 30 nts of 5’SS) of the DIs (IRI > 0.2) (Fig. 6L), slightly differing from the binding site of core EJC (−24 to −20 nts from the 5’SS) (47, 54). For SF3a60 - a protein component of the SF3a complex - the CLIP-seq reads concentrated between -40 to 0 nts of the 3’SS of DIs, with a peak at -30 nts (Fig. 6M). However, such RNPS1- and SF3a60-CLIP-seq peaks were less represented at the positions of the efficiently spliced introns (IRI < 0.05) or all the introns we examined (All).

For representative CLIP peaks at DIs, IGV illustrated that intron 4 of RPL10A and intron 13 of HSP90B1 are the highest confidence DIs compared to neighboring introns in both Hela and HepG2 cells (Figs. 6N and 6O). The most stringent RNPS1 binding peaks identified by CLIPper appeared at the 5’ exon/intron boundary of the highest confidence DIs. The SF3a60 peaks were located inside of the highest confidence DIs close to the 3’SS. Taken together, our analysis suggests that RNPS1 a peripheral EJC component and SF3a60 an SF3a complex component preferentially dock at DIs.

Conditional KO of Snip1 in cerebellar granule cells leads to IDT accumulation and neurodegeneration

SNIP1 is expressed in granule cells in the developing and adult cerebellum (Figs. 4A and 4B) and homozygous Snip1 KO leads to embryonic lethality (Table S2). To reveal the biological consequences of Snip1 KO in the cerebellum, we generated a Snip1 floxed mouse line and crossed it to Nse-CreER^T2, a tamoxifen-inducible Cre line specific for Cre expression mainly in cerebellar granule cells and a few granule neurons in the hippocampal dentate gyrus (Figs. S9A and S9B) (55). We confirmed the specificity of this system by crossing the Nse-CreER^T2 to a Cre-dependent reporter line, H2B mCherry (56) (Fig. S9C). After tamoxifen injections on P3, P4 and P5, we observed a deletion of Snip1 exon 3 at both the genomic DNA and mRNA levels, specifically in the cerebellum and hippocampus but not in the cortex of Snip1^fl^/^fl;NseCreER^T2 mice on P9 (Fig. S9D). Such deletions did not appear in the brain regions in age- matched Snip1^fl^/^fl controls. We harvested P7 cerebella from Snip1^fl^/^fl;NseCreER^T2 and age-matched Snip1^fl^/^fl mice after tamoxifen injections and performed RNA-seq (Fig. 7A). Accumulation of intron-containing transcripts were evident in the Snip1 conditional KO (cKO) cerebella compared to controls. Genes transcribing these transcripts are functionally involved in histone modification, DNA repair, RNA splicing, ribosome biogenesis, and synaptic vesicle transport (Fig. 7B), similar to what we observed in the NMF291 mutant cerebellum (Fig. 2H). To understand the details of incompletely-spliced introns induced by Snip1 cKO at the single transcript level, we sent the cKO cerebella for nanopore sequencing. Similar to what we observed in wildtype and NMF291 mutant cerebella (Figs. 3K-3L), most intron-containing transcripts had one or two intron(s) in both Snip1 cKO (for one, 74%; for two, 16%) and an age-matched control (Snip1^fl^/^fl, for one, 78%; for two, 17%) (Fig. 7C). These data suggest that like the NMF291 mutation, Snip1 cKO in cerebellar granule cells leads to accumulation of IDTs, but has less effect on constitutive splicing. Indeed, the majority of intron-containing transcripts (65.7%, 3959/6026) present in Snip1 cKO cerebella on P7 are also found in NMF291 cerebella on P30 (Fig. 7D).

Figure 7. Snip1 cKO in cerebellar granule cells leads to IDT accumulation and neurodegeneration.

(A) A schematic diagram for Cre-dependent Snip1 cKO (upper). Cerebellar RNAs extracted from the indicated genotypes on P7 (n = 3) were applied for RNA-seq. Events with IRI > 0.1 were included for clustering.

(B) GO analysis of genes transcribing intron-containing transcripts that are over-represented in Snip1 cKO cerebellum.

(C) The cerebellar RNAs described in (A) were applied for nanopore sequencing. Intron number distribution of the indicated genotypes was plotted.

(D) Intron-containing transcripts detected by nanopore sequencing in P7 Snip1 cKO cerebellum are shared by that in P30 NMF291^-/- cerebellum.

(E) Our experimental procedure: tamoxifen injections on day 1 (D1), 2, 3, 4, and 5 after 3 months (Mos.) of age; mouse phenotyping on D9 (upper). Hematoxylin and eosin-stained cerebellar sagittal sections of the indicated genotypes (lower).

(F) Previously reported DECKO (Double Excision CRISPR Knockout, PMID: 26493208) approach was adopted for Snip1 KO in N2a cells. Snip1 dual gRNAs target the exon 3 (upper). The dual gRNAs are driven by U6 and H1 promoters, separately (lower). F and R, primers for Crispr/Cas9-mediated editing site fusion.

(G and H) N2a cells expressing L1cam minigenes were transfected with Mu- U2 or Snip1 dual gRNAs plasmids. Cas9 was constitutively expressed in these cells. DIs were measured by RT-PCR. Unspl., unspliced transcripts; spl., spliced transcripts. Δ, Crispr/Cas9-mediated editing site fusion at RNA level measured by RT-PCR. Primers used for fusion detection were labeled in (F). The data summary shown in (H).

In E, the corresponding high magnification images from lobule II. Scale bar, upper 200 μm; lower, 100 μm. In H, data are presented as mean ± SEM, **p<0.01, n = 4, one-way ANOVA, SPSS.

Discussion

Here, we describe interaction between SNIP1 and RNPS1 as a molecular brake for spliceosome pausing at highly regulated DIs. Together with a previous report (28), we suggest that misregulation of this process contributes to pathogenesis of neurodegeneration.

The yeast homolog of SNIP1 is Pml1 (pre-mRNA leakage protein 1). Compared to human SNIP1 (396 a.a.), yeast Pml1 (204 a.a.) lacks 1-180 a.a. N-terminus and has as low as ∼30% similarity to human SNIP1. Yeast RES complex contains Snu17, Bud13, and Pml1, corresponding to human RBMX2, BUD13, and SNIP1 (58). RES complex is initially recruited to the B/B^act complex and released from B* (6, 7, 38, 44, 59-61). We demonstrate that SNIP1 interacts with protein components found in B^act but not B* (Figs. 5A-5D). In yeast, unlike spliceosome basal components, such as SF3b subunits, the RES complex is not essential for yeast growth (58, 62). In zebrafish, individual KO of bud13, snip1, and rbmx2 leads to mis-splicing a subset of introns (63). In mammalian cells, BUD13 binds to a group of retained-introns and regulates their splicing (64). It is plausible that SNIP1 functions together with the other two RES complex proteins to regulate spliceosome pausing at DIs, although our SNIP1 co-IP/MS experiments did not achieve reliable RBMX2 and BUD13 hits (Table S3). Given that histone modification plays an important role in alternative splicing regulation and SNIP1 regulates p300 histone acetyltransferase activity (33, 34, 65, 66), SNIP1 could also indirectly regulate DI splicing.

Previous studies demonstrated that RNPS1 interacts with NMD machinery for mRNA surveillance (54, 67–69). RNPS1 interacts with SAP18, ACIN1 or PNN to form the heterotrimeric apoptosis and splicing-associated protein (ASAP) complex (ACIN1, SAP18 and RNPS1) or the PSAP complex (PNN, SAP18 and RNPS1) (45, 70, 71). Both ACIN1 and PNN contain a similar RNPS1-SAP18-binding (RSB) motif to interact with RNPS1 and SAP18, suggesting that the formation of the ASAP and PSAP complexes is mutually exclusive and the resulting complexes determine the specificity of RNA substrate binding (45). This may explain why knock down of Rnps1 or Pnn but not Acin1 significantly reduced the intron detentions induced by the expression of mutant U2 (Figs. 6A and 6B). Our experiments did not support that SNIP1 reliably interact with the core EJC components (Figs. 5A and S6F). Given that ACIN1 and PNN are located in nuclear speckles - where post-transcriptional splicing occurs - it is plausible that RNPS1 regulates post-transcriptional splicing by temporarily forming the ASAP or PSAP complex, which is replaced by the core EJC complex during mRNA export (70, 72–75).

RNPS1 docking at DIs is sufficient to induce intron detention but has little effect on constitutive splicing (Figs. 6J and 6K). In addition, the interaction between SNIP1 and RNPS1 is required for spliceosome pausing at DIs (Figs. 6C-6J). We propose that SNIP1 and RNPS1 function a molecular brake for the spliceosome pausing (Fig. S10). That is why partial loss-of-function of SNIP1 or RNPS1, probably through releasing the molecular brake, rescues intron detentions caused by mutant U2. SNIP1 contains an FHA domain, which is involved in phospho-dependent protein/protein interaction (43, 44), and the FHA domain is required for the interaction between SNIP1 and RNPS1 (Fig. 6C). Therefore, phosphorylation/dephosphorylation status of RNPS1, especially in its S domain, may participates in the spliceosome pausing/resuming at highly regulated DIs. In fact, RNPS1 phosphorylation close to its S domain was proposed to regulate splicing in vitro and inhibition of SR protein kinases (Clk family members) were reported to promote the post- transcriptional splicing of DIs (15, 20, 76).

Like the NMF291 mutation, Snip1 KO decreases the splicing efficiency of DIs but has little effect on that of constitutively-spliced introns (Figs. 3A-3C and Fig. 7). These indicate that splicing of constitutively-spliced introns and DIs has different kinetics. Most likely, constitutive introns are co-transcriptional spliced but DIs undergo post-transcriptional splicing (72) (Fig. S10). In yeast, RES complex is not required for B complex formation but for the efficient transformation from B to B^act (77). Here, we suggest that SNIP1 may have two sides in regulation of DI splicing: 1) to form molecular brake together with RNPS1 to pause the spliceosome at DIs; 2) to facilitate the spliceosome transformation at DIs. Due to different splicing kinetics, DI but not constitutive intron splicing is primarily affected in presence of mutant (the NMF291^-/-) or dysfunction (Snip1^-/-) spliceosome (Fig. S10). Partial loss of SNIP1 or RNPS1 rescues mutant-U2 caused intron detentions probably through releasing the molecular brake, while complete removal of Snip1 may impair the spliceosome transformation at DIs, thus lower the splicing efficiency (Fig. S10). Global accumulation of DIs will sequester spliceosome pausing complex, which in turn lowers the available spliceosome abundance and worsens DI splicing. Indeed, decreased amounts of core and non-core spliceosome components leads to accumulation of intron-containing transcripts (78). In addition, accumulation of intron-containing transcripts has been documented in patient brains with neurodegenerative diseases (79, 80). Given that genes involved in RNA metabolism/processing and cellular response to stress tend to transcribe IDTs, global accumulation of DIs will in turn further damages these gene functions. The mechanisms underlying post-transcriptional DI splicing we describe here may help to improve the understanding of pathogenesis of neurodegenerative diseases and to develop intervention strategies in the future.

Materials and methods

Mice

All the mice described in this work were maintained on the C57BL/6J background (JAX, Stock No. 000664) and housed in Tsinghua Animal Facility that has been accredited by the Association for the Assessment and Accreditation of Laboratory Animal Care International (AAALAC) since 2014. All animal protocols were approved by the Institutional Animal Care and Use Committee (IACUC) at Tsinghua University based on Guide for the Care and Use of Laboratory Animals (Eighth Edition, NHR). Mice were housed in isolated ventilated cages (maximum six mice per cage), on a 12/12-hour light/dark cycle, 22-26°C, 40–70% humidity with sterile pellet food and water ad libitum. Cages were checked daily to ensure animal welfare. Body weight was assessed regularly to ensure no weight loss. Whenever animals were used for research, we followed the 3Rs (replacement, refinement or reduction) rules. The C57BL/6J and ICR mice were purchased from Charles River Laboratories, Beijing, China. The Nse-CreER^T2 line was imported from The Jackson Laboratory (JAX, Stock No. 022763) (55). For tamoxifen injection, we followed previous reports (81, 82). In brief, tamoxifen base (T5648-Sigma) was dissolved in corn oil and prepared freshly before use.

ENU mutagenesis and modifier identification

For ENU-induced mutagenesis, the NMF291^-/+ males were i.p. injected with ENU (80-110 mg/kg B. W.) for three consecutive weeks, as previously reported (83, 84). After an infertility test, the ENU-treated males were crossed to untreated NMF291^-/+ females for G1. Of these offsprings, the NMF291^-/- mice were used for behavioral tests, and the mice with less ataxia phenotype and improved lifespan were selected for family pedigree determination.

For identification of the modifier candidates, the ENU family members with or without phenotypic improvement were applied for exome sequencing. For exome capture and library construction, the instructions of NimbleGen SeqCap EZ Exome Library SR Platform (Roche) were followed. Briefly, genomic DNA was fragmented to 200-300 bps with ultrasonic shearing. End-repair, A-tailing, adapter ligation, and pre-capture ligation were performed by using the KAPA LTP Library Preparation Kit (Roche). After exome capture, the resulting samples were amplified by KAPA HiFi HotStart Ready Mix using Pre-LM-PCR Oligos 1 & 2. Libraries passing QC were sequenced (Illumina HiSeq 2500 platform) and data processing and variant discovery were performed by using GATK platform (85). Variant annotation and classification were achieved using ANNOVAR (86).

Generation of Snip1 KO, Flag-tag KI, and conditional KO mice

The CRISPR design website (http://crispr.mit.edu) was used to design gRNAs and avoid off-targets (87, 88). Cas9 mRNA, gRNA (s), and/or donor DNA were injected to C57BL/6J embryos. Then the injected embryos were transferred to the oviduct ampulla of pseudo-pregnant ICR female mice. For generation of the Snip1 KO mouse, the gRNA target sequence was: AGTGAGCGAGACCGGCACCGGGG (with PAM site underlined). For generation of the Snip1 Flag-tagged mouse, the gRNA target sequence was: GGGACGGTTTCTAACAGTAGAGG. Donor DNA had two homology arms (∼200bps each) flanking the mutant PAM site. For generation of the Snip1 floxed mouse, we employed multiple gRNAs to increase homologous recombination. The gRNAs were: gRNA-A1: GAGTCTAACTGGCCCTTCGGGGG, gRNA-A2: CAATGGTACCATCCTTTAACAGG, gRNA-B1: AGTGTGGTTCTTCCCCCGAAGGG, gRNA-B2: CTGTTAAAGGATGGTACCATTGG. Two LoxP sites were placed in the Snip1 intron2 and intron3, respectively, away from conserved intronic sequences. The donor DNA contained the targeted exon3, the flanking two LoxP sites, and two homology arms (∼800bps each). C57BL/6J mouse genomic DNA was used as a template to amplify the sequences and the homology arms in the donor DNAs. Mice with the right genotypes were further crossed to C57BL/6J mice for at least three generations to establish the lines.

Hematoxylin and eosin staining

Mice were anesthetized and transcardially perfused with PBS and then Bouin’s solution (Sigma-Aldrich). After post-fix in Bouin’s solution, the brain tissues were paraffin embedded. The paraffin sections were applied for hematoxylin and eosin (H&E) staining. The stained sections were scanned with Pannoramic Digital Slide Scanners (3DHISTECH).

Cell culture, plasmid construction, transfection, lentiviral infection, shRNA knockdown, and NMD reporter assay

HEK293FT and Neuro-2a cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM, Corning) complemented with 10% FBS, 1% penicillin–streptomycin in 5% CO2 at 37℃. For plasmid construction, genes were cloned into pCMV-3tag-4A and pCMV-3tag-1A (Agilent Technologies). In addition, we replaced EGFP and Cas9 from pLJ-EGFP and LentiCas9-Blast (Addgene) with genes of interest. Cell transfection was performed using Lipofectamine 3000 (Thermo Fisher Scientific). shRNA clones were purchased from lentiviral vector-based shRNA libraries (MISSION shRNA library, Sigma-Aldrich). Lentivirus packaging was performed as previous reported (89). The NMD reporter assay in N2a cells was performed as previous described (90). Briefly, N2a cells were transfected with pCI-Renilla/β-globin and pCI-firefly reporters (gifted from Drs. Gabriele Neu-Yilik and Andreas E. Kulozik). After 24 hours, cells were treated with100 μg/ml CHX or 0.01% DMSO for 5 hours. Renilla and firefly luciferase were detected using Dual-Luciferase Reporter System (Promega).

Antibodies and immunofluorescence

Antibodies used in this study were: Mouse anti-FLAG (Abmart, M2008H), rabbit anti-FLAG (Sigma, F7425), mouse anti-TUBULIN (Sigma, T6793), mouse anti- HA (Abcam, ab130275), rabbit anti-HA (Invitrogen, 715500), mouse anti-Myc (Abmart, M20002), rabbit anti-NeuN (Cell Signaling, 24307), anti-c-Myc Magnetic Beads (Thermo Fisher Scientific, 88842), donkey anti-mouse IgG secondary antibody (Alexa Fluor 555, Thermo Fisher Scientific, A31570), donkey anti-rabbit IgG secondary antibody (Alexa Fluor 555, Thermo Fisher Scientific, A31572), donkey anti-goat IgG secondary antibody (Alexa Fluor 488, Thermo Fisher Scientific, A11055). PTBP1 N-terminal antibody (PTB-NT, generated from 1-15 a.a.) was gifted from Dr. Douglas Black.

For tissue immunostaining, mice were anesthetized and transcardially perfused with PBS and then 4% paraformaldehyde. Brain tissues were submerged in 10%, 20%, and 30% sucrose solution for gradient hydration. The resulting tissues were embedded in OCT and cut on a cryostat. The brain sections were blocked in 3% BSA and then incubated with primary antibody overnight. After 0.5% PBS-T washes, the sections were incubated with Alexa Fluor-conjugated secondary antibodies (Thermo Fisher Scientific). For cultured cell staining, cells were fixed with 4% paraformaldehyde and blocked with blocking buffer. The fixed cells were incubated with primary antibodies overnight. Images were taken by Nikon A1 confocal microscopy.

Immunoblot, IP, and co-IP/MS

For immunoblot, fresh tissues were homogenized on ice with RIPA buffer (25 mM Tris-HCl pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS) complemented with protease inhibitor cocktail (Roche) and phosphatase inhibitor cocktail (PhosSTOP, Roche). For cell lysate, the cultured cells were washed with PBS first and then lysed with RIPA buffer. The resulting lysates were centrifuged at 12,000 × g for 10 minutes. Supernatant was boiled with 2 × SDS loading buffer for immunoblot loading.

For IP, mouse cerebellum was homogenized on ice with IP buffer (150 mM KCl, 25 mM Tris pH 7.4, 5 mM EDTA, 1% Nonidet P-40) complemented with protease and phosphatase inhibitor cocktail (Roche). Cultured cells were washed with ice-cold PBS three times and lysed with IP buffer on ice. Tissue and cell lysates were rotated at 4℃ for 0.5 hour and then centrifuged to remove cell debris. The supernatant was collected and incubated with anti-FLAG M2 magnetic beads (M8823, Merck) 4℃ overnight. The magnetic beads were washed with IP buffer twice and TBS buffer (50 mM Tris-HCl, 150 mM NaCl, pH 7.4) twice. The resulting samples were boiled with 2 × SDS loading buffer.

For MS, samples were loaded on SDS-PAGE gels for separation, stained with SimplyBlue™ SafeStain (Life technologies), and excised. The gel slices were reduced and in-gel digested with sequencing grade modified trypsin (Promega). The resulting samples were quenched by 10% trifluoroacetic acid and peptides were extracted and dissolved in 0.1% trifluoroacetic acid. For LC- MS/MS, the purified peptides were separated by a C18 column (75 μm inner diameter, 150 mm length, 5 μm, 300 Å) and directly connected with an Orbitrap Fusion Lumos mass spectrometer (Thermo Fisher Scientific). Mobile phase A was an aqueous solution of 0.1% formic acid, and mobile phase B was 0.1% formic acid in acetonitrile. The MS/MS spectra were searched against the Uniport mouse database using Proteome Discoverer (version PD1.4, Thermo Scientific™). The peptide spectrum match (PSM) was calculated by Percolator provided by Proteome Discoverer, and only peptide FDR less than 0.01 was included for further analysis. Peptides only assigned to a given protein group were considered as unique. FDR was also set 0.01 for protein identifications.

Proteome analysis for potential peptides encoded by DIs and their upstream exons

For proteome analysis, cerebellum was quickly removed and placed in a Dounce homogenizer. Tissues were lysed with 8 M urea in PBS supplemented with protease and phosphatase inhibitor cocktail (Roche) at room temperature. Lysates were centrifuged at 12,000 rpm for 10 minutes and supernatant was collected. Protein concentration was determined by BCA Protein Assay Kit (Pierce™). The tryptic peptides were fractionated with a XBridgeTM BEH300 C18 column (Waters, MA). LC-MS/MS analysis was similar to what we described above. To identify potential peptides encoded by DIs, a customized peptide database was generated as previously reported (29). Briefly, all DI events were extracted from wildtype and NMF291^-/- RNA-Seq results. The RNA sequences were in silico translated to peptides from 60 nucleotides upstream of DIs, until a PTC is encountered. The MS/MS spectra from each LC-MS/MS run were searched against the customized peptide database by the Sequest HT search engine of Proteome Discoverer, to identify potential peptides encoded by intronic sequences. Trypsin was specified as the proteolytic enzyme, and we allowed up to two missed cleavage sites. PSM was validated using Percolator, and only FDR < 0.01 was considered correct.

Total RNA extraction, reverse transcription, real-time qPCR, and nuclear and cytoplasmic RNA extraction

Total RNA was extracted with TRIzol reagent (Thermo Fisher Scientific). The extracted RNA was dissolved in nuclease-free water and then treated with DNase (RQ1 RNase-Free DNase, Promega). Reverse transcription was achieved with M-MLV Reverse Transcriptase (Promega). Real-time qPCR was achieved with qPCR SYBR Green Mix (Yeasen).

For nuclear and cytoplasmic RNA extraction (91), N2a cells were placed on ice, washed with ice-cold PBS three times, and then collected by spinning at 1,000 rpm for 10 minutes. The resulting cells were resuspended in 200 μl lysis buffer A (pH 8.0 10 mM Tris, 140 mM NaCl, 1.5 mM MgCl2, 0.5% NP-40, 1 mM DTT, 100 U/ml RNasin), incubated on ice for 5 minutes, then centrifuged at 1,000 × g for three minutes. The supernatant was collected for cytoplasmic RNA extraction. The pellets were further washed twice with the lysis buffer A and once with the lysis buffer A complemented with 1% Tween-40 and 0.5% deoxycholic acid. The resulting pellets were resuspended in TRIzol for nuclear extraction.

RNA-Seq

Total RNA was dissolved in nuclease-free water, treated with DNase (Ambion, Thermo Fisher Scientific), and quantified by Qubit RNA Assay Kit (Thermo Fisher Scientific). Agilent 2100 Bioanalyzer (Agilent Technologies) was employed to check the quality of RNA. Total RNA (3 μg) was applied for poly(A) mRNA purification by using oligo-d(T) magnetic beads (S1419S, NEB). RNA fragmentation, cDNA synthesis, terminal repair, A-tailing, and adapter ligation were performed by using RNA library prep kit (E7530L, NEB). DNA products were cleaned using AMPure XP beads (Beckman). Library quality was checked by Agilent 2100 Bioanalyzer and quantified by real-time PCR. Sequencing was performed on the Illumina HiSeq platform.

IR analysis

The quality of RNA-Seq reads was assessed by FastQC. The adaptors and low-quality reads were removed by Cutadapt to obtain clean reads. The resulting reads were aligned to mouse genome mm10 using HISAT2 (92). IR was analyzed as previously reported with slight modifications (28–30). In brief, every intron in the genome was considered as a potential retained intron, while only introns with both flanking exons covered by the aligned reads were included for further analysis. To avoid the influence of small non-coding RNAs and unknown exons located inside of the introns and low mappability regions in large introns, including high GC regions and repetitive sequence, only reads covering exon-intron junctions and exon-exon junctions were used to calculate the IRI (intron retention index). IRI for 5’ SS and 3’ SS were calculated separately. We considered the exon-intron junction reads as intronic reads, and the sum of exon-intron and exon-exon junction reads as total reads. IRI = (5’SS intronic reads / 5’SS total reads + 3’SS intronic reads / 3’SS total reads)/2. For RNA-Seq data, IRI > 0.1 and intron coverage > 0.9 were set to identify reliable IR events. For the comparison of differential intron usage, DEXSeq was employed for the statistical analysis, which offers reliable control of padj by estimation of dispersion for each counting bin with generalized linear models (93).

Heatmap, GO analysis, and conservation analysis

Heatmaps of Z-scores were plotted using Heatmap.2 of the R package “gplots”. To compare reliable DI events across different conditions, only events with IRI > 0.1 were included in the analysis. For GO enrichment analysis, the PANTHER Classification System was used to test the overrepresentation of genes (94). Statistical overrepresentation testing was performed for the GO biological process complete category and the enriched GO terms were ranked by fold- enrichment and FDR and the redundant GO terms were manually removed. To determine whether DIs are conserved between human and mouse, intron coordinate conversions were performed using UCSC LiftOver (from mm10 to hg38). The highly conserved introns (Liftover remap ratio > 90% between human and mouse) were included for the conservation comparison (p values were calculated with two-sided proportion tests).

Nanopore full-length mRNA sequencing

Mouse cerebellar total RNA was extracted using TRIzol and treated with DNase. RNA quality was assessed using an Agilent 2100 Bioanalyzer to ensure the integrity of RNA. Total RNA (1 μg) was used for cDNA library construction, following the protocols suggested by Oxford Nanopore Technologies (ONT). Reverse transcription and strand-switching were performed using a cDNA-PCR Sequencing Kit (SQK-PCS109, ONT). cDNA was PCR amplified for 14 cycles by using LongAmp Taq (NEB). ONT adaptor ligation was performed using T4 DNA ligase (NEB). The resulting DNA was purified by Agencourt XP beads. The final libraries were added to FLO-MIN109 flowcells (ONT), sequenced using PromethION platform. Raw ONT reads were filtered by read quality score (> 7) and minimal reads length (> 500bps). Clean ONT reads were aligned to mouse genome reference mm10. Aligned reads were converted to bam files, sorted, and indexed by SAMtools (95). Reliable full-length transcripts were filtered to contain 5’UTR and 3’UTR regions by using BEDTools (96). To identify reliable DIs, minimal IRI was set as 0.05 with intronic region coverage larger than 0.9.

Native RIP-seq

Native RIP-seq was performed as previously reported with minor modifications (97). For mouse tissues, mouse cerebellum was removed and placed in a Dounce homogenizer on ice. Lysis was in RIP buffer (150 mM KCl, 25 mM Tris pH 7.4, 5 mM EDTA, 0.5 mM DTT, 0.5% Nonidet P-40) complemented with protease and phosphatase inhibitor cocktail (Roche) and 100 U/ml RNasin (Promega). For cultured cells, cells were washed with ice-cold PBS three times and lysed in RIP buffer on ice. Tissue and cell lysates went through a 23-gauge needle, rotated at 4℃ for 0.5 hour, and centrifuged to remove cell debris. The resulting supernatant was incubated with anti-FLAG M2 magnetic beads (M8823, Merck). After incubation for 12 hours at 4℃, M2 beads were washed with RIP buffer twice and TBS buffer twice. Elution was achieved by using 250 μg/ml 3 × Flag peptides (F4799, Sigma). The resulting eluates were resuspended in TRIzol for the following RNA extraction. Smart-seq2 was performed following a standard protocol as previously described (98). The amplified DNA (40 ng) was fragmented into ∼350 bp fragments using a Bioruptor® Sonication System (Diagenode Inc.). Illumina library construction was performed and qualified libraries were sequenced on the Illumina HiSeq platform. As described above, we used exon-intron junction and exon-exon junction reads to calculate IRI. For ratio comparison of IRI, IRI > 0.1 and intron coverage > 0.8 were set to identify reliable IR events.

CLIP analysis

The RNPS1-GFP iCLIP and SF3a60 eCLIP were performed by Hauer et al. and Van Nostrand et al., previously (52–54), and the raw data were downloaded from ArrayExpress and ENCODE (99–101). The CLIP and corresponding RNA- Seq reads were aligned to the human reference genome hg19 by STAR v2.7 (102). The second read (R2) in each eCLIP read pair were extracted by SAMtools for downstream analysis, as described in the eCLIP-seq processing pipeline (103). To generate a CLIP read density plot, biological replicates were merged for combined reads count. Introns were grouped based on IR analysis in CLIP-related RNA-Seq data. DIs with high confidence were filtered by IRI > 0.1 and intron region coverage > 0.95, while efficiently spliced introns were filtered by IRI < 0.05. CLIP read coverage of introns and intron flanking exons were calculated using BEDTools (96), then read coverage were summed and normalized according to their relative position to 5’ SS and 3’ SS. The final read density was plotted using R (version 3.4.3). CLIP peak calling was achieved with CLIPper (CLIP peak enrichment recognition) by the default parameters (104). Binding stringency was ranked by the p-value calculated by CLIPper.

AUTHOR CONTRIBUTIONS

Y.J. and D.M. conceived the study and wrote the manuscript. D.M. performed the mouse forward genetic screening and identified the genetic modifier. L.L. performed the CRISPR-Cas9-mediated Snip1 KO and Snip1-Flag KI in mice. D.M. performed the IR analysis and analyzed the RNA-Seq, nanopore sequencing, and CLIP data. X.Z. performed the immunostaining. D.M. performed the MS and RIP-seq and analyzed the data. D.M. and Q.Z. validated the RNA-Seq and MS results. D.M. and Q.Z. mapped the SNIP1/RNPS1 interaction regions.

Supplemental information

Figure S1. An ENU-induced mutagenesis screening for dominant modifier(s) that rescue(s) the NMF291^-/- phenotypes.

G0 male NMF291^-/+ mice were injected with ENU (N-ethyl-N-nitrosourea) once a week for 3 weeks. After their fertilities recovered, they were bred to ENU- untreated female NMF291^-/+ mice. The G1 NMF291^-/- mice carrying less ataxia phenotype and improved lifespan were crossed to ENU-untreated NMF291^-/+ mice to determine the family pedigree.

View this table:

Table S1. Modifier candidates for the NMF291 phenotypes.

View this table:

Table S2. The abnormal Mendelian ratio seen in the progenies from Snip1^M+ and Snip1^-/+ intercrosses.

Figure S2. Haploinsufficiency of Snip1 partially rescues the IRs and their corresponding gene expressions.

(A-C) Validation of the IRs rescued by Snip1^-/+ by RT-PCR. Cerebella were harvested at one month of age with the indicated genotypes. Unspl., unspliced transcripts; spl., spliced transcripts.

(D) The representative IRs visualized by IGV. Primers used for the RT-PCR validation in (C) were illustrated. F, forward primer; R, reverse primer.

(E) Corresponding genes (1455) of the rescued IRs (1961) shown in Figure 2C were included for expression analysis. The Z-score was used to normalize expression level in each row.

Figure S3. Intron-containing transcripts overrepresented in NMF291 mutant cerebellum are likely IDTs.

(A) The DIs amplified by Mu-U2 but not WT-U2 in N2a cells.

(B) Previously described a NMD reporter (PMID: 1693475) in which Renilla luciferase contains a PTC in the second last exon. Application of CHX significantly inhibited NMD measured by relative Renilla activity. The Firefly luciferase was used to normalize transfection efficiency.

(C) The intron-containing transcripts were insensitive to application of CHX in culture medium of N2a cells expressing WT-U2 or Mu-U2.

(D and E) N2a cells were infected with Upf1 and scrambled shRNA (MissionRNAi, Sigma). The relative expression level of Upf1 measured by quantitative-PCR (D). The intron-containing transcripts were detected by RT- PCR (E).

(F) The nuclear and cytosolic fractions evidenced by the enrichments of U99 and 12s rRNA, respectively. Nu., nucleus; Cyt., cytosol.

In B and D, the values are presented as mean ± SEM, *p<0.05, **p<0.01, n = 4, t-test, SPSS. In A, C, E, and F, unspl., unspliced transcripts; spl., spliced transcripts.

Figure S4. DIs are likely not code proteins.

(A) Detained intron 5 of Ptbp1 was evidenced by both RNA-seq (upper) and RT-PCR (lower) in wildtype (+/+) and NMF291 mutant (-/-) cerebella.

(B and C) The expression of PTBP1 by immunoblot. GAPDH served as loading control. Asterisk marked the corresponding molecular weight of PTBP1 supposedly coded by the intron 5-containing transcript.

(D) A pipeline combines RNA-seq and MS (mass spectrometry) for seeking intron-coded peptide. Based on the DIs we identified in the NMF291^-/- cerebellum, we generated customized peptide database, which are coded by the DIs and their upstream exons.

(E) Protein lysates from +/+ (n = 3) and NMF291^-/- mutant (n = 3) cerebella were applied for MS. The MS hits of our customized peptide database from one +/+ and mutant pair were pooled and summarized as one smaple. Mouse, one month of age.

In A, unspl., unspliced transcripts; spl., spliced transcripts. In C, the values are presented as mean ± SEM. NS, no statistical significance (n = 3, t-test, SPSS).

Figure S5. IDTs overrepresented in the NMF291 mutant mouse are also accumulated in aged cerebellum, revealed by nanopore sequencing.

(A) The majority of DIs accumulated in aged wildtype cerebellum are overlapped with that of the P30 NMF291^-/- cerebellum.

(B) Percentage of IDTs in young (P30), aged (2.5 years), and P30 NMF291^-/- cerebella. All full-length nanopore reads containing 5’ UTR and 3’ UTR were included for the calculation. p values correspond to two-sided proportion tests.

Figure S6. Generation of Snip1-Flag mouse.

(A) Crispr/Cas9-based Snip1-Flag knockin (KI) design. The mouse Snip1 locus and Snip1 last coding exon (exon 4, Ex4) are shown (Upper). The stop codon (TAA) labeled with an asterisk (*); the target sequences of gRNA and PAM site (NGG) labeled in red and blue, respectively. In the donor DNA, a 3- time Flag tag was fused with Snip1 last coding exon. A linker placed between Ex4 and the Flag sequences, which codes 2-time G4S (G4S)₂.

(B) Genomic DNA PCR confirmed the right KI. Primers for the PCR were labeled in (A) by arrows. F, forward; R, reverse. f/+, mouse heterozygous for Snip1-Flag KI.

(D) Flag-IP was performed with cerebellar protein lysates from Snip1-Flag KI mouse.

Figure S7. SNIP1-interacting partners are protein components found in B^act but not B*.

(A-F) Interactions between SNIP1 and protein components found in B^act (SF3a66, SF3a60, SF3b130, and Syf1) but not those in B* (CWC25 and EIF4A3) were validated in N2a cells expressing the indicated tagged proteins.

Figure S8. SNIP1 and RNPS1 function as a molecular brake to pause spliceosome at DIs.

(A) Knockdown of Snip1 in N2a cells by lentiviral shRNA (shSnip1). Note the target sequence of shSnip1 located in the 3’UTR (upper). The relative expression level of Snip1 measured by quantitative-PCR (lower). The scrambled shRNA served as a control.

(B) Knockdown of Snip1 reduced DIs amplified by Mu-U2. The shRNA infected N2a cells were transfected with WT- and Mu-U2 expression plasmids, respectively. Scrambled shRNA infection served as a control.

(C) Knockdown of genes encoding peripheral EJC components (Acin1, Pnn, and Rnps1) and a spicing factor (SRm300) measured by quantitative-PCR.

(D) The interactions of SNIP1, PNN, and RNPS1 were evidenced by co-IP in N2a cells expressing PNN-HA, Flag-SNIP1, and RNPS1-Myc, simultaneously.

(E) The shSnip1 targets 3’-UTR of endogenous Snip1, which does not affect the expression of exogenous full-length and ΔFHA SNIP1 in N2a cells.

(F) Flag-RIP was performed with protein lysates from N2a cells infected with Flag-RNPS1 and transfected with Mu-U2 expression plasmid.

In A and C, the values are presented as mean ± SEM, n = 4. *p<0.05, **p<0.01, ***p<0.001, N.S., no statistical significance, t-test or ANOVA, SPSS. In B and F, DIs were measured by RT-PCR. Unspl., unspliced transcripts; spl., spliced transcripts.

Figure S9. Generation of Snip1 cKO mouse.

(A) Crispr/Cas9-based Snip1 cKO design. Genomic structure of mouse Snip1 is illustrated. Homology arms (Left and Right) in donor DNA were labeled. Four gRNAs (2 gRNAs each side) flanking the Snip1 exon3 (Ex3) were employed to increase homologous recombination. Primers for genotyping and detecting Cre-induced Ex3 deletion were labeled. Primers (R1 and F2) covering LoxP sites were labeled in red. F1 and R2 are located outside the homology arms.

(B) Genomic PCR detected right LoxP site insertions in mice heterozygous for the floxed allele (+/fl). Primers used were labeled in (A).

(C) Confirmation of the NseCreER expression by a previously described Cre- reporter, H2B mCherry (PMID: 25913859). The NseCreER were predominantly expressed in cerebellar granule cells and a few expressed in the granule neurons in hippocampal dentate gyrus.

(D) Cre-dependent Ex3 deletion was detected at both genomic DNA (gDNA) and RNA levels in cerebellum and hippocampus but not in cortex in Snip1^fl^/^fl;NseCreER mouse. Snip1^fl^/^fl mouse served as negative control. Tamoxifen injections on P3, 4, and 5; DNA/RNA harvest on P7.

Figure S10. Our working model for post-transcriptional spliceosome pausing at highly regulated DIs and its contribution to neurodegeneration.

We suggest that DI splicing is paused at B^act state, an active spliceosome but not catalytically primed, and SNIP1 and RNPS1 function as a molecular brake for the spliceosome pausing. The whole process can be divided into several steps. 1) Spliceosome loaded at both constitutive and detained introns. After constitutive introns are spliced, pre-mRNAs are polyadenylated to complete the co-transcriptional splicing. 2) The splicing of DIs is paused at B^act state (spliceosome pausing), which is mediated by SNIP1/RNPS1 containing complex. SNIP1, RNPS1 and B^act component preferentially dock at DIs, forming a molecular brake to modulate spliceosome pausing. RNPS1 recognizes the DIs and its neighboring sequences through its RRM (RNA recognition motif). Interaction between SNIP1 and RNPS1 is mediated by SNIP1 FHA (forkhead-associated) domain and RNPS1 S (serine-rich) domain. 3) The paused spliceosome is resumed, and DIs are spliced. The completely spliced transcripts are exported to cytosol for protein translation. Mutant (the NMF291^-/-) or dysfunctional (Snip1^-/-) spliceosome decreases splicing efficiency of DIs but has little effect on that of constitutively-spliced introns. That leads to accumulation of DIs, which in turn worsens DI splicing by sequester of spliceosome pausing complex and depletion of the available complex components and further damages the corresponding gene functions, and eventually causes neurodegeneration. Partial loss of SNIP1 or RNPS1 rescues intron detentions caused by expression of mutant U2, probably through releasing the molecular brake.

ACKNOWLEDGMENTS

We thank Dr. Susan L. Ackerman for the NMF291 mouse line. We thank the staff members in the Tsinghua Laboratory Animal Research Center (LARC) for mouse housing. We thank Dr. Haiteng Deng, Dr. Yi Huo, and Chongchong Zhao in the Tsinghua Proteomics Resource Center for mass spectrometry. We thank Dr. Douglas Black for the PTB-NT antibody and Drs. Gabriele Neu-Yilik and Andreas E. Kulozik for the NMD reporters. We also thank Dr. Michael Q. Zhang and Dr. Susan L. Ackerman for their comments on the manuscript. This work was supported by the National Natural Science Foundation of China (NSFC grants 31571097 and 81371361), the Thousand Talent Program for Young Outstanding Scientists, the Tsinghua-Peking Joint Center for Life Sciences, and IDG/McGovern Institute for Brain Research at Tsinghua.

REFERENCES

1.↵
R. A. Padgett, P. J. Grabowski, M. M. Konarska, S. Seiler, P. A. Sharp, Splicing of messenger RNA precursors. Annual review of biochemistry 55, 1119–1150 (1986).
OpenUrl CrossRef PubMed Web of Science
2.↵
M. C. Wahl, C. L. Will, R. Luhrmann, The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 (2009).
OpenUrl CrossRef PubMed Web of Science
3.
M. E. Wilkinson, C. Charenton, K. Nagai, RNA Splicing by the Spliceosome. Annual review of biochemistry 89, (2020).
4.↵
Y. Shi, Mechanistic insights into precursor messenger RNA splicing by the spliceosome. Nature reviews Molecular cell biology 18, 655 (2017).
5.↵
R. M. Lardelli, J. X. Thompson, J. R. Yates, 3rd., S. W. Stevens, Release of SF3 from the intron branchpoint activates the first step of pre-mRNA splicing. RNA 16, 516–528 (2010).
OpenUrl Abstract/FREE Full Text
6.↵
T. Ohrt et al., Prp2-mediated protein rearrangements at the catalytic core of the spliceosome as revealed by dcFCCS. RNA 18, 1244–1256 (2012).
OpenUrl Abstract/FREE Full Text
7.↵
X. Zhang et al., Structure of the human activated spliceosome in three conformational states. Cell Res 28, 307–322 (2018).
OpenUrl CrossRef PubMed
8.↵
S. Bessonov, M. Anokhina, C. L. Will, H. Urlaub, R. Lührmann, Isolation of an active step I spliceosome and composition of its RNP core. Nature 452, 846–850 (2008).
OpenUrl CrossRef PubMed Web of Science
9.↵
E. T. Wang et al., Alternative isoform regulation in human tissue transcriptomes. Nature 456, 470–476 (2008).
OpenUrl CrossRef PubMed Web of Science
10.↵
Q. Pan, O. Shai, L. J. Lee, B. J. Frey, B. J. Blencowe, Deep surveying of alternative splicing complexity in the human transcriptome by high-throughput sequencing. Nature genetics 40, 1413–1415 (2008).
OpenUrl CrossRef PubMed Web of Science
11.↵
T. W. Nilsen, B. R. Graveley, Expansion of the eukaryotic proteome by alternative splicing. Nature 463, 457–463 (2010).
OpenUrl CrossRef PubMed Web of Science
12.↵
H. Keren, G. Lev-Maor, G. Ast, Alternative splicing and evolution: diversification, exon definition and function. Nat Rev Genet 11, 345–355 (2010).
OpenUrl CrossRef PubMed Web of Science
13.↵
M. W. Popp, L. E. Maquat, Organizing principles of mammalian nonsense-mediated mRNA decay. Annu Rev Genet 47, 139–165 (2013).
OpenUrl CrossRef PubMed Web of Science
14.↵
A. G. Jacob, C. W. J. Smith, Intron retention as a component of regulated gene expression programs. Hum Genet 136, 1043–1057 (2017).
OpenUrl CrossRef
15.↵
P. L. Boutz, A. Bhutkar, P. A. Sharp, Detained introns are a novel, widespread class of post-transcriptionally spliced introns. Genes Dev 29, 63–80 (2015).
OpenUrl Abstract/FREE Full Text
16.↵
K. Yap, Z. Q. Lim, P. Khandelia, B. Friedman, E. V. Makeyev, Coordinated regulation of neuronal mRNA steady-state levels through developmentally controlled intron retention. Genes Dev 26, 1209–1223 (2012).
OpenUrl Abstract/FREE Full Text
17.
O. Mauger, F. Lemoine, P. Scheiffele, Targeted Intron Retention and Excision for Rapid Gene Regulation in Response to Neuronal Activity. Neuron 92, 1266–1278 (2016).
OpenUrl CrossRef
18.
J. Gill et al., Regulated intron removal integrates motivational state and experience. Cell 169, 836–848. e815 (2017).
OpenUrl CrossRef PubMed
19.
C. Naro et al., An Orchestrated Intron Retention Program in Meiosis Controls Timely Usage of Transcripts during Germ Cell Differentiation. Dev Cell 41, 82–93 e84 (2017).
OpenUrl CrossRef
20.↵
K. Ninomiya, N. Kataoka, M. Hagiwara, Stress-responsive maturation of Clk1/4 pre- mRNAs promotes phosphorylation of SR splicing factor. J Cell Biol 195, 27–40 (2011).
OpenUrl Abstract/FREE Full Text
21.
K. E. Pendleton et al., The U6 snRNA m(6)A Methyltransferase METTL16 Regulates SAM Synthetase Intron Retention. Cell 169, 824–835 e814 (2017).
OpenUrl CrossRef PubMed
22.
S. K. Park et al., A Conserved Splicing Silencer Dynamically Regulates O-GlcNAc Transferase Intron Retention and O-GlcNAc Homeostasis. Cell Rep 20, 1088–1099 (2017).
OpenUrl CrossRef PubMed
23.↵
Z. W. Tan et al., O-GlcNAc regulates gene expression by controlling detained intron splicing. Nucleic Acids Res 48, 5656–5669 (2020).
OpenUrl CrossRef
24.↵
E. S. Lander et al., Initial sequencing and analysis of the human genome. Nature 409, 860–921 (2001).
OpenUrl CrossRef PubMed Web of Science
25.
C. N. Tennyson, H. J. Klamut, R. G. Worton, The human dystrophin gene requires 16 hours to be transcribed and is cotranscriptionally spliced. Nat Genet 9, 184–190 (1995).
OpenUrl CrossRef PubMed Web of Science
26.
X. Darzacq et al., In vivo dynamics of RNA polymerase II transcription. Nat Struct Mol Biol 14, 796–806 (2007).
OpenUrl CrossRef PubMed Web of Science
27.↵
J. Singh, R. A. Padgett, Rates of in situ transcription and splicing in large human genes. Nat Struct Mol Biol 16, 1128–1133 (2009).
OpenUrl CrossRef PubMed Web of Science
28.↵
Y. Jia, J. C. Mu, S. L. Ackerman, Mutation of a U2 snRNA gene causes global disruption of alternative splicing and neurodegeneration. Cell 148, 296–308 (2012).
OpenUrl CrossRef PubMed Web of Science
29.↵
J. J. Wong et al., Orchestrated intron retention regulates normal granulocyte differentiation. Cell 154, 583–595 (2013).
OpenUrl CrossRef PubMed Web of Science
30.↵
U. Braunschweig et al., Widespread intron retention in mammals functionally tunes transcriptomes. Genome Res 24, 1774–1786 (2014).
OpenUrl Abstract/FREE Full Text
31.↵
L. E. Maquat, Nonsense-mediated mRNA decay: splicing, translation and mRNP dynamics. Nat Rev Mol Cell Biol 5, 89–99 (2004).
OpenUrl CrossRef PubMed Web of Science
32.↵
B. M. Venkatesan, R. Bashir, Nanopore sensors for nucleic acid analysis. Nat Nanotechnol 6, 615–624 (2011).
OpenUrl CrossRef PubMed
33.↵
R. H. Kim et al., SNIP1 inhibits NF-κB signaling by competing for its binding to the C/H1 domain of CBP/p300 transcriptional co-activators. Journal of Biological Chemistry 276, 46297–46304 (2001).
OpenUrl Abstract/FREE Full Text
34.↵
R. H. Kim et al., A novel smad nuclear interacting protein, SNIP1, suppresses p300- dependent TGF-β signal transduction. Genes & development 14, 1605–1616 (2000).
OpenUrl Abstract/FREE Full Text
35.↵
C. P. Bracken et al., Regulation of cyclin D1 RNA stability by SNIP1. Cancer Res 68, 7621–7628 (2008).
OpenUrl Abstract/FREE Full Text
36.↵
S. Picelli et al., Smart-seq2 for sensitive full-length transcriptome profiling in single cells. Nat Methods 10, 1096–1098 (2013).
OpenUrl CrossRef PubMed Web of Science
37.↵
D. Haselbach et al., Structure and Conformational Dynamics of the Human Spliceosomal B(act) Complex. Cell 172, 454–464 e411 (2018).
OpenUrl CrossRef PubMed
38.↵
R. Wan, R. Bai, C. Yan, J. Lei, Y. Shi, Structures of the Catalytically Activated Yeast Spliceosome Reveal the Mechanism of Branching. Cell 177, 339–351 e313 (2019).
OpenUrl CrossRef PubMed
39.↵
C. Townsend et al., Mechanism of protein-guided folding of the active site U2/U6 RNA during spliceosome activation. Science 370, (2020).
40.
K. Bertram et al., Cryo-EM Structure of a Pre-catalytic Human Spliceosome Primed for Activation. Cell 170, 701–713.e711 (2017).
OpenUrl CrossRef PubMed
41.↵
X. Zhan, C. Yan, X. Zhang, J. Lei, Y. Shi, Structure of a human catalytic step I spliceosome. Science 359, 537–545 (2018).
OpenUrl Abstract/FREE Full Text
42.↵
V. L. Reichert, H. Le Hir, M. S. Jurica, M. J. Moore, 5’ exon interactions within the human spliceosome establish a framework for exon junction complex structure and assembly. Genes Dev 16, 2778–2791 (2002).
OpenUrl Abstract/FREE Full Text
43.↵
D. Durocher, S. P. Jackson, The FHA domain. FEBS Lett 513, 58–66 (2002).
OpenUrl CrossRef PubMed Web of Science
44.↵
P. Wysoczanski et al., Cooperative structure of the heterotrimeric pre-mRNA retention and splicing complex. Nat Struct Mol Biol 21, 911–918 (2014).
OpenUrl CrossRef PubMed
45.↵
A. G. Murachelli, J. Ebert, C. Basquin, H. Le Hir, E. Conti, The structure of the ASAP core complex reveals the existence of a Pinin-containing PSAP complex. Nat Struct Mol Biol 19, 378–386 (2012).
OpenUrl CrossRef PubMed
46.↵
V. Boehm et al., Exon Junction Complexes Suppress Spurious Splice Sites to Safeguard Transcriptome Integrity. Mol Cell 72, 482–495 e487 (2018).
OpenUrl CrossRef PubMed
47.↵
H. Le Hir, J. Sauliere, Z. Wang, The exon junction complex as a node of post- transcriptional networks. Nat Rev Mol Cell Biol 17, 41–54 (2016).
OpenUrl CrossRef PubMed
48.
R. Hayashi, D. Handler, D. Ish-Horowicz, J. Brennecke, The exon junction complex is required for definition and excision of neighboring introns in Drosophila. Genes Dev 28, 1772–1785 (2014).
OpenUrl Abstract/FREE Full Text
49.
C. D. Malone et al., The exon junction complex controls transposable element activity by ensuring faithful splicing of the piwi transcript. Genes Dev 28, 1786–1799 (2014).
OpenUrl Abstract/FREE Full Text
50.
L. Blazquez et al., Exon Junction Complex Shapes the Transcriptome by Repressing Recursive Splicing. Mol Cell 72, 496–509 e499 (2018).
OpenUrl CrossRef
51.↵
T. Gonatopoulos-Pournatzis et al., Genome-wide CRISPR-Cas9 Interrogation of Splicing Networks Reveals a Mechanism for Recognition of Autism-Misregulated Neuronal Microexons. Mol Cell 72, 510–524 e512 (2018).
OpenUrl CrossRef PubMed
52.↵
E. L. Van Nostrand et al., Principles of RNA processing from analysis of enhanced CLIP maps for 150 RNA binding proteins. Genome Biol 21, 90 (2020).
53.
E. L. Van Nostrand et al., A large-scale binding and functional map of human RNA- binding proteins. Nature 583, 711–719 (2020).
OpenUrl CrossRef PubMed
54.↵
C. Hauer et al., Exon Junction Complexes Show a Distributional Bias toward Alternatively Spliced mRNAs and against mRNAs Coding for Ribosomal Proteins. Cell Rep 16, 1588–1603 (2016).
OpenUrl CrossRef
55.↵
T. Pohlkamp et al., Generation and characterization of an Nse-CreERT2 transgenic line suitable for inducible gene manipulation in cerebellar granule cells. PLoS One 9, e100384 (2014).
OpenUrl
56.↵
S. P. Peron, J. Freeman, V. Iyer, C. Guo, K. Svoboda, A Cellular Resolution Map of Barrel Cortex Activity during Tactile Behavior. Neuron 86, 783–799 (2015).
OpenUrl CrossRef PubMed
57.↵
E. Aparicio-Prat et al., DECKO: Single-oligo, dual-CRISPR deletion of genomic elements including long non-coding RNAs. BMC Genomics 16, 846 (2015).
OpenUrl CrossRef PubMed
58.↵
A. Dziembowski et al., Proteomic analysis identifies a new complex required for nuclear pre-mRNA retention and splicing. EMBO J 23, 4847–4856 (2004).
OpenUrl Abstract/FREE Full Text
59.↵
P. Fabrizio et al., The evolutionarily conserved core design of the catalytic activation step of the yeast spliceosome. Mol Cell 36, 593–608 (2009).
OpenUrl CrossRef PubMed Web of Science
60.
C. Yan, R. Wan, R. Bai, G. Huang, Y. Shi, Structure of a yeast activated spliceosome at 3.5 å resolution. Science 353, 904–911 (2016).
OpenUrl Abstract/FREE Full Text
61.↵
C. Schneider et al., Dynamic Contacts of U2, RES, Cwc25, Prp8 and Prp45 Proteins with the Pre-mRNA Branch-Site and 3’ Splice Site during Catalytic Activation and Step 1 Catalysis in Yeast Spliceosomes. PLoS Genet 11, e1005539 (2015).
OpenUrl CrossRef PubMed
62.↵
A. Gottschalk, C. Bartels, G. Neubauer, R. Luhrmann, P. Fabrizio, A novel yeast U2 snRNP protein, Snu17p, is required for the first catalytic step of splicing and for progression of spliceosome assembly. Mol Cell Biol 21, 3037–3046 (2001).
OpenUrl Abstract/FREE Full Text
63.↵
J. P. Fernandez et al., RES complex is associated with intron definition and required for zebrafish early embryogenesis. PLoS Genet 14, e1007473 (2018).
OpenUrl CrossRef
64.↵
L. Frankiw et al., BUD13 Promotes a Type I Interferon Response by Countering Intron Retention in Irf7. Mol Cell 73, 803–814 e806 (2019).
OpenUrl CrossRef
65.↵
R. F. Luco et al., Regulation of alternative splicing by histone modifications. Science 327, 996–1000 (2010).
OpenUrl Abstract/FREE Full Text
66.↵
Z. Xing et al., lncRNA directs cooperative epigenetic regulation downstream of chemokine signals. Cell 159, 1110–1125 (2014).
OpenUrl CrossRef PubMed
67.↵
J. Lykke-Andersen, M. D. Shu, J. A. Steitz, Communication of the position of exon-exon junctions to the mRNA surveillance machinery by the protein RNPS1. Science 293, 1836–1839 (2001).
OpenUrl Abstract/FREE Full Text
68.
H. Le Hir, E. Izaurralde, L. E. Maquat, M. J. Moore, The spliceosome deposits multiple proteins 20-24 nucleotides upstream of mRNA exon-exon junctions. EMBO J 19, 6860–6869 (2000).
OpenUrl Abstract/FREE Full Text
69.↵
N. H. Gehring et al., Exon-junction complex components specify distinct routes of nonsense-mediated mRNA decay with differential cofactor requirements. Mol Cell 20, 65–75 (2005).
OpenUrl CrossRef PubMed Web of Science
70.↵
C. Schwerk et al., ASAP, a novel protein complex involved in RNA processing and apoptosis. Molecular and Cellular Biology 23, 2981–2990 (2003).
OpenUrl Abstract/FREE Full Text
71.↵
T. O. Tange, T. Shibuya, M. S. Jurica, M. J. Moore, Biochemical analysis of the EJC reveals two new factors and a stable tetrameric protein core. RNA 11, 1869–1883 (2005).
OpenUrl Abstract/FREE Full Text
72.↵
C. Girard et al., Post-transcriptional spliceosomes are retained in nuclear speckles until splicing completion. Nat Commun 3, 994 (2012).
73.
C. Li, R. I. Lin, M. C. Lai, P. Ouyang, W. Y. Tarn, Nuclear Pnn/DRS protein binds to spliced mRNPs and participates in mRNA processing and export via interaction with RNPS1. Mol Cell Biol 23, 7363–7376 (2003).
OpenUrl Abstract/FREE Full Text
74.
A. Mayeda et al., Purification and characterization of human RNPS1: a general activator of pre-mRNA splicing. EMBO J 18, 4560–4570 (1999).
OpenUrl Abstract/FREE Full Text
75.↵
E. Sakashita, S. Tatsumi, D. Werner, H. Endo, A. Mayeda, Human RNPS1 and its associated factors: a versatile alternative pre-mRNA splicing regulator in vivo. Mol Cell Biol 24, 1174–1187 (2004).
OpenUrl Abstract/FREE Full Text
76.↵
J. H. Trembley et al., Activation of pre-mRNA splicing by human RNPS1 is regulated by CK2 phosphorylation. Mol Cell Biol 25, 1446–1457 (2005).
OpenUrl Abstract/FREE Full Text
77.↵
P. Bao, C. L. Will, H. Urlaub, K. L. Boon, R. Luhrmann, The RES complex is required for efficient transformation of the precatalytic B spliceosome into an activated B(act) complex. Genes Dev 31, 2416–2429 (2017).
OpenUrl Abstract/FREE Full Text
78.↵
S. Ullrich, R. Guigo, Dynamic changes in intron retention are tightly associated with regulation of splicing factors and proliferative activity during B-cell development. Nucleic Acids Res 48, 1327–1340 (2020).
OpenUrl
79.↵
S. Adusumalli, Z. K. Ngian, W. Q. Lin, T. Benoukraf, C. T. Ong, Increased intron retention is a post-transcriptional signature associated with progressive aging and Alzheimer’s disease. Aging Cell 18, e12928 (2019).
80.↵
Q. Wang, E. G. Conlon, J. L. Manley, D. C. Rio, Widespread intron retention impairs protein homeostasis in C9orf72 ALS brains. Genome Res 30, 1705–1715 (2020).
OpenUrl Abstract/FREE Full Text
81.↵
B. Lizen, M. Claus, L. Jeannotte, F. M. Rijli, F. Gofflot, Perinatal induction of Cre recombination with tamoxifen. Transgenic Res 24, 1065–1077 (2015).
OpenUrl CrossRef
82.↵
M. E. Pitulescu, I. Schmidt, R. Benedito, R. H. Adams, Inducible gene targeting in the neonatal vasculature and analysis of retinal angiogenesis in mice. Nat Protoc 5, 1518–1534 (2010).
OpenUrl CrossRef PubMed
83.↵
A. P. Salinger, M. J. Justice, Mouse mutagenesis using N-ethyl-N-nitrosourea (ENU). Cold Spring Harbor Protocols 2008, pdb. prot4985 (2008).
84.↵
R.-Z. Chen et al., An ENU-induced mutation in Twist1 transactivation domain causes hindlimb polydactyly with complete penetrance and dominant-negatively impairs E2A-dependent transcription. Scientific reports 10, 1–12 (2020).
OpenUrl
85.↵
A. McKenna et al., The Genome Analysis Toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data. Genome research 20, 1297–1303 (2010).
OpenUrl Abstract/FREE Full Text
86.↵
K. Wang, M. Li, H. Hakonarson, ANNOVAR: functional annotation of genetic variants from high-throughput sequencing data. Nucleic acids research 38, e164–e164 (2010).
OpenUrl CrossRef PubMed
87.↵
P. D. Hsu et al., DNA targeting specificity of RNA-guided Cas9 nucleases. Nature biotechnology 31, 827–832 (2013).
OpenUrl CrossRef PubMed
88.↵
F. A. Ran et al., Genome engineering using the CRISPR-Cas9 system. Nature protocols 8, 2281–2308 (2013).
OpenUrl
89.↵
O. Shalem et al., Genome-scale CRISPR-Cas9 knockout screening in human cells. Science 343, 84–87 (2014).
OpenUrl Abstract/FREE Full Text
90.↵
S. Boelz, G. Neu-Yilik, N. H. Gehring, M. W. Hentze, A. E. Kulozik, A chemiluminescence- based reporter system to monitor nonsense-mediated mRNA decay. Biochemical and biophysical research communications 349, 186–191 (2006).
OpenUrl CrossRef PubMed Web of Science
91.↵
H. W. Hwang, E. A. Wentzel, J. T. Mendell, A hexanucleotide element directs microRNA nuclear import. Science 315, 97–100 (2007).
OpenUrl Abstract/FREE Full Text
92.↵
D. Kim, J. M. Paggi, C. Park, C. Bennett, S. L. Salzberg, Graph-based genome alignment and genotyping with HISAT2 and HISAT-genotype. Nature biotechnology 37, 907–915 (2019).
OpenUrl CrossRef PubMed
93.↵
S. Anders, A. Reyes, W. Huber, Detecting differential usage of exons from RNA-seq data. Genome Res 22, 2008–2017 (2012).
OpenUrl Abstract/FREE Full Text
94.↵
H. Mi, A. Muruganujan, J. T. Casagrande, P. D. Thomas, Large-scale gene function analysis with the PANTHER classification system. Nat Protoc 8, 1551–1566 (2013).
OpenUrl CrossRef PubMed
95.↵
H. Li et al., The sequence alignment/map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).
OpenUrl CrossRef PubMed Web of Science
96.↵
A. R. Quinlan, BEDTools: The Swiss-Army Tool for Genome Feature Analysis. Curr Protoc Bioinformatics 47, 11 12 11–34 (2014).
OpenUrl CrossRef
97.↵
J. L. Rinn et al., Functional demarcation of active and silent chromatin domains in human HOX loci by noncoding RNAs. cell 129, 1311–1323 (2007).
OpenUrl CrossRef PubMed Web of Science
98.↵
S. Picelli et al., Full-length RNA-seq from single cells using Smart-seq2. Nature protocols 9, 171–181 (2014).
OpenUrl
99.↵
E. P. Consortium, An integrated encyclopedia of DNA elements in the human genome. Nature 489, 57 (2012).
100.
J. Zhang et al., An integrative ENCODE resource for cancer genomics. Nature communications 11, 1–11 (2020).
OpenUrl
101.↵
A. Athar, et al., ArrayExpress update - from bulk to single-cell expression data. Nucleic Acids Res 47, D711–D715 (2019).
OpenUrl CrossRef PubMed
102.↵
A. Dobin et al., STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
OpenUrl CrossRef PubMed Web of Science
103.↵
E. L. Van Nostrand et al., Robust transcriptome-wide discovery of RNA-binding protein binding sites with enhanced CLIP (eCLIP). Nat Methods 13, 508–514 (2016).
OpenUrl CrossRef PubMed
104.↵
M. T. Lovci et al., Rbfox proteins regulate alternative mRNA splicing through evolutionarily conserved RNA bridges. Nature structural & molecular biology 20, 1434–1442 (2013).
OpenUrl

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[1] 1.↵
R. A. Padgett, P. J. Grabowski, M. M. Konarska, S. Seiler, P. A. Sharp, Splicing of messenger RNA precursors. Annual review of biochemistry 55, 1119–1150 (1986).
OpenUrl CrossRef PubMed Web of Science

[2] 2.↵
M. C. Wahl, C. L. Will, R. Luhrmann, The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 (2009).
OpenUrl CrossRef PubMed Web of Science

[3] 3.
M. E. Wilkinson, C. Charenton, K. Nagai, RNA Splicing by the Spliceosome. Annual review of biochemistry 89, (2020).

[4] 4.↵
Y. Shi, Mechanistic insights into precursor messenger RNA splicing by the spliceosome. Nature reviews Molecular cell biology 18, 655 (2017).

[5] 5.↵
R. M. Lardelli, J. X. Thompson, J. R. Yates, 3rd., S. W. Stevens, Release of SF3 from the intron branchpoint activates the first step of pre-mRNA splicing. RNA 16, 516–528 (2010).
OpenUrl Abstract/FREE Full Text

[6] 6.↵
T. Ohrt et al., Prp2-mediated protein rearrangements at the catalytic core of the spliceosome as revealed by dcFCCS. RNA 18, 1244–1256 (2012).
OpenUrl Abstract/FREE Full Text

[7] 7.↵
X. Zhang et al., Structure of the human activated spliceosome in three conformational states. Cell Res 28, 307–322 (2018).
OpenUrl CrossRef PubMed

[8] 8.↵
S. Bessonov, M. Anokhina, C. L. Will, H. Urlaub, R. Lührmann, Isolation of an active step I spliceosome and composition of its RNP core. Nature 452, 846–850 (2008).
OpenUrl CrossRef PubMed Web of Science

[9] 9.↵
E. T. Wang et al., Alternative isoform regulation in human tissue transcriptomes. Nature 456, 470–476 (2008).
OpenUrl CrossRef PubMed Web of Science

[10] 10.↵
Q. Pan, O. Shai, L. J. Lee, B. J. Frey, B. J. Blencowe, Deep surveying of alternative splicing complexity in the human transcriptome by high-throughput sequencing. Nature genetics 40, 1413–1415 (2008).
OpenUrl CrossRef PubMed Web of Science

[11] 11.↵
T. W. Nilsen, B. R. Graveley, Expansion of the eukaryotic proteome by alternative splicing. Nature 463, 457–463 (2010).
OpenUrl CrossRef PubMed Web of Science

[12] 12.↵
H. Keren, G. Lev-Maor, G. Ast, Alternative splicing and evolution: diversification, exon definition and function. Nat Rev Genet 11, 345–355 (2010).
OpenUrl CrossRef PubMed Web of Science

[13] 13.↵
M. W. Popp, L. E. Maquat, Organizing principles of mammalian nonsense-mediated mRNA decay. Annu Rev Genet 47, 139–165 (2013).
OpenUrl CrossRef PubMed Web of Science

[14] 14.↵
A. G. Jacob, C. W. J. Smith, Intron retention as a component of regulated gene expression programs. Hum Genet 136, 1043–1057 (2017).
OpenUrl CrossRef

[15] 15.↵
P. L. Boutz, A. Bhutkar, P. A. Sharp, Detained introns are a novel, widespread class of post-transcriptionally spliced introns. Genes Dev 29, 63–80 (2015).
OpenUrl Abstract/FREE Full Text

[16] 16.↵
K. Yap, Z. Q. Lim, P. Khandelia, B. Friedman, E. V. Makeyev, Coordinated regulation of neuronal mRNA steady-state levels through developmentally controlled intron retention. Genes Dev 26, 1209–1223 (2012).
OpenUrl Abstract/FREE Full Text

[17] 17.
O. Mauger, F. Lemoine, P. Scheiffele, Targeted Intron Retention and Excision for Rapid Gene Regulation in Response to Neuronal Activity. Neuron 92, 1266–1278 (2016).
OpenUrl CrossRef

[18] 18.
J. Gill et al., Regulated intron removal integrates motivational state and experience. Cell 169, 836–848. e815 (2017).
OpenUrl CrossRef PubMed

[19] 19.
C. Naro et al., An Orchestrated Intron Retention Program in Meiosis Controls Timely Usage of Transcripts during Germ Cell Differentiation. Dev Cell 41, 82–93 e84 (2017).
OpenUrl CrossRef

[20] 20.↵
K. Ninomiya, N. Kataoka, M. Hagiwara, Stress-responsive maturation of Clk1/4 pre- mRNAs promotes phosphorylation of SR splicing factor. J Cell Biol 195, 27–40 (2011).
OpenUrl Abstract/FREE Full Text

[21] 21.
K. E. Pendleton et al., The U6 snRNA m(6)A Methyltransferase METTL16 Regulates SAM Synthetase Intron Retention. Cell 169, 824–835 e814 (2017).
OpenUrl CrossRef PubMed

[22] 22.
S. K. Park et al., A Conserved Splicing Silencer Dynamically Regulates O-GlcNAc Transferase Intron Retention and O-GlcNAc Homeostasis. Cell Rep 20, 1088–1099 (2017).
OpenUrl CrossRef PubMed

[23] 23.↵
Z. W. Tan et al., O-GlcNAc regulates gene expression by controlling detained intron splicing. Nucleic Acids Res 48, 5656–5669 (2020).
OpenUrl CrossRef

[24] 24.↵
E. S. Lander et al., Initial sequencing and analysis of the human genome. Nature 409, 860–921 (2001).
OpenUrl CrossRef PubMed Web of Science

[25] 25.
C. N. Tennyson, H. J. Klamut, R. G. Worton, The human dystrophin gene requires 16 hours to be transcribed and is cotranscriptionally spliced. Nat Genet 9, 184–190 (1995).
OpenUrl CrossRef PubMed Web of Science

[26] 26.
X. Darzacq et al., In vivo dynamics of RNA polymerase II transcription. Nat Struct Mol Biol 14, 796–806 (2007).
OpenUrl CrossRef PubMed Web of Science

[27] 27.↵
J. Singh, R. A. Padgett, Rates of in situ transcription and splicing in large human genes. Nat Struct Mol Biol 16, 1128–1133 (2009).
OpenUrl CrossRef PubMed Web of Science

[28] 28.↵
Y. Jia, J. C. Mu, S. L. Ackerman, Mutation of a U2 snRNA gene causes global disruption of alternative splicing and neurodegeneration. Cell 148, 296–308 (2012).
OpenUrl CrossRef PubMed Web of Science

[29] 29.↵
J. J. Wong et al., Orchestrated intron retention regulates normal granulocyte differentiation. Cell 154, 583–595 (2013).
OpenUrl CrossRef PubMed Web of Science

[30] 30.↵
U. Braunschweig et al., Widespread intron retention in mammals functionally tunes transcriptomes. Genome Res 24, 1774–1786 (2014).
OpenUrl Abstract/FREE Full Text

[31] 31.↵
L. E. Maquat, Nonsense-mediated mRNA decay: splicing, translation and mRNP dynamics. Nat Rev Mol Cell Biol 5, 89–99 (2004).
OpenUrl CrossRef PubMed Web of Science

[32] 32.↵
B. M. Venkatesan, R. Bashir, Nanopore sensors for nucleic acid analysis. Nat Nanotechnol 6, 615–624 (2011).
OpenUrl CrossRef PubMed

[33] 33.↵
R. H. Kim et al., SNIP1 inhibits NF-κB signaling by competing for its binding to the C/H1 domain of CBP/p300 transcriptional co-activators. Journal of Biological Chemistry 276, 46297–46304 (2001).
OpenUrl Abstract/FREE Full Text

[34] 34.↵
R. H. Kim et al., A novel smad nuclear interacting protein, SNIP1, suppresses p300- dependent TGF-β signal transduction. Genes & development 14, 1605–1616 (2000).
OpenUrl Abstract/FREE Full Text

[35] 35.↵
C. P. Bracken et al., Regulation of cyclin D1 RNA stability by SNIP1. Cancer Res 68, 7621–7628 (2008).
OpenUrl Abstract/FREE Full Text

[36] 36.↵
S. Picelli et al., Smart-seq2 for sensitive full-length transcriptome profiling in single cells. Nat Methods 10, 1096–1098 (2013).
OpenUrl CrossRef PubMed Web of Science

[37] 37.↵
D. Haselbach et al., Structure and Conformational Dynamics of the Human Spliceosomal B(act) Complex. Cell 172, 454–464 e411 (2018).
OpenUrl CrossRef PubMed

[38] 38.↵
R. Wan, R. Bai, C. Yan, J. Lei, Y. Shi, Structures of the Catalytically Activated Yeast Spliceosome Reveal the Mechanism of Branching. Cell 177, 339–351 e313 (2019).
OpenUrl CrossRef PubMed

[39] 39.↵
C. Townsend et al., Mechanism of protein-guided folding of the active site U2/U6 RNA during spliceosome activation. Science 370, (2020).

[40] 40.
K. Bertram et al., Cryo-EM Structure of a Pre-catalytic Human Spliceosome Primed for Activation. Cell 170, 701–713.e711 (2017).
OpenUrl CrossRef PubMed

[41] 41.↵
X. Zhan, C. Yan, X. Zhang, J. Lei, Y. Shi, Structure of a human catalytic step I spliceosome. Science 359, 537–545 (2018).
OpenUrl Abstract/FREE Full Text

[42] 42.↵
V. L. Reichert, H. Le Hir, M. S. Jurica, M. J. Moore, 5’ exon interactions within the human spliceosome establish a framework for exon junction complex structure and assembly. Genes Dev 16, 2778–2791 (2002).
OpenUrl Abstract/FREE Full Text

[43] 43.↵
D. Durocher, S. P. Jackson, The FHA domain. FEBS Lett 513, 58–66 (2002).
OpenUrl CrossRef PubMed Web of Science

[44] 44.↵
P. Wysoczanski et al., Cooperative structure of the heterotrimeric pre-mRNA retention and splicing complex. Nat Struct Mol Biol 21, 911–918 (2014).
OpenUrl CrossRef PubMed

[45] 45.↵
A. G. Murachelli, J. Ebert, C. Basquin, H. Le Hir, E. Conti, The structure of the ASAP core complex reveals the existence of a Pinin-containing PSAP complex. Nat Struct Mol Biol 19, 378–386 (2012).
OpenUrl CrossRef PubMed

[46] 46.↵
V. Boehm et al., Exon Junction Complexes Suppress Spurious Splice Sites to Safeguard Transcriptome Integrity. Mol Cell 72, 482–495 e487 (2018).
OpenUrl CrossRef PubMed

[47] 47.↵
H. Le Hir, J. Sauliere, Z. Wang, The exon junction complex as a node of post- transcriptional networks. Nat Rev Mol Cell Biol 17, 41–54 (2016).
OpenUrl CrossRef PubMed

[48] 48.
R. Hayashi, D. Handler, D. Ish-Horowicz, J. Brennecke, The exon junction complex is required for definition and excision of neighboring introns in Drosophila. Genes Dev 28, 1772–1785 (2014).
OpenUrl Abstract/FREE Full Text

[49] 49.
C. D. Malone et al., The exon junction complex controls transposable element activity by ensuring faithful splicing of the piwi transcript. Genes Dev 28, 1786–1799 (2014).
OpenUrl Abstract/FREE Full Text

[50] 50.
L. Blazquez et al., Exon Junction Complex Shapes the Transcriptome by Repressing Recursive Splicing. Mol Cell 72, 496–509 e499 (2018).
OpenUrl CrossRef

[51] 51.↵
T. Gonatopoulos-Pournatzis et al., Genome-wide CRISPR-Cas9 Interrogation of Splicing Networks Reveals a Mechanism for Recognition of Autism-Misregulated Neuronal Microexons. Mol Cell 72, 510–524 e512 (2018).
OpenUrl CrossRef PubMed

[52] 52.↵
E. L. Van Nostrand et al., Principles of RNA processing from analysis of enhanced CLIP maps for 150 RNA binding proteins. Genome Biol 21, 90 (2020).

[53] 53.
E. L. Van Nostrand et al., A large-scale binding and functional map of human RNA- binding proteins. Nature 583, 711–719 (2020).
OpenUrl CrossRef PubMed

[54] 54.↵
C. Hauer et al., Exon Junction Complexes Show a Distributional Bias toward Alternatively Spliced mRNAs and against mRNAs Coding for Ribosomal Proteins. Cell Rep 16, 1588–1603 (2016).
OpenUrl CrossRef

[55] 55.↵
T. Pohlkamp et al., Generation and characterization of an Nse-CreERT2 transgenic line suitable for inducible gene manipulation in cerebellar granule cells. PLoS One 9, e100384 (2014).
OpenUrl

[56] 56.↵
S. P. Peron, J. Freeman, V. Iyer, C. Guo, K. Svoboda, A Cellular Resolution Map of Barrel Cortex Activity during Tactile Behavior. Neuron 86, 783–799 (2015).
OpenUrl CrossRef PubMed

[57] 57.↵
E. Aparicio-Prat et al., DECKO: Single-oligo, dual-CRISPR deletion of genomic elements including long non-coding RNAs. BMC Genomics 16, 846 (2015).
OpenUrl CrossRef PubMed

[58] 58.↵
A. Dziembowski et al., Proteomic analysis identifies a new complex required for nuclear pre-mRNA retention and splicing. EMBO J 23, 4847–4856 (2004).
OpenUrl Abstract/FREE Full Text

[59] 59.↵
P. Fabrizio et al., The evolutionarily conserved core design of the catalytic activation step of the yeast spliceosome. Mol Cell 36, 593–608 (2009).
OpenUrl CrossRef PubMed Web of Science

[60] 60.
C. Yan, R. Wan, R. Bai, G. Huang, Y. Shi, Structure of a yeast activated spliceosome at 3.5 å resolution. Science 353, 904–911 (2016).
OpenUrl Abstract/FREE Full Text

[61] 61.↵
C. Schneider et al., Dynamic Contacts of U2, RES, Cwc25, Prp8 and Prp45 Proteins with the Pre-mRNA Branch-Site and 3’ Splice Site during Catalytic Activation and Step 1 Catalysis in Yeast Spliceosomes. PLoS Genet 11, e1005539 (2015).
OpenUrl CrossRef PubMed

[62] 62.↵
A. Gottschalk, C. Bartels, G. Neubauer, R. Luhrmann, P. Fabrizio, A novel yeast U2 snRNP protein, Snu17p, is required for the first catalytic step of splicing and for progression of spliceosome assembly. Mol Cell Biol 21, 3037–3046 (2001).
OpenUrl Abstract/FREE Full Text

[63] 63.↵
J. P. Fernandez et al., RES complex is associated with intron definition and required for zebrafish early embryogenesis. PLoS Genet 14, e1007473 (2018).
OpenUrl CrossRef

[64] 64.↵
L. Frankiw et al., BUD13 Promotes a Type I Interferon Response by Countering Intron Retention in Irf7. Mol Cell 73, 803–814 e806 (2019).
OpenUrl CrossRef

[65] 65.↵
R. F. Luco et al., Regulation of alternative splicing by histone modifications. Science 327, 996–1000 (2010).
OpenUrl Abstract/FREE Full Text

[66] 66.↵
Z. Xing et al., lncRNA directs cooperative epigenetic regulation downstream of chemokine signals. Cell 159, 1110–1125 (2014).
OpenUrl CrossRef PubMed

[67] 67.↵
J. Lykke-Andersen, M. D. Shu, J. A. Steitz, Communication of the position of exon-exon junctions to the mRNA surveillance machinery by the protein RNPS1. Science 293, 1836–1839 (2001).
OpenUrl Abstract/FREE Full Text

[68] 68.
H. Le Hir, E. Izaurralde, L. E. Maquat, M. J. Moore, The spliceosome deposits multiple proteins 20-24 nucleotides upstream of mRNA exon-exon junctions. EMBO J 19, 6860–6869 (2000).
OpenUrl Abstract/FREE Full Text

[69] 69.↵
N. H. Gehring et al., Exon-junction complex components specify distinct routes of nonsense-mediated mRNA decay with differential cofactor requirements. Mol Cell 20, 65–75 (2005).
OpenUrl CrossRef PubMed Web of Science

[70] 70.↵
C. Schwerk et al., ASAP, a novel protein complex involved in RNA processing and apoptosis. Molecular and Cellular Biology 23, 2981–2990 (2003).
OpenUrl Abstract/FREE Full Text

[71] 71.↵
T. O. Tange, T. Shibuya, M. S. Jurica, M. J. Moore, Biochemical analysis of the EJC reveals two new factors and a stable tetrameric protein core. RNA 11, 1869–1883 (2005).
OpenUrl Abstract/FREE Full Text

[72] 72.↵
C. Girard et al., Post-transcriptional spliceosomes are retained in nuclear speckles until splicing completion. Nat Commun 3, 994 (2012).

[73] 73.
C. Li, R. I. Lin, M. C. Lai, P. Ouyang, W. Y. Tarn, Nuclear Pnn/DRS protein binds to spliced mRNPs and participates in mRNA processing and export via interaction with RNPS1. Mol Cell Biol 23, 7363–7376 (2003).
OpenUrl Abstract/FREE Full Text

[74] 74.
A. Mayeda et al., Purification and characterization of human RNPS1: a general activator of pre-mRNA splicing. EMBO J 18, 4560–4570 (1999).
OpenUrl Abstract/FREE Full Text

[75] 75.↵
E. Sakashita, S. Tatsumi, D. Werner, H. Endo, A. Mayeda, Human RNPS1 and its associated factors: a versatile alternative pre-mRNA splicing regulator in vivo. Mol Cell Biol 24, 1174–1187 (2004).
OpenUrl Abstract/FREE Full Text

[76] 76.↵
J. H. Trembley et al., Activation of pre-mRNA splicing by human RNPS1 is regulated by CK2 phosphorylation. Mol Cell Biol 25, 1446–1457 (2005).
OpenUrl Abstract/FREE Full Text

[77] 77.↵
P. Bao, C. L. Will, H. Urlaub, K. L. Boon, R. Luhrmann, The RES complex is required for efficient transformation of the precatalytic B spliceosome into an activated B(act) complex. Genes Dev 31, 2416–2429 (2017).
OpenUrl Abstract/FREE Full Text

[78] 78.↵
S. Ullrich, R. Guigo, Dynamic changes in intron retention are tightly associated with regulation of splicing factors and proliferative activity during B-cell development. Nucleic Acids Res 48, 1327–1340 (2020).
OpenUrl

[79] 79.↵
S. Adusumalli, Z. K. Ngian, W. Q. Lin, T. Benoukraf, C. T. Ong, Increased intron retention is a post-transcriptional signature associated with progressive aging and Alzheimer’s disease. Aging Cell 18, e12928 (2019).

[80] 80.↵
Q. Wang, E. G. Conlon, J. L. Manley, D. C. Rio, Widespread intron retention impairs protein homeostasis in C9orf72 ALS brains. Genome Res 30, 1705–1715 (2020).
OpenUrl Abstract/FREE Full Text

[81] 81.↵
B. Lizen, M. Claus, L. Jeannotte, F. M. Rijli, F. Gofflot, Perinatal induction of Cre recombination with tamoxifen. Transgenic Res 24, 1065–1077 (2015).
OpenUrl CrossRef

[82] 82.↵
M. E. Pitulescu, I. Schmidt, R. Benedito, R. H. Adams, Inducible gene targeting in the neonatal vasculature and analysis of retinal angiogenesis in mice. Nat Protoc 5, 1518–1534 (2010).
OpenUrl CrossRef PubMed

[83] 83.↵
A. P. Salinger, M. J. Justice, Mouse mutagenesis using N-ethyl-N-nitrosourea (ENU). Cold Spring Harbor Protocols 2008, pdb. prot4985 (2008).

[84] 84.↵
R.-Z. Chen et al., An ENU-induced mutation in Twist1 transactivation domain causes hindlimb polydactyly with complete penetrance and dominant-negatively impairs E2A-dependent transcription. Scientific reports 10, 1–12 (2020).
OpenUrl

[85] 85.↵
A. McKenna et al., The Genome Analysis Toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data. Genome research 20, 1297–1303 (2010).
OpenUrl Abstract/FREE Full Text

[86] 86.↵
K. Wang, M. Li, H. Hakonarson, ANNOVAR: functional annotation of genetic variants from high-throughput sequencing data. Nucleic acids research 38, e164–e164 (2010).
OpenUrl CrossRef PubMed

[87] 87.↵
P. D. Hsu et al., DNA targeting specificity of RNA-guided Cas9 nucleases. Nature biotechnology 31, 827–832 (2013).
OpenUrl CrossRef PubMed

[88] 88.↵
F. A. Ran et al., Genome engineering using the CRISPR-Cas9 system. Nature protocols 8, 2281–2308 (2013).
OpenUrl

[89] 89.↵
O. Shalem et al., Genome-scale CRISPR-Cas9 knockout screening in human cells. Science 343, 84–87 (2014).
OpenUrl Abstract/FREE Full Text

[90] 90.↵
S. Boelz, G. Neu-Yilik, N. H. Gehring, M. W. Hentze, A. E. Kulozik, A chemiluminescence- based reporter system to monitor nonsense-mediated mRNA decay. Biochemical and biophysical research communications 349, 186–191 (2006).
OpenUrl CrossRef PubMed Web of Science

[91] 91.↵
H. W. Hwang, E. A. Wentzel, J. T. Mendell, A hexanucleotide element directs microRNA nuclear import. Science 315, 97–100 (2007).
OpenUrl Abstract/FREE Full Text

[92] 92.↵
D. Kim, J. M. Paggi, C. Park, C. Bennett, S. L. Salzberg, Graph-based genome alignment and genotyping with HISAT2 and HISAT-genotype. Nature biotechnology 37, 907–915 (2019).
OpenUrl CrossRef PubMed

[93] 93.↵
S. Anders, A. Reyes, W. Huber, Detecting differential usage of exons from RNA-seq data. Genome Res 22, 2008–2017 (2012).
OpenUrl Abstract/FREE Full Text

[94] 94.↵
H. Mi, A. Muruganujan, J. T. Casagrande, P. D. Thomas, Large-scale gene function analysis with the PANTHER classification system. Nat Protoc 8, 1551–1566 (2013).
OpenUrl CrossRef PubMed

[95] 95.↵
H. Li et al., The sequence alignment/map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).
OpenUrl CrossRef PubMed Web of Science

[96] 96.↵
A. R. Quinlan, BEDTools: The Swiss-Army Tool for Genome Feature Analysis. Curr Protoc Bioinformatics 47, 11 12 11–34 (2014).
OpenUrl CrossRef

[97] 97.↵
J. L. Rinn et al., Functional demarcation of active and silent chromatin domains in human HOX loci by noncoding RNAs. cell 129, 1311–1323 (2007).
OpenUrl CrossRef PubMed Web of Science

[98] 98.↵
S. Picelli et al., Full-length RNA-seq from single cells using Smart-seq2. Nature protocols 9, 171–181 (2014).
OpenUrl

[99] 99.↵
E. P. Consortium, An integrated encyclopedia of DNA elements in the human genome. Nature 489, 57 (2012).

[100] 100.
J. Zhang et al., An integrative ENCODE resource for cancer genomics. Nature communications 11, 1–11 (2020).
OpenUrl

[101] 101.↵
A. Athar, et al., ArrayExpress update - from bulk to single-cell expression data. Nucleic Acids Res 47, D711–D715 (2019).
OpenUrl CrossRef PubMed

[102] 102.↵
A. Dobin et al., STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
OpenUrl CrossRef PubMed Web of Science

[103] 103.↵
E. L. Van Nostrand et al., Robust transcriptome-wide discovery of RNA-binding protein binding sites with enhanced CLIP (eCLIP). Nat Methods 13, 508–514 (2016).
OpenUrl CrossRef PubMed

[104] 104.↵
M. T. Lovci et al., Rbfox proteins regulate alternative mRNA splicing through evolutionarily conserved RNA bridges. Nature structural & molecular biology 20, 1434–1442 (2013).
OpenUrl