Phosphorylation of SF3B1 by CDK11 orchestrates spliceosome activation via SNIP1-dependent RES complex recruitment

Domain: biology
Published: 2026-03-29
Editorial outcome: major_revision

Abstract
Splicing Factor 3b Subunit 1 (SF3B1), a core component of the spliceosome, undergoes dynamic
phosphorylation and dephosphorylation during the splicing cycle to regulate pre-mRNA splicing.
Twenty-eight threonine/proline repeats are phosphorylated by CDK11 during spliceosome
activation and remain phosphorylated in the catalytically active spliceosomes. The function of
phosphorylated SF3B1 (P-SF3B1), and the identity of spliceosomes stalled by CDK11 inhibition
remain unclear. Using quantitative proteomics of chromatin-associated spliceosomes, we identify
a previously uncharacterized intermediate complex BOTS964, arrested by CDK11 inhibitor OTS964,
that incorporates the nineteen-related (NTR) but not nineRteen (NTC) complex. iCLIP-seq revealed
that P-SF3B1 engages with the U6 snRNA internal stem-loop (ISL), suggesting a potential role in
stabilizing the RNA catalytic core. We further demonstrate that P-SF3B1 is recognized by
forkhead-associated (FHA) domain of SNIP1, which promotes recruitment of retention and
splicing (RES) complex during spliceosome activation. Acute SNIP1 depletion disrupts RES
incorporation, causes widespread splicing defects, and promotes hyperphosphorylation of SF3B1
by CDK11. Mutations in SNIP1 FHA domain, including the neurodevelopmental disorder-
associated E366G variant, impair P-SF3B1 binding, pre-mRNA splicing, and cell viability.
Together, these findings uncover the phosphorylation-dependent CDK11/P-SF3B1/SNIP1
signaling axis that is critical for pre-mRNA splicing and cellular proliferation and provide a
mechanistic insight into its dysregulation in disease.

Introduction
Transcription of eukaryotic genes produces RNA precursors called pre-mRNAs, which contain
non-coding introns. Introns are removed by spliceosome in the process of pre-mRNA splicing. The
spliceosome is a dynamic RNA-protein complex that assembles on intron sequences through a
series of steps involving the sequential recruitment and release of five small nuclear
ribonucleoprotein particles (snRNPs): U1, U2, U4, U5, and U6, along with numerous non-snRNP
proteins. This process is accompanied by significant structural rearrangements. As a result, a series
of intermediate spliceosome complexes are formed, including ES, A, B, Bact, B*, C, C*, P, and ILS.
Pre-mRNA splicing consists of four key steps: spliceosome assembly (E→B), catalytic activation
(B→B*), two-step catalysis of splicing (B*→P), and spliceosome disassembly (P→ILS) 1-6. While
the roles of kinases and their substrates in facilitating transitions between intermediates are
essential, they remain poorly understood 7-9.

Activation of the spliceosome occurs as it transitions from the B to the Bact complex, leading to
the formation of the catalytically active U2/U6 snRNA structure. This activation progresses via
two intermediates, pre-Bact1 and pre-Bact2, and requires the release and recruitment of about 20 and
25 proteins, respectively 10-12. A key event in this activation is the unwinding of the U4/U6 snRNA
duplex, followed by the release of the U4 snRNP and B-specific proteins, and the recruitment of
various protein complexes such as the NTC (Nineteen), NTR (Nineteen-related), RES (retention
and splicing), and IBC (intron-binding) 12-16. In the activated spliceosome (Bact), the newly formed
U2/U6 snRNA catalytic center is stabilized by various proteins, including components of the NTR
and NTC complexes 12,17-19. The catalytic center remains static until the ILS complex 4. In higher
eukaryotes, this activation also requires the action of CDK11, which phosphorylates SF3B1, a core
splicing factor and component of the U2 snRNP. The inhibition of CDK11 with OTS964 traps the
spliceosome in a B-like intermediate, in which U4 snRNA and B-specific proteins remain
associated, and RES and IBC complexes unbound 8,20. The precise composition of the stalled
spliceosome however, remains unknown.

Human SF3B1 consists of an N-terminal intrinsically disordered region (IDR) and a C-terminal
twenty HEAT-repeats domain 21. These regions serve distinct and essential roles in the
spliceosome lifecycle and are targeted by small molecules that disrupt spliceosome assembly and
have antiproliferative effects 8,22-26. While the functions of the HEAT-repeats are relatively well-
understood 21, the roles of the IDR, due to its structural flexibility, which prevents its visualization
in cryo-EM structures, remains largely unclear 20. The IDR acts as a dynamic interaction hub for
proteins and RNA 8,27-30, containing a cluster of 28 threonine/proline (Thr/Pro) CDK-consensus
sites that overlap with several U2AF homology ligand motifs (ULM)s 20,27,28,31,32. These motifs
serve as binding sites for splicing factors (such as U2AF65, PUF60, TAT-SF1, RBM39, and
SPF45(RBM17) when IDR is unphosphorylated in the A intermediate 31,33-36. Upon the formation
of the Bact complex, the IDR becomes fully phosphorylated, a state that persists until the second
catalytic reaction (C complex) 8,20,27, when SF3B1 is dephosphorylated by the PP1 phosphatase
37,38. RNA immunoprecipitation coupled to qPCR (RIP-qPCR) showed that phosphorylated SF3B1
(P-SF3B1) associates with U6 snRNA 8. However, it remains unclear whether this interaction is
direct and where at U6 snRNA the P-SF3B1 binds. Additionally, interacting proteins (readers) of
P-SF3B1 and their functions in the activated and catalytically active spliceosomes (B*→C) also
remain elusive.

SNIP1, a component of the trimeric RES complex (which also includes RBMX2 and BUD13) 39,40,
is integrated into the spliceosome during activation and is essential for the assembly of Bact in both
yeast and humans 12,13,40,41. The knockdown of SNIP1 and other RES subunits results in retention
of mostly short and GC-rich introns in human cells 42. A similar phenotype is observed in zebrafish
models with loss-of-function mutations in RES components 43. SNIP1 contains a conserved
forkhead-associated (FHA) domain, a putative P-Thr-binding module 44,45, with unknown
function. A recent study suggested that it is required for the interaction with RNPS1, possibly
affecting the post-transcriptional splicing of detained introns 46. A biallelic mutation (E366G) in
SNIP1, located near the FHA domain, has been associated with a complex neurodevelopmental
disorder in 35 individuals from the Amish community 47,48, although the underlying mechanistic
defect caused by the mutation is unknown. Other aberrations of SNIP1 have been linked to
neurogenesis and neurodegeneration, both in human and zebrafish 43,46,48,49. SNIP1 has also been
implicated in transcription regulation and oncogenesis 50-55, highlighting its clinical relevance.

In this study, we demonstrate that the CDK11 inhibitor OTS964 traps the spliceosome in a novel
intermediate, with the NTR complex integrated but the NTC complex not associated. We show
that P-SF3B1 localizes to U6 snRNA´s internal stem-loop (ISL), potentially contributing to the
formation and stabilization of the U2/U6 snRNA catalytic center in the activated spliceosomes.
Additionally, we identify the FHA domain of SNIP1 as a “reader” of P-SF3B1, guiding the
recruitment and assembly of the RES complex during spliceosome activation, and show that
mutations in the FHA domain, including the neurodevelopmental disorder-linked E366G variant,
impair this interaction, resulting in defects in splicing and cellular proliferation. Overall, our
findings define the roles of CDK11 kinase activity and the phosphorylated form of SF3B1 during
spliceosome activation and in the catalytically active spliceosome.

Results
OTS964 blocks the spliceosome in a B-Bact intermediate with integrated NTR and unbound NTC.
Previously, we showed that CDK11 inhibition by OTS964 blocks spliceosome during B to Bact
transition, in a B-like intermediate 8. To define a CDK11-dependent rate-limiting step in
spliceosome activation, we decided to determine the precise protein composition of spliceosomes
blocked by OTS964 in cells. We took advantage of the fact that most pre-mRNA splicing occurs
co-transcriptionally and that spliceosomes are tightly associated with chromatin during this process
8,27,56-58. Splicing inhibitors cause the accumulation of stalled spliceosome intermediates on
chromatin, enabling their isolation and characterization directly from living cells 8,59. We treated
HCT116 cells with either DMSO (control) or 50 nM OTS964, isolated chromatin fractions, and
digested them with micrococcal nuclease (MNase) 8,59,60. The released spliceosome complexes
were immunoprecipitated with SF3B1 antibody, and the composition of SF3B1-associated
complexes was determined by mass spectrometry (Fig. 1a). Since SF3B1 is a core component of
the spliceosome (Fig. 1b), the analyses of SF3B1-associated proteins reflect the composition of a
spliceosome intermediate.

In DMSO-treated cells, approximately 30% of SF3B1-associated proteins were classified as RNA
splicing factors (GO biological process: RNA splicing) (Supplementary Data 1, sheet “IP SF3B1
(Chromatin) all data”). CDK11 inhibition significantly altered the association of 78 splicing factors
with SF3B1, with 52 proteins decreased and 26 increased in abundance (-0.5 > log2FC > 0.5, Padj
<0.05) (Fig. 1c, Supplementary Data 1, sheet “IP SF3B1 (Chromatin) all data”). Since our
previous work identified a critical role for CDK11 in the B to Bact transition 8, we followed changes
in snRNP and non-snRNP complexes involved in this transition 12. As expected, the levels of U2
snRNP and SF3B1 remained unchanged after OTS964 treatment (Fig. 1d, Supplementary Data
1, sheet “IP SF3B1 (Chromatin) splicing”). In contrast, the components of B intermediate-
associated complexes, namely U5 and U4/U6 snRNPs and B-specific proteins, increased in
abundance (Fig. 1d, Supplementary Data 1, sheet “IP SF3B1 (Chromatin) splicing”), consistent
with their accumulation upstream of a blocked transition during spliceosome activation 8. Protein
complexes that are normally recruited during spliceosome activation, such as RES, NTC and IBC,
were significantly reduced, with the notable exception of NTR, which remained unchanged,
suggesting that it was already integrated into the spliceosome. This result suggests that OTS964
arrests spliceosome progression during activation in a previously uncharacterized intermediate.

To validate this result by a method that omits the immunoprecipitation step, we isolated soluble
and chromatin fractions from cells and quantified protein levels as proxies for the presence of their
respective spliceosome complexes 8,59 (Supplementary Fig. 1a). After CDK11 inhibition, the
chromatin abundance of proteins associated with E (U2AF65), and A (RBM17) intermediates as
well as the U4/U6 snRNP (NHP2L1) complex remained equal. Meanwhile, the levels of U5
snRNP (EFTUD2, PRPF4, TXNL4A) and B-specific proteins (SMU1, IK, WBP4 and MFAP1) 
increased, and the amount of NTR (SNW1, RBM22, BUD31) complex remained unchanged
(Supplementary Fig. 1a). In contrast, proteins from RES (SNIP1, BUD13, RBMX2), NTC
(CDC5L, PRPF19), IBC (AQR) and exon-junction (EJC) (RBM8A) complexes exhibited
decreased chromatin association (Supplementary Fig. 1a). The optimal fractionation of
chromatin and effect of treatment with OTS964 were verified by histone H2A and SF3B1
phosphorylation, respectively (Supplementary Fig. 1a).

This result suggests that OTS964 arrests spliceosome progression during activation in an
intermediate with integrated NTR complex, and NTC complex still not associated. We refer to this
intermediate as BOTS964.

P-SF3B1 is enriched at U6 snRNA ISL in the activated spliceosome.
The N-terminal region of SF3B1 is a large IDR containing multiple phosphorylatable Thr/Pro
repeats 8,20. As for most other IDRs of core spliceosome factors that are not resolved in cryo-EM
structures, the functional roles of the P-SF3B1 in the activated and catalytically active
spliceosomes remain unknown. Here we applied UV crosslinking and an immunoprecipitation
(iCLIP), a method that identifies direct protein-RNA interactions at single nucleotide resolution
61,62 to investigate the interaction of P-SF3B1 with U6 snRNA 8, a component of catalytically
active U2/U6 snRNA structure 63-66.

P-SF3B1 was immunoprecipitated from HCT116 cells, treated or untreated with OTS964. To
ensure the specificity of the P-SF3B1 iCLIP binding profiles, we used two phospho-specific
antibodies recognizing either P-Thr211 or P-Thr235 of SF3B1 8. Because iCLIP replicates
generated with the P-Thr211 and P-Thr235 antibodies correlated well, we combined the replicates
for each antibody and used them for further genome-wide analyses (Supplementary Fig. 2a).

To determine the contacts between P-SF3B1 and U6 snRNA, we analyzed crosslinking patterns of
P-Thr211 and P-Thr235 SF3B1 iCLIPs within U6 snRNA. Both datasets revealed a distinct peak
around nucleotide 66 in the ISL of U6 snRNA (Fig. 2a, b), suggesting an association of P-SF3B1
with this catalytically important RNA element of U6 snRNA formed in the activated spliceosome
12 (Fig. 2c). Previously published SF3B1 iCLIP data 8 showed a detectable peak at the same
position in the U6 ISL, which was insensitive to OTS964 treatment (Fig. 2d). In contrast, OTS964
treatment reduced contacts between P-SF3B1 and the ISL U6 two- to three-fold relative to total
SF3B1 at this specific position (Fig. 2a, b, d), providing further evidence for the specificity of P-
SF3B1 association with this U6 snRNA element.

As expected, neither P-SF3B1 nor total SF3B1 crosslinked to U1 snRNA which binds to 5´ splice
sites during early stages of spliceosome assembly (Supplementary Fig. 2b). By contrast, U2
snRNA, another component of the catalytically active U2/U6 snRNA spliceosome structure, was
bound predominantly by total SF3B1 and to lesser extent by P-SF3B1 (Supplementary Fig. 2c)8.

P-SF3B1 interacts with SNIP1 protein
To identify proteins that preferentially associate with P-SF3B1 in the spliceosome, we performed
a pull-down assay followed by mass spectrometry (Fig. 3a). We expressed and purified the GST-
tagged N-terminus of SF3B1 (residues 1-463) from E. coli, and phosphorylated it in vitro using
flag-tagged CDK11 purified from HEK293 cells. A non-phosphorylated version of the protein was
used as a control. GST-SF3B1 proteins were immobilized on glutathione resin and incubated with
soluble nuclear lysate from HCT116 cells. To minimize unspecific interactions with pre-assembled
spliceosome complexes, we removed the chromatin fraction and used soluble nuclear lysate treated
with RNase A to further promote the release of proteins from various RNA-dependent
subcomplexes of the spliceosome. After stringent washes, SF3B1-interacting proteins were
identified by mass spectrometry (Fig. 3a).

A comparison of proteins enriched (-1 > log2FC > 1, Padj <0.05) with phosphorylated or
unphosphorylated SF3B1 identified 8 and 26 interactors, respectively (Fig. 3b, Supplementary
Data 2). As expected, known ULM-binding proteins U2AF65 and PUF60 were detected, and
RBM39 and RBM17 31,33,34,36 were also enriched on unphosphorylated SF3B1, albeit below the
statistical significance threshold (Fig. 3b, Supplementary Data 2). These proteins are associated
with the A spliceosome intermediate in which SF3B1 remains unphosphorylated (Fig. 1b) 20,
validating the assay´s biological relevance. Among proteins preferentially associated with P-
SF3B1 was identified NIPP1, a PP1 adaptor protein that interacts with P-SF3B1 via its FHA
domain 67,68 and SNIP1, another FHA domain-containing protein and component of the RES
complex 39-41 (Fig. 3b). The presence of CDK11 and components of its complex, CCNL1/L2 and
SAP30BP 69,70, is a consequence of P-SF3B1 sample preparation (Fig. 3a, b). To validate these
interactions, we performed GST pull-down assays using (P)-SF3B1 and analyzed bound proteins
by western blotting. U2AF65 and RBM17 preferentially bound unphosphorylated SF3B1
(Supplementary Fig. 3a) and SNIP1 preferred the phosphorylated one (Fig. 3c). Notably, CDK11
remained associated with P-SF3B1 even after multiple washes, in line with the proteomics results
(Fig. 3c). Since SNIP1 contains a P-Thr binding FHA domain and is recruited to the spliceosome
during its activation (B to Bact), we focused on its functional characterization.

Acute SNIP1 depletion leads to a widespread splicing defect
To investigate the role of SNIP1 in pre-mRNA splicing and spliceosome assembly, we tagged the
endogenous SNIP1 with an inducible-degron (dTAG) system 71. dTAG facilitates the acute and
often near-complete depletion of a protein of interest, allowing for the examination of its primary
function in dynamic processes such as splicing 8,72,73. We introduced a cassette encoding a
hemagglutinin (HA) tag and the FKBP12F36V degron into both alleles of the SNIP1 gene by
CRISPR/Cas9-mediated genome editing in HCT116 cells (Supplementary Fig. 4a). Accurate and
homozygous integration into genomic DNA was verified by sequencing (Supplementary Fig.
4b). Comparative western blot analysis of cell lysates from HCT116 and HCT116 dTAG SNIP1
cells confirmed the absence of the wild-type and the appropriate size of the tagged SNIP1 in the
edited cell line (Supplementary Fig. 4c). Upon addition of the small molecule inducer dTAG-v1,
SNIP1 was depleted specifically in the dTAG-tagged cell line, with > 90% degradation observed
within 2 h of the treatment (Supplementary Fig. 4c).

To assess how acute SNIP1 depletion impacts pre-mRNA splicing, we performed total RNA-seq
on nuclear extracts from HCT116 dTAG SNIP1 cells treated with either 500 nM dTAG-v1 or the
DMSO control for 4 h. Visual inspection of individual genes showed widespread intron retention
in transcripts from cells treated with dTAG-v1 (Fig. 4a, Supplementary Fig. 4d). For global
splicing analyses, we only considered the 6222 major isoforms of protein-coding genes with a
prevalence higher than 70%, no overlap with other genes, and with read counts per kilobase (RPK)
higher than 50, as previously described 8,73. The splicing ratio was calculated separately for 5´SS
and 3´SS by dividing the number of spliced reads by the total number of spliced and unspliced
reads crossing the respective SS. We observed significant (Wilcoxon signed rank test; p<2.2.10-
16) accumulations of unspliced reads when all the introns were considered (Fig. 4b) and also when
first, middle, and last introns were analyzed separately (Supplementary Fig. 4e). To rigorously
compare the effects of CDK11 inhibition and SNIP1 loss on splicing we compared fold changes
of intron ratios between OTS964- and DMSO-treated HCT116 cells from our previous study 8 and
between dTAG-induced SNIP1 depletion and DMSO control in dTAG SNIP1 HCT116 cells
across 12727 protein-coding genes. Intron ratios per gene were calculated as the ratio between the
total number of intronic and exonic reads for each gene as described previously 8. Only genes for
which intron ratios varied by less than two-fold between replicates in each condition were
included. This analysis revealed a strong correlation between CDK11 inhibition and SNIP1
depletion (Spearman correlation coefficient = 0.79) providing evidence that CDK11 and SNIP1
affect splicing in a similar manner (Fig. 4c).

Highly similar splicing defects, observed both globally (Fig. 4b, c, Supplementary Fig. 4e) and
in individual transcripts (Fig. 4a, Supplementary Fig. 4d) after SNIP1 depletion and CDK11
inhibition 8, strongly suggest that both proteins act in the same step of spliceosome regulation. To
determine the composition of a spliceosome, intermediate after acute SNIP1 depletion and
compare it to BOTS964, we isolated chromatin fractions as in Supplementary Fig. 1a.
Immunoblotting analyses of chromatin-bound spliceosome complexes after acute SNIP1 depletion
revealed reduced levels of RES components (BUD13 and RBMX2), indicating the failure to recruit
the RES complex during the transition from B to Bact (Fig. 4d). Notably, while some subunits of
spliceosome complexes normally recruited during its activation (e.g. NTR, NTC, IBC) were still
incorporated (e.g. SNW1, RBM22, PRPF19, AQR), the others were decreased (e.g. BCAS2,
CDC5L, CWC22). Components recruited at or after the Bact stage, such as those from the EJC, C
and C* 12 complexes, were reduced or absent (Fig. 4d). The optimal fractionation of chromatin
and SNIP1 depletion were verified by using histone H4 and SNIP1 antibodies, respectively (Fig.
4d).

In summary, our results demonstrate that SNIP1 is essential for the proper recruitment of the RES
complex, and its depletion results in the accumulation of a mis-assembled, likely dead-end
spliceosome intermediate leading to widespread splicing defects.

Acute SNIP1 depletion induces OTS964-sensitive SF3B1 hyperphosphorylation
While analyzing the composition of chromatin-associated spliceosomes after either CDK11
inhibition or SNIP1 depletion, we observed an increase in U5 snRNP and B-specific proteins in
both conditions (Supplementary Fig. 1a and Fig. 4d). We interpret this accumulation as a
consequence of spliceosome stalling at the B to Bact transition. However, these perturbations had
the opposite effects on SF3B1 phosphorylation: while CDK11 inhibition diminished
phosphorylation 8 (Supplementary Fig. 1a), SNIP1 depletion caused a marked increase (Fig. 4d,
Supplementary Fig. 4c). We interpret this increase as a rapid compensatory response to the
absence of SNIP1, indicating that SF3B1 phosphorylation is a prerequisite for proper SNIP1
recruitment. An alternative explanation for the SF3B1 hyperphosphorylation is a “spliceosome
cycle arrest” with SF3B1 dephosphorylation by phosphatases being blocked while CDK11 activity
remains high. Altogether, these results are consistent with CDK11 acting as a primary kinase for
SF3B1 phosphorylation 8, and suggest that P-SF3B1 is required for the efficient integration of
SNIP1 and the RES complex to the spliceosome during its activation. To directly determine
whether this hyperphosphorylation is CDK11-dependent, we first depleted SNIP1 to induce
SF3B1 phosphorylation and subsequently treated cells with OTS964 (Fig. 5a, b, c,
Supplementary Fig. 5a). Inhibition of CDK11 for 30 min fully abrogated the phosphorylation of
SF3B1 as well as phosphorylation of Thr806 in SPT5, a recently identified CDK11 substrate 74. In
contrast, phosphorylation of the RNA Polymerase II (RNAPII) C-terminal domain (CTD) on
serine 2, 5 and 7 (Ser2, Ser5 and Ser7) were reduced gradually and less specifically, as expected
upon constitutive splicing inhibition 8,73 (Fig. 5c, Supplementary Fig. 5a).

Given that previous studies have implicated transcriptional CDKs (e.g., CDK7, CDK9,
CDK12/13) in SF3B1 phosphorylation 75-79, we compared the effects of their inhibition with
OTS964. Treatment with the CDK9 inhibitor NVP-2 80, CDK12/13 inhibitor THZ531 81 or CDK7
inhibitor SY-351 75 for 30 min did not reduce P-SF3B1 levels (Fig. 5a, c, Supplementary Fig.
5a). Only a modest decrease in SF3B1 phosphorylation was observed after 1 h of CDK9 and CDK7
inhibition, whereas longer treatments (≥ 2 h) led to a substantial reduction in P-SF3B1 levels for
all inhibitors (Fig. 5c, Supplementary Fig. 5a). The decrease in SF3B1 phosphorylation observed
after longer inhibition (≥ 1 h) can be attributed to roles of these kinases in transcription, a process
upstream of splicing 8,20,72, and, in the case of CDK7 to its recently described role in CDK11
activation 75,82,83. Effective inhibition of CDK7, CDK9 and CDK12 was confirmed by the
expected changes in RNAPII CTD and SPT5 phosphorylation. Specifically, phosphorylation of
Ser5 and Ser7 in the CTD was reduced upon CDK7 inhibition 84-86; phosphorylation of Thr806 in
SPT5 and also in the CTD of RNAPII were decreased after CDK9 inhibition 80,87,88, and modest
changes in phosphorylation of Ser2 and Ser7 were detected upon longer (≥ 1 h) CDK12/13
inhibition 78,81,89-95 (Fig. 5c, Supplementary Fig. 5a). In summary, our data demonstrate that
SF3B1 hyperphosphorylation induced by SNIP1 depletion is specifically mediated by CDK11.

Two P-Thrs in SF3B1 are required for interaction with SNIP1´s FHA domain
Given that SNIP1 harbors a candidate P-Thr-binding FHA domain at its C-terminus, we tested
whether this domain is necessary for interaction with P-SF3B1 (Fig. 6a). We used in vitro binding
assays (Fig. 3a) and incubated GST-tagged (P)-SF3B1 (1-463) with his-tagged wild-type or FHA-
deletion mutant (del FHA) SNIP1 proteins expressed in E. coli (Fig. 6a). As expected, wild-type
SNIP1 associated with P-SF3B1, while deletion of the FHA domain abolished this binding
(Supplementary Fig. 6a), confirming the essential role of the FHA domain in the interaction.

FHA domains typically consist of eleven ß-strands (ß1- ß11) interspaced with six loop regions
(L1-L6), and recognize a single P-Thr via residues on apical surfaces of the loops 44,45. We used
AlphaFold 3 (AF3) 96 to model the interaction between the SNIP1 FHA domain (amino acids 259-
371) (Fig. 6b) and P-SF3B1 (amino acids 142-436). The model predicted that the SNIP1 FHA
domain interacts with two P-Thr residues in P-SF3B1, P-Thr358 and P-Thr369. The P-Thr358 is
in close proximity to SNIP1 K301 in loop L3 and P-Thr369 interacts with SNIP1 K265 in loop L1
and R365 in loop L6 (Fig. 6c).

To determine the relevance of these amino acid residues for the interaction, we generated SNIP1
mutants K265G, K301G and R365G and tested their binding to P-SF3B1 in an in vitro binding
assay (Supplementary Fig. 6b). Both K265G and K301G mutations abolished the binding, and
R365G mutation reduced binding by approximately 3-fold (Supplementary Fig. 6b). SF3B1
contains 28 Thr/Pro repeats (Supplementary Fig. 6c) that can be phosphorylated 8,20, so we next
asked whether P-Thr358 and P-Thr369 are essential for SNIP1 binding or whether they can be
substituted by the remaining P-Thr. Therefore, we mutated these two amino acids to alanine
(Thr358A and Thr369A) in the GST-SF3B1 and tested their binding to wild-type SNIP1 in an in
vitro binding assay. Since the binding of SNIP1 to the wild-type and mutated (Thr358A and
Thr369A) SF3B1 proteins remained equal, this indicates that the other P-Thr residues in SF3B1
can compensate for the loss of P-Thr358 and P-Thr369 (Supplementary Fig. 6d). These findings
suggest that the SNIP1 FHA domain recognizes multiple P-Thr residues in SF3B1, with flexibility
in which sites are utilized.

Since cryo-EM structures of the activated spliceosome (Bact) containing P-SF3B1 and RES
complex were published 13,16 we wanted to determine whether association of P-SF3B1 with FHA
domain of SNIP1 can be visualized in these structures. Close examination of the Bact complexes
(PDB: 8I0R 16 and PDB: 5Z57 13) identified a contact point between SF3B1 and SNIP1 and non-
interpreted density in the loops of the FHA domain of SNIP1 (Fig. 6d). An AF3 model of SNIP1
FHA in complex with SF3B1-derived phospho-peptide motif (P)ThrPro[X] (P)ThrPro from the
repetitive sequence of 28 Thr/Pro (Supplementary Fig. 6c) demonstrated that the predicted motif
containing two P-Thr fits exactly to the region of the unassigned cryo-EM density (CCC=0.85 at
4 Å map resolution for the AF3 prediction of SNIP1 FHA domain in complex with the SF3B1
phospho-peptide) (Fig. 6e). This observation independently validates our in vitro mutational
analysis of SNIP1-loop mutants (K265G, K301G and R365G) and supports the findings that the
FHA domain of SNIP1 binds two P-Thr in P-SF3B1 to guide the recruitment and/or anchoring of
the RES complex during the spliceosome activation.

Neurodevelopmental disorder-associated SNIP1 mutation E366G reduces binding to P-SF3B1
A homozygous SNIP1 mutation (E366G) found in ~ 35 individuals from the Amish population
has been identified as the genetic cause of a neurodevelopmental disorder characterized by
hypotonia, craniofacial abnormalities, and seizures (NEDHCS) 47,48. However, the molecular basis
of this mutation´s effect remains unclear. As E366 is adjacent to R365 in loop L6 of the FHA
domain, a region critical for P-Thr recognition, we hypothesized that the E366G mutant could
impair SNIP1 binding to P-SF3B1. To test this, we compared the interactions of wild-type, E366G
and R365G SNIP1 proteins with P-SF3B1 in in vitro binding assays (Supplementary Fig. 6e).

The E366G mutant exhibited an approximately 3-fold reduced binding, similar to the R365G
mutation (Supplementary Fig. 6e). Close inspection of our SNIP1-P-SF3B1 interaction model
also indicates that E366 is in close proximity to the binding interface and its mutation to glycine
(E366G) induces most likely local conformational changes in the FHA domain leading to
suboptimal P-SF3B1 binding (Fig. 6e). Altogether, these findings strongly suggest that impaired
SNIP1-P-SF3B1 interaction is the likely molecular defect underlying NEDHCS.

SNIP1 mutations in the FHA domain affect cell viability and pre-mRNA splicing
To determine whether mutations in the SNIP1 FHA domain affect incorporation of SNIP1 into the
activated spliceosome, we expressed flag-tagged SNIP1 (F-SNIP1) WT, single mutants K265G,
K301G, R365G, E366G, and the double mutant K265G/K301G in cells and compared their
association with components of the activated spliceosome (Fig. 7a, b, Supplementary Fig. 7a).
Immunoprecipitation of endogenous BUD13, a component of the RES complex, revealed
interactions with P-SF3B1 and CWC22, but not with U1-70K and U2AF65, which associate with
the spliceosome at stages preceding activation 17 (Fig. 7a, b, Supplementary Fig. 7a). These
results indicate full integration of BUD13 into the activated spliceosome. Notably, BUD13
immunoprecipitated WT F-SNIP1 more efficiently than the F-SNIP1 mutants, indicating a reduced
ability of the mutants to associate with the activated spliceosome (Fig. 7a, b, Supplementary Fig.
7a). Because substitution with glycine (G) may disrupt the FHA domain conformation, we
additionally generated alanine (A) substitutions (K265A, K301A, E365A, and K265A/K301A).
These variants also showed reduced associations with the spliceosome in BUD13
immunoprecipitates compared with WT F-SNIP1 (Supplementary Fig. 7a), confirming the
importance of residues K265, K301, and E365 for mediating interaction with P-SF3B1 in cells.
Consistent with these findings, immunoprecipitation of F-SNIP1 E366G, K265G, and double
mutant K265G/K301G revealed moderate (for E366G) and severe (for K265G and
K265G/K301G) reductions in association with components of the activated spliceosome (P-
SF3B1, CWC22, RBM22 and BUD13) (Supplementary Fig. 7b, c).

To assess the physiological importance of these FHA domain residues for cell viability and pre-
mRNA splicing, we stably integrated F-SNIP1 constructs encoding WT or mutant SNIP1 into the
FRT/TO integration site of the HCT116 dTAG SNIP1 cell line. Immunoblot analysis showed that
F-SNIP1 K265G, E366G and double K265G/K301G mutants were expressed at levels comparable
to WT F-SNIP1 (Supplementary Fig. 7d, e); therefore, subsequent analyses were performed in
these cell lines.

Cells were treated with dTAG-v1 or DMSO, and cell viability was assessed either indirectly, by
measuring metabolic activity of mitochondrial dehydrogenases by WST assay (Fig. 7c), or directly
by crystal violet staining (Fig. 7d). In the absence of ectopic F-SNIP1 expression, cells were not
viable, consistent with the essential role of SNIP1 in constitutive splicing. Expression of WT F-
SNIP1 fully rescued both metabolic activity and cellular proliferation. In contrast, the K265G and
the double K265G/K301G mutants severely impaired metabolic activity and cell viability, whereas
the patient-derived E366G mutation caused only a moderate reduction of both (Fig. 7c, d).

We next examined whether these metabolic and viability phenotypes reflected defects in pre-
mRNA splicing (Fig. 7e, f, Supplementary Fig. 7f). Splicing of endogenous RIOK3, ARRDC1,
and CCNL1 transcripts was analyzed by RT-PCR following dTAG-v1 treatment. Accumulation of
unspliced transcripts was evident within 2 h of SNIP1 depletion in HCT116 dTAG-SNIP1 cells
(Fig. 7e, f). This splicing defect was partially rescued by expression of WT F-SNIP1, confirming
that the phenotype was specific to SNIP1 depletion (Fig. 7e, f). In contrast, expression of F-SNIP1
K265G, E366G, or the double K265G/K301G mutant failed to rescue, or rescued less efficiently,
the splicing defects compared with WT F-SNIP1 (Fig. 7e, f). Notably, the double K265G/K301G
mutant consistently showed a more severe defect than the patient-derived E366G mutation (Fig.
7e). These differences were further supported by quantitative PCR analysis of splicing of long
introns in the CCNL1 and SFSWAP genes (Supplementary Fig. 7f), indicating that the observed
splicing defects are general and not dependent on intron length. Taken together, these results
demonstrate that FHA domain residues mediating interaction with P-SF3B1 are essential for
efficient pre-mRNA splicing and cell viability.

Methods
Antibodies, Primers, Plasmids and Reagents
Lists of antibodies, primers, plasmids and reagents used in the study are provided in
Supplementary Data 3.

Cell culture
HEK293 Flp-in cell line (Thermo Fisher Scientific, R75007), human colorectal HCT116 cell line
(ATCC) and its variant HCT116 Flp-in cell line (a gift from B. G. Wouters), were maintained in
DMEM/high-glucose medium supplemented with L-glutamine, sodium pyruvate and 5% FBS at
37 °C and 5% CO . The cell lines were negatively tested for mycoplasma contamination with
2
MycoStrip Mycoplasma Detection Kit (InvivoGen, rep-mys-20).

To prepare HCT116 Flp-in cell line with inducible degradation of SNIP1 protein, we used
CRISPR-Cas9 genome editing with the PITCh system 119, and proceeded according to the
published protocol 120. sgRNA (CAGCCAAAATCACAGAATGA) targeting N-terminus of
SNIP1 gene was cloned into pX330-BbsI-PITCh (addgene #127875) 121 and microhomology arms
surrounding CRISPR-Cas9 cleavage site (each 20bp long) were introduced into donor plasmid
pCRIS-PITChv2-Puro-dTAG (addgene #91793) 71.

HCT116 Flp-in cells (seeded in 10 cm plates, 30% confluency) were co-transfected with 9 μg of
donor plasmid and 3 μg of sgRNA-Cas9 plasmid, and 24 μl Turbofect (Thermo Scientific, R0533)
in 2 ml Opti-MEM (Thermo Scientific, 11058-021). Medium was changed 2 h post-transfection.
Cells were transferred to 15 cm plates two days post-transfection and resistant colonies were
selected with 0.3 µg/ml puromycin (Merck, P8833) for at least 2 weeks. Individual colonies were
expanded in 12 well plates, genomic DNA was extracted with PureLink Genomic DNA Mini Kit
(Thermo Scientific, K182002), as well as protein lysates were prepared in RIPA buffer (50 mM
Tris-Cl, pH 8, 150 mM NaCl, 5 mM EDTA, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS,
supplemented with protease inhibitors, Sigma, P8340). Clones with homozygous integration of the
dTAG cassette at the SNIP1 locus were validated by Sanger sequencing and immunoblotting.

We selected clone (#28) with proper integration of dTAG cassette (named as HCT116 dTAG
SNIP1) for further experiments and verified rapid degradation of SNIP1 protein by treating the
cells with 500 nM dTAGv-1 (MedChemExpress, HY-145514).

We modified HCT116 dTAG SNIP1 cells by stable integration of flag-tagged SNIP1 (F-SNIP1)
gene into flp-in site. We prepared five derivative cell lines expressing different F-SNIP1 mutants
(WT, K265G, K265G&K301G and E366G). We verified both ectopic integration of F-SNIP1
variants by Sanger sequencing of their genomic DNA and F-SNIP1 expression by western blotting
with SNIP1 antibody (after induction with 1μg/ml doxycycline for 24 h).

Cell proliferation assays
All clones were seeded into four 96-well plate (400 cells / well; 6 wells per clone per plate) and
expression of F-SNIP1 variants was induced with doxycycline (1 mg/1 ml of media). The next day
dTAG-v1 (500 nM final concentration) or DMSO (solvent) were added and cells were cultured for
another 24, 48 and 72 hours, respectively. Each day cell growth was measured by WST assay
(ab65473, Abcam).

For crystal violet staining individual clones were seeded into 100 mm dishes (4 x 105 cells / dish) and
expression of F-SNIP1 variants was induced with doxycycline (1 mg/1 ml of media). The next
day dTAG-v1 (500 nM final concentration) or DMSO (solvent) were added and cells were cultured for
another 72 hours. Media was replaced every day and after three days the cells were fixed with
methanol: acetic acid (3:1) and stained with crystal violet.

Chromatin isolation and mass spectrometry analyses of SF3B1-associated proteins
Chromatin-associated protein complexes were isolated according to mNET-seq protocol 60.
Protein complexes were released from chromatin by mild MNase digestion, and finally
immunoprecipitated with endogenous antibodies. Immunoprecipitated proteins were detected by
mass spectrometry.

HCT116 cells were plated onto 15 cm plates (three plates per condition) to reach 80% confluency
at the day of experiment. Plates were washed twice with ice-cold PBS, scraped, pelleted 250 g at
4°C for 3 min and treated on ice for 5 min with 4 ml of ice-cold HLB+N buffer (10 mM Tris-HCl
(pH 7.5), 10 mM NaCl, 2.5 mM MgCl , 0.5% NP-40, 1x phosphatase inhibitor cocktail (Sigma,
2
P0044) and 1x protease inhibitor cocktail (Sigma, P8340)). The cell pellet was underlaid with 1
ml of ice-cold HLB+NS (10 mM Tris-HCl (pH 7.5), 10 mM NaCl, 2.5 mM MgCl , 0.5% NP-40,
2
10% sucrose, 1x phosphatase inhibitor cocktail and 1x protease inhibitor cocktail). Nuclei were
pelleted by centrifugation (420 g, 4 °C for 5 min) and supernatant was discarded. Pellets were
resuspended by vortexing in 100 μl NUN1 buffer containing 20 mM Tris-HCL (pH = 7.5), 75 mM
NaCl, 0.5 mM EGTA, 50 % glycerol, 1x protease and 1x phosphatase inhibitors. Chromatin was
extracted by addition of 1 ml of NUN2 buffer containing 20 mM HEPES (pH = 7.6), 7.5 mM
MgCl , 0.2 mM EGTA, 300 mM NaCl, 1 M urea and 1 % NP-40, 1x protease and 1x phosphatase
2
inhibitors and rotated at 4°C for 10 min. After centrifugation (1600 g, 4°C for 5 min), supernatant
was completely removed. Chromatin pellet was washed once with 1 ml of MNase buffer, digested
with MNase at 37 °C for 6 min at 1400 rpm. MNase was inactivated by EGTA (20 mM final
concentration). After centrifugation (1600 g, 4°C for 5 min), supernatant was diluted by two
volumes of NET-2 buffer (50 mM Tris-HCl (pH=7.5), 150 mM NaCl, 0.05% NP-40, 1x protease
and 1x phosphatase inhibitors) and pre-cleared by protein G Dynabeads at 4 °C for 1 h.
Immunoprecipitation was performed with SF3B1 antibody (MBL, MB-221-3, 5 μl).

Immunoprecipitated samples were washed four times with 1 ml of NET-2 buffer and stored in 10
mM Tris-HCl (pH=8.1) at -80 °C.

Beads-bound protein complexes were mixed with 2 % SDS solution and heated to 50 ℃ for 20 min
following protein reduction using 100 mM DITT at 95 ℃ for 15 min. Resulting protein solutions
were subjected to filter-aided sample preparation (FASP) as described elsewhere 122 covering
proteins alkylation (100 mM iodoacetamide (IAA) for 20 min in dark at laboratory temperature)
and proteins digestion using 0.75 μg of trypsin (sequencing grade; Promega). Peptides were then
cleaned by liquid-liquid extraction (3 iterations) using water saturated ethyl acetate 123. Cleaned
FASP eluates were evaporated completely in SpeedVac concentrator (Thermo Fisher Scientific)
and transferred into LC-MS vial containing PEG (final concentration 0.001%, v/v) using 25 μl of
50% acetonitrile (ACN) with 2.5% formic acid (FA) and 100 μl of ACN. SpeedVac was used for
transferred peptides concentration and ACN removal with final volume adjustment using MilliQ
water. Resulting peptide solutions were analyzed by liquid chromatography–tandem mass
spectrometry (LC–MS/MS) performed using UltiMate 3000 RSLCnano system (Thermo Fisher
Scientific) on-line coupled with Orbitrap Fusion Lumos Tribrid mass spectrometer (Thermo Fisher
Scientific).

LC-MS/MS analyses
LC-MS/MS analyses were done using Ultimate 3000 RSLCnano liquid chromatograph connected
to Orbitrap Fusion Lumos Tribrid mass spectrometer (Thermo Fisher Scientific). Prior to LC
separation, tryptic digests were online concentrated and desalted using trapping column
(Acclaim™ PepMap™ 100 C18, dimensions 300 μm ID, 5 mm long, 5 μm particles, Thermo
Fisher Scientific). After washing of trapping column with 0.1% FA, the peptides were eluted (flow
rate - 300 nl/min) from the trapping column onto an analytical column (Aurora C18, 75 μm ID,
250 mm long, 1.6 μm particles, Ion Opticks) by 101 min long linear gradient (2-37% of mobile
phase B; mobile phase A: 0.1% FA in water; mobile phase B: 0.1% FA in 80% ACN).
Equilibration of the trapping column and the analytical column was done prior the sample injection
to sample loop. The analytical column outlet was directly connected to the Digital PicoView 550
(New Objective) ion source with sheath gas option. ABIRD (Active Background Ion Reduction
Device, ESI Source Solutions) was installed.

MS data were acquired in a data-dependent strategy (cycle time 2.5 s). Survey scan range was set
to m/z 350-2000 with the resolution of 120,000 (at m/z 200), normalized target value of 250 %
and maximum injection time of 500 ms. HCD MS/MS spectra (isolation window m/z 1.2, 30%
relative fragmentation energy) were acquired from 110 m/z with a target value of 5×104, resolution
of 15,000 (at m/z 200) and maximum injection time of 50 ms. Dynamic exclusion was enabled for
30 s.

The analysis of the mass spectrometric RAW data files was carried out using the MaxQuant
software (version 2.0.1.0) 124 using default settings unless otherwise noted. MS/MS ion searches
were done against modified cRAP database (based on http://www.thegpm.org/crap) containing
protein contaminants like keratin, trypsin etc., and UniProtKB protein database for Homo sapiens
(downloaded 2021-06-16, number of protein sequences: 20,600). Oxidation of methionine,
deamidation (N, Q) and acetylation (protein N-terminus) as optional modification,
carbamidomethylation (C) as a fixed modification and trypsin/P enzyme with two allowed miss
cleavages were set. Peptides and proteins with FDR threshold <0.01 and proteins having at least
one unique or razor peptide were considered only. Match between runs was set among all analyzed
samples. Protein abundance was assessed using protein intensities calculated by MaxQuant.

Reported protein intensities were further processed using the software container environment
(https://github.com/OmicsWorkflows). Processing workflow is available upon request. Briefly, it
covered: a) removal of decoy hits and contaminant protein groups, b) protein group intensities log2
transformation, c) loessF normalization, d) quantitative filtering and global minimum imputation,
e) differential expression using LIMMA statistical test.

Isolation and analyses of chromatin-associated proteins by immunoblotting
This protocol was used to analyze chromatin-associated proteins by western blotting without
immunoprecipitation step. Cell pellets (approximately 20-30 µl of packed cell pellet) were re-
suspended in 1 ml buffer containing 20 mM HEPES (pH = 7.5), 10 mM KCl, 250 mM sucrose, 5
mM MgCl , 1 mM EGTA, 1x phosphatase inhibitor cocktail (Sigma, P0044) and 1x protease
2
inhibitor cocktail (Sigma, P8340), and lysed by addition of digitonin (Sigma, D141), 200 µg/ml
final concentration at 4°C for 10 min. Nuclei were pelleted by centrifugation (650 g, 5 min),
supernatant was discarded and the pellet was resuspended in 100 μl NUN1 buffer containing 20
mM Tris-HCL (pH = 7.5), 75 mM NaCl, 0.5 mM EGTA, 50 % glycerol, 1x protease and 1x
phosphatase inhibitors. Chromatin was extracted by the addition of 1 ml of NUN2 buffer
containing 20 mM HEPES (pH = 7.6), 7.5 mM MgCl , 0.2 mM EGTA, 300 mM NaCl, 1 M urea
2
and 1 % NP-40, 1x protease and 1x phosphatase inhibitors and rotated at 4°C for 10 min. After
centrifugation (1600 g, 4°C for 5 min), supernatant was completely removed. Chromatin was
diluted (without touching the pellet) in 200 µl of RIPA buffer (50 mM Tris-Cl (pH = 8), 150 mM
NaCl, 5 mM EDTA, 1 % NP-40, 0.5 % sodium deoxycholate, 0.1% SDS, 1x protease and 1x
phosphatase inhibitors, followed by addition of 100 µl of 3×Laemmli sample buffer. Samples were
kept at room temperature for 30 min with occasional shaking, sonicated 30 times for 1 s (amplitude
0.30) using a 5/64 probe (QSonica Q55A) and boiled for 15 min at 95°C. Protein levels were
analyzed by SDS-PAGE. The antibodies used for protein detection and their dilutions are listed in
Supplementary Data 3. Uncropped and unprocessed scans of all immunoblots are available in
the Source Data file.

iCLIP–seq
iCLIP was performed according to the previously published protocol 8. Protein-RNA complexes
were immunoprecipitated with 15 μl of affinity purified rabbit antisera against P-Thr211-SF3B1
or P-Thr235-SF3B1 8. Multiplexed libraries were sequenced as 50 bp single-end reads on the
Illumina NextSeq system (VBCF).

Preparation of whole cell lysates and analysis by western blotting
HCT116 dTAG-SNIP1 cells were plated onto 10 cm plates to reach 50% confluency at the day of
experiment. Plates were treated with 500 nM dTAG-V1, 50 nM OTS964 or DMSO (as a solvent
control) for the indicated time. The plates were then washed twice with ice-cold PBS, scraped,
pelleted 250 g at 4°C for 3 min and lysed with RIPA buffer (50 mM Tris-Cl (pH = 8), 150 mM
NaCl, 5 mM EDTA, 1 % NP-40, 0.5 % sodium deoxycholate, 0.1% SDS, 1x protease and 1x
phosphatase inhibitors). During lysis samples were sonicated three times for 1 s (amplitude 0.30)
using a 5/64 probe (QSonica Q55A), cleared by centrifugation (10,000 g at 4 °C for 10 min) and
protein concentration was equalized between samples. For western blot analysis samples were
separated by SDS-PAGE and transferred to 0.45 μm nitrocellulose membranes. The membranes
were blocked for 1 h in 5% low-fat milk in PBS-0.05% Tween 20 (PBS-T) and incubated overnight
with primary antibodies at 4 °C in 5% low-fat milk in PBS-T. Next day membranes were washed
3× 10 min in PBS-T and incubated for 1 h in 5% low-fat milk in PBS-T with appropriate secondary
antibody. Membranes were washed 3 ×10 min in PBS-RT and visualized using Western Blotting
Luminol Reagent (Santa Cruz, sc-2048) and/or Clarity™ Western ECL Substrate (Bio-Rad). The
antibodies used for protein detection and their dilutions are listed in Supplementary Data 3.

Co-immunoprecipitation assay
HCT116 dTAG SNIP1 cells and its derivative stable cell lines expressing F-SNIP1 variants (WT,
K265G, K265G&K301G, E366G) were plated onto 15 cm plates to reach 90% confluency at the
day of experiment. Expression of F-SNIP1 variants was induced with doxycycline (1 mg/1 ml of
media) 18 h prior harvesting cells. Plates were washed twice with ice-cold PBS, scraped, pelleted
250 g at 4°C for 3 min. Cell pellets were lysed with 800 μl of buffer C (20 mM Tris-Cl (pH = 8.0),
150 mM NaCl, 1.5mM MgCl2, 10mM KCl, 10% Glycerol, 0.5 mM EDTA, 0.5 % NP-40, 1x
protease and 1x phosphatase inhibitors). During lysis samples were sonicated 10 x 1 s (amplitude
0.30) using a 5/64 probe (QSonica Q55A), cleared by centrifugation (10,000 g for 10 min at 4 °C)
and protein concentration was equalized between samples. Immunoprecipitation was performed
with Flag agarose M2 affinity beads (Sigma-Aldrich, A2220, 15 μl per sample) or with BUD13
antibody (A303-321A) prebound to protein G Dynabeads beads (20 μl per sample).
Immunoprecipitated samples were washed three times with 1 ml of Buffer C. After the last wash,
all liquid was carefully removed, the beads were resuspended in 35 μl of 1x Laemmli buffer and
boiled for 1 min. For western blot analysis samples were separated by SDS-PAGE and transferred
to 0.45 μm nitrocellulose membranes. The antibodies used for protein detection and their dilutions
are listed in Supplementary Data 3.

RNA-seq
HCT116 dTAG-SNIP1 cells were plated onto 10 cm plates to reach 50 % confluency at the day of
experiment. Next day cells were treated with 500 nM dTAG-V1 for 4 h or with DMSO (as a solvent
control). Plates were washed twice with ice-cold PBS, scraped, pelleted 250 g at 4°C for 3 min
and treated for 5 min with 500 µl of cytoplasmic lysis buffer (10 mM Tris-Cl, pH 8, 0.32 M
Sucrose, 3 mM CaCl , 2 mM MgCl , 0.1 mM EDTA, 0.5 % TritonX-100) supplemented with 20
2 2
U/ml SUPERase inhibitor (Thermo Scientific, AM2694). Supernatant was removed by
centrifugation (650 g at 4°C for 3 min). The pellet was washed once more with cytoplasmic lysis
buffer. Remaining nuclear pellet was carefully resuspended in 1 ml Tri-Reagent (Sigma, 9424)
and RNA was isolated according to the manufacturer protocol. 250 ng of DNase treated RNA
(Sigma, AMPD1) was used for rRNA depletion using RiboCop rRNA Depletion Kit V1.2
(Lexogen, #144). Sequencing libraries were prepared using the NEBNext UltraExpress RNA
Library Prep Kit for Illumina (NEB, E3330) according to the manufacturer protocol and sequenced
on DNBSEQ-G400, MGI (150 bp paired-end, minimum 100 mil reads per condition, 2 biological
replicates).

RT-PCR analysis
HCT116 dTAG-SNIP1 cells and its derivative stable cell lines expressing F-SNIP1 variants (WT,
K265G, K265GK301G and E366G) were treated for 2 h with DMSO or 500 nM dTAG-V1.
Nuclear RNA was extracted using Tri-Reagent (9424, Sigma), as described in the RNA-seq
section. After DNase treatment (AMPD1, Sigma), 1,000 ng of RNA was reverse-transcribed using
random hexamers and Superscript II reverse transcriptase (Thermo Scientific). PCR was
performed in 20 μl reaction mixture containing 5 ng of the obtained cDNA (or 50 ng of human
genomic DNA as a control), 1x Phusion HF buffer, 0.2 mM dNSTP mix, 0.5 μM of each primer and
0.4 U Phusion DNA polymerase (F530S, Thermo Scientific). PCR conditions were 94 °C for 30
s; 32 cycles of 94 °C for 10 s, 62 °C for 20 s, and 72 °C for 30 s; followed by 72 °C for 10 min.
Primer sequences are specified in Supplementary Data 3. PCR products were separated on a
1.5% agarose gel and stained with Midori Green (MG03, Nippon Genetics). Band intensities were
analyzed by ImageJ software (v.1.54f2).

Purification of His-SNIP1 proteins
The 6×His-SNIP1 protein fragments (WT (1-396), del FHA (241-371), K265G, K301G, E366G,
R365G, SSR363GAG) were expressed in BL21 RIPL E. coli strain using pET-M11 plasmids.
Cultures were induced at OD600 = 0.6 by 0.5 mM IPTG (Sigma-Aldrich, I5502) and kept at 16
°C for 16 h. Cells were collected by centrifugation at 5,000 g at 4 °C for 10 min. Cell extracts were
prepared by sonication in 10 mL of lysis buffer (50 mM NaH PO , pH 8.0, 500 mM NaCl, 10 mM
2 4
imidazole, 1% Triton X-100, 1 mM PMSF). Lysates were clarified by centrifugation, and the
resulting supernatant was incubated with 1 mL of Ni-NTA Agarose (Qiagen, 30210) for 1 h at 4
°C. The beads were washed with 40 ml of wash buffer (50 mM NaH PO , pH 8.0, 500 mM NaCl,
2 4
35 mM imidazole, 10% glycerol, 1% TritonX-100). SNIP1 proteins were eluted 5 times with 300
μl of elution buffer (50 mM NaH PO , pH 8.0, 500 mM NaCl, 250 mM imidazole, 10 % glycerol,
2 4
1 % Triton X-100). Fractions 2-5 were pooled together and stored in aliquots at –80 °C until use.

Purification of GST-SF3B1 proteins
The GST-SF3B1(1–463) and GST-SF3B1(1–463) T358A T369A were expressed in BL21 RIPL
E. coli strain using the pGEX plasmid. Cultures were induced at OD600 = 0.6 by 0.5 mM IPTG
(Sigma-Aldrich, I5502) and kept at 16 °C for 16 h. Cells were collected by centrifugation at 5,000
g at 4 °C for 10 min. Cell extracts were prepared by sonication in bacteria lysis buffer (50 mM
Tris, 20% glycerol, 150 mM NaCl, 0.5 mM EDT A, 1 % NP-40, 2 mM PMSF, cOmplete Mini
Protease Inhibitor Cocktail (1 tablet per 50 ml, Roche, 04693124001)). Lysates were clarified by
centrifugation at 10,000g at 4 °C for 10 min, and the resulting supernatant was incubated with 1
ml of Glutathione Sepharose 4B beads (Cytiva, GE17- 0756-01) for 1 h at 4 °C. The beads were
washed with 10 ml of bacteria lysis buffer followed by 40 ml of wash buffer (50 mM Tris, 20 %
glycerol, 200 mM NaCl, 0.5 mM EDTA). GST-tagged SF3B1(1–463) was eluted eight times with
750 µl of elution buffer (50 mM Tris, 20 % glycerol, 150 mM NaCl, 0.5 mM EDTA, 10 mM
glutathione). Fractions containing SF3B1 were pooled together, washed with PBS and
concentrated in an Amicon Ultra-4 Centrifugal Filter Unit (Sigma-Aldrich, UFC801024). The
samples were stored at −80 °C until use.

F-CDK11 kinase purification from HEK293 cells
HEK293 cells were plated onto 150 cm2 plates to reach 40 % confluency on the day of the
transfection. The cells were co-transfected with flag-tagged CDK11 WT 125/Xpress-tagged cyclin
L1 125/HA-tagged SAP30BP (10 μg/5 μg /5 μg) using PEI transfection reagent. Media were
changed 2 h post-transfection. The cells were collected 48 h post-transfection, washed twice with
PBS and lysed in 1 ml of HEPES buffer (20 mM HEPES-KOH (pH 7.9), 300 mM KCl, 0.2 % NP-
40, 15 % glycerol, 1 mM DTT and protease inhibitors, Sigma-Aldrich, P8340) for 20 min. During
the lysis, protein extracts were mildly sonicated (10 times for 1 s, amp 0.30, using 5/64 probe,
QSonica Q55A). Clarified extracts (10,000g for 10 min at 4 °C) were subsequently rotated with
15 μl of packed Flag agarose M2 affinity beads (Sigma-Aldrich, A2220) for 1 h at 4 °C. The beads
were washed three times with 1 ml of high-salt buffer (20 mM HEPES-KOH (pH 7.9), 500 mM
KCl, 0.2 % NP-40, 15 % glycerol, 1 mM DTT) with rotation for 5 min at 4 °C during each wash,
followed by one detergent-free wash (20 mM HEPES-KOH (pH 7.9), 150 mM KCl, 15% glycerol,
1 mM DTT). The remaining buffer was carefully removed and 3×flag-tagged proteins were eluted
using 90 μl of flag elution buffer (20 mM HEPES-KOH (pH 7.9), 150 mM KCl, 1 mM DTT and
1 mg/ml 3×flag peptide, Sigma-Aldrich, F4799).

Isolation of soluble nuclear lysate from HCT116 cells
HCT116 cells were plated onto 15 cm plates to reach 80 % confluency. Plates were washed twice
with ice-cold PBS, scraped, pelleted 250 g at 4°C for 3 min and treated on ice for 5 min with 500
µl of cytoplasmic lysis buffer (10 mM Tris-Cl, pH 8, 0.32 M Sucrose, 3 mM CaCl , 2 mM MgCl ,
2 2
0.1 mM EDTA, 0.5 % TritonX-100). Supernatant was removed by centrifugation (650 g at 4°C
for 3 min). The pellet was once more washed with cytoplasmic lysis buffer and once with
cytoplasmic lysis buffer without detergent. Nuclear pellet was that lysed with 300 μl EDTA-EGTA
buffer (3 mM EDTA, 0.2 mM EGTA, 450 mM KCl, 1mM DTT) and rotated for 30 min at 4°C.
Chromatin pellet was removed by centrifugation (1700 g at 4 °C for 10 min), supernatant (soluble
nuclear lysate) was saved and treated with 1.5 μl of RNase A (Qiagen, 19101) at room temperature
for 15 min. Finally, soluble nuclear lysate was diluted with two volumes of the dilution buffer (60
mM HEPES-KOH (pH = 7.9), 1 mM DTT, 30 % glycerol, 0.4 % NP-40, 1 x phosphatase inhibitor
cocktail (Sigma, P0044) and 1 x protease inhibitor cocktail (Sigma, P8340)) to lower KCL
concentration to 150 mM. The lysate was kept on ice until needed in the SF3B1 pull down assay.
Usually, lysate from one 15 cm plate was added to one in-vitro kinase reaction.

SF3B1 pull down assay coupled to mass spectrometry analysis
1 μg of purified GST-tagged SF3B1(1–463) in 48 μl of kinase buffer (20 mM HEPES-KOH
pH=7.9, 5 mM MgCl , 2 mM DTT, 2 mM ATP, 1x phosphatase inhibitor cocktail (Sigma, P0044))
2
was incubated with 12 µl flag-tagged CDK11 kinase (immunoprecipitated from HEK293 cells) or
with 12 µl of elution buffer (20 mM HEPES-KOH pH=7.9, 150 mM KCl, 1 mM DTT). After 60
min shaking at 450 rpm at 30 °C, 20 μl of 3× washed glutathione Sepharose 4B resin (GE
Healthcare, 17075601) was added to each sample and they were rotated for another 60 min at 4
°C. After 60 min incubation the samples were washed 3x with 1ml of binding buffer (20mM
HEPES-KOH (pH = 7.9), 150 mM KCl, 0.2 mM EDTA, 1 mM DTT 15 % glycerol, 0.2 % NP-
40), mixed with 800 μl of RNase-treated soluble nuclear lysate prepared from HCT116 cells (see
protocol above) and rotated for 60 min at 4°C. Finally, the beads were washed 5 times with 1 ml
of modified binding buffer (20 mM HEPES-KOH (pH = 7.9), 200 mM KCl, 0.2 mM EDTA, 1
mM DTT, 15 % glycerol, 0.2 % NP-40) and stored in 10 mM Tris-HCl (pH=8.1) at -80 °C.
Samples used for western blot verification were prepared similarly, after final washing step the
beads were diluted in 1x Laemmli buffer and boiled for 3 min. After SDS–PAGE, the following
antibodies were used: CDK11 (rabbit antiserum, 1:3000), P-T235-SF3B1 (rabbit antiserum,
1:3000), GST (Santa Cruz, sc-138, 1:200), U2AF65 (Santa Cruz, sc-53942, 1:400), RBM17
(Sigma, HPA037478, 1:1000). For detection of SNIP1 (Thermo Scientific, PA5-29412, 1:1000)
and BUD13 (Bethyl, A303-321A, 1:2000) proteins by western blotting, the SF3B1 pull down
protocol was modified in a way that six in-vitro kinase reactions were pooled together for each
condition (6 times with and 6x without flag-tagged CDK11 kinase). Beads-bound protein
complexes were processed as described in section Chromatin isolation and mass spectrometry
analyses of SF3B1-associated proteins.

LC-MS/MS analyses of all peptides were done using UltiMate 3000 RSLCnano liquid
chromatograph (Thermo Fisher Scientific) connected to timsTOF Pro spectrometer (Bruker). Prior
to LC separation, tryptic digests were online concentrated and desalted using trapping column
(Acclaim™ PepMap™ 100 C18, dimensions 300 μm ID, 5 mm long, 5 μm particles, Thermo
Fisher Scientific). After washing of the trapping column with 0.1% trifluoroacetic acid (TFA), the
peptides were eluted (flow rate - 300 nl/min) from the trapping column onto an analytical column
(Aurora C18, 75μm ID, 250 mm long, 1.7 μm particles, heated to 50°C, Ion Opticks) by 60 min
linear gradient program (3-42 % of mobile phase B; mobile phase A: 0.1 % FA in water; mobile
phase B: 0.1 % FA in 80 % ACN). Equilibration of the trapping column and the analytical column
was done prior to sample injection to sample loop. The analytical column was placed inside the
Column Toaster (Bruker) and its emitter side was installed inside the CaptiveSpray ion source
(Bruker) according to the manufacturer instructions with the column temperature set to 50 ºC.
MSn data were acquired in data independent acquisition (DIA) mode with base method m/z range
of 100-1700 and 1/k0 range of 0.6-1.4 V×s×cm-2. Precursor range was defined to m/z 400-1000
with equal windows sizes of 26 Th (including 1 Th overlap) using two steps each PASEF scan and
cycle time of 100ms locked to 100% duty cycle.

DiaPASEF data were processed in DIA-NN4 (version 1.8.1) 126 in library free mode against
modified cRAP database (based on http://www.thegpm.org/crap; 112 sequences in total) and and
UniProtKB protein database for Homo sapiens (downloaded 2024-01-24, number of protein
sequences: 20,597). No variable, carbamidomethylation as fixed modification and trypsin/P
enzyme with 1 allowed missed cleavages were set during the library preparation. False discovery
rate (FDR) control was set to 1% FDR. MS1 and MS2 accuracies as well as scan window
parameters were set based on the initial test searches (median value from all samples ascertained
parameter values). MBR was switched on.

Reported protein intensities were further processed using the software container environment
(https://github.com/OmicsWorkflows). Processing workflow is available upon request. Briefly, it
covered: a) removal of low-quality precursors and contaminant protein groups, b) precursor
intensities normalization by loessF algorithm c) protein group MaxLFQ intensities calculation and
log2 transformation, and d) differential expression analysis using LIMMA statistical test.

SF3B1 in-vitro binding assays
1 μg of purified GST-tagged SF3B1(1–463) in 48 μl of kinase buffer (20 mM Tris-HCl, pH=8.0,
5 mM MgCl , 2 mM DTT, 2 mM ATP) was incubated with 12 ul flag-tagged CDK11 kinase
2
(immunoprecipitated from HEK293 cells) or with 12 ul of elution buffer (20 mM Tris-HCl
(pH=7.5), 150 mM NaCl, 1 mM DTT). After 60 min shaking at 450 rpm at 30 °C, 20 μl of 3×
washed glutathione magnetic agarose beads (Thermo Scientific, 78601) were added to each sample
and they were rotated for another 60 min at 4 °C. After 60 min incubation the samples were washed
2x with 1 ml of binding buffer (125 mM Tris-HCl (pH = 8.0), 150 mM NaCl, 1 mM EDTA, 1 mM
DTT, 0.1% Tween 20), mixed with 1 ug of SNIP1 peptide in 300 ul of binding buffer and rotated
for 60 min at 4°C. Finally, the beads were washed 3 times with 1 ml of modified binding buffer
(125 mM Tris-HCl (pH = 8.0), 250 mM NaCl, 1 mM EDTA, 1 mM DTT, 0.1% Tween 20).
Samples used for western blot analysis were diluted in 1x Laemmli buffer and boiled for 3 min.
After SDS–PAGE, the following antibodies were used: CDK11 (rabbit antiserum, 1:3000), P-
T235-SF3B1 (rabbit antiserum, 1:3000), GST (Santa Cruz, sc-138, 1:200), SNIP1 (Proteintech,
14950-1-AP).

Major Isoform Selection and Splicing Ratio Analysis
The primary transcript isoforms were determined following established protocols 73. RNA-
sequencing data was processed using the NF-Core pipeline 127 (Nextflow v23.04.1) from the NF-
Core consortium (v3.14, available at https://github.com/nf-core/rnaseq; archived at
https://zenodo.org/records/10471647). To identify the predominant isoform for each gene,
sequencing reads were mapped with Salmon 128 (v1.10.1) against curated UCSC RefSeq
annotations (GRCh38, downloaded July 2024). The isoform exhibiting the highest mean
transcript-per-million (TPM) value across all samples was designated as the major isoform,
provided it accounted for at least 70% of the gene’s isoform expression. Isoforms associated with
overlapping genes and those located on chromosomes X, Y, and M were excluded.

Splicing ratios were computed via an exon-centric splice junction analysis 58. Only exons from
major isoforms with an initial exon exceeding 100 base pairs were considered, while isoforms
annotated as single-exon were excluded. A window spanning ±4 base pairs around each splice
junction (2 bp in the exon and 2 bp in the intron) was defined to assess both spliced and unspliced
reads using the findOverlaps function from the GenomicRanges 129 R package (v1.56). Reads
overlapping at least 3 bp of this window were classified as unspliced, whereas reads overlapping
at least 2 bp were counted as total reads (spliced and unspliced). Spliced reads were determined
by subtracting the unspliced count from the total. The splicing ratio was calculated as the
proportion of spliced reads relative to the total, spliced and unspliced reads 73.

iCLIP Data Analysis
iCLIP sequencing data, including total, P-Thr211- and P-Thr235 SF3B1 samples, were analyzed
on the flow.bio platform (https://app.flow.bio/) using the nf-core/clipseq 130 pipeline (v1.0;
https://github.com/nf-core/clipseq/tree/1.0.0). Initial processing involved removing experimental
barcodes (iCount version 2.0.0, de-multiplex function https://github.com/tomazc/iCount),
followed by read alignment with STAR 131 against the human reference genome (hg38,
GRCh38.p13 primary assembly, GENCODE version 39), allowing up to two mismatches. Unique
molecular identifiers (UMIs) were applied to identify and eliminate PCR duplicates. Cross-linking
events were determined by identifying the nucleotide immediately preceding each uniquely
mapped sequencing read.

Genome-wide coverage tracks for combined iCLIP replicates were generated using deepTools 132,
normalized by sequencing depth and total read count, and binned per nucleotide. cDNA counts at
individual iCLIP cross-link sites were normalized to the total number of reads in each experimental
library and scaled to counts per million (CPM). Metagene density plots were generated from
normalized coverage data across all RNUs. To account for variations in region lengths between
main and pseudogene RNUs, the mode distance was used to scale the signal for each RNU class
as follows: RNU1 (116 genes, 163 nt), RNU2 (39 genes, 190 nt), RNU4 (54 genes, 140 nt), RNU5
(25 genes, 115 nt), and RNU6 (505 genes, 106 nt).

Data Availability
All next-generation-sequencing raw and processed data are available at the NCBI’s Gene
Expression Omnibus under the GEO series accession numbers GSE295433
[https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE295433] (iCLIP-seq) and
GSE295435 [https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE295435] (RNA-seq). All
proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner
repository with the dataset identifier PXD063160
[http://proteomecentral.proteomexchange.org/cgi/GetDataset?ID=PXD063160] and PXD063162
[http://proteomecentral.proteomexchange.org/cgi/GetDataset?ID=PXD063162]. Source data are
provided with this paper.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this study, the authors sought to explain a requirement for SF3B1 phosphorylation by cyclin-dependent kinase 11 (CDK11) in pre-mRNA splicing, which they established in their previous work (ref. 8). Here they use mass spectrometry and iCLIP to analyze protein-protein and protein-RNA interactions, respectively, of SF3B1 in human colon cancer cells in the absence or presence of the selective CDK11 inhibitor OTS964. They uncover a novel intermediate in spliceosome assembly/ activation, designated as “BOTS964”, which is blocked after association with U4 and U6 snRNPs but prior to recruitment of the RES complex and subsequent enzyme activation. They show that the RES complex subunit SNIP1 is needed for this interaction—and to support normal splicing—by introducing a degron tag (dTAG) into SNIP1 in HCT116 cells; and that the forkhead-associated (FHA) domain of SNIP1 is necessary for phosphorylation-dependent binding to the intrinsically disordered region (IDR) of SF3B1 in pulldown assays in vitro. Finally, the authors present evidence that a SNIP1-FHA mutant variant implicated in a neurodevelopmental disorder is defective in binding phosphorylated SF3B1 (P-SF3B1) in vitro. Overall, this is a well-designed and –executed study. Most of the major conclusions are well supported by the data, and will be of high interest to the readers of Nature Communications. There are, however, some deficiencies—most significantly, a missed opportunity to test effects of SNIP1-FHA mutations on splicing in living cells—that should be addressed in a revised manuscript.\nSpecific concerns:\n1. Line 2: The title is generally a bit acronym-heavy, and in particular I’m afraid readers outside the field will not know what “RES” is. I would recommend giving them at least a clue by saying “RES complex” (minor).\n2. In Figure 1c, there are a couple of extreme outliers whose association with SF3B1 drops more drastically than the others in the same complexes (NTC and IBC). What are these (they should probably be labeled)? Are they known to be more loosely associated with their respective complexes, or more labile?\n3. Lines 172-173: Looking at the correlation analysis in Fig S2b, it is an overstatement to say that the iCLIP replicates for the two anti-P-SF3B1 antibodies “correlated well” (Pearson correlation coefficients for the DMSO treated samples are mostly around 0.5, whereas the OTS964-treated samples show lower correlation, presumably because of lower yield). Compare this to the strong correlations reported in ref. 8 for the total SF3B1 antibody (ED Fig. 9d).\n4. Lines 186-189: I am not sure I follow the logic that leads the authors to state that the reduction of P-SF3B1 binding to the U6 internal stem-loop (ISL) “is likely an underestimate.” As I understand the normalization scheme, the OTS964-treated sample was overrepresented (in terms of cell equivalents) to account for decreased protein-RNA binding, but shouldn’t this boost both ISL and non-ISL read counts equally? It is the relative enrichment of ISL in the two conditions (DMSO vs. OTS964) that is being measured here, so I’m not really sure what is being underestimated.\n5. Relatedly, I was unable to deduce from the data shown what the quantitative effect of CDK11 inhibition on U6 binding was (as opposed to the qualitative effect on sequence specificity depicted in Fig. 2). Can this be estimated based on the iCLIP data with the pan-specific anti-SF3B1 antibody?\n6. Lines 213-216: The enrichment of CDK11 and its binding partners (cyclin L, SAP30BP) in the pulldown of P-SF3B1 seems likely to be a trivial consequence of sample preparation differences, i.e., pre-treating only this sample with purified, FLAG-tagged CDK11 complexes. Concluding anything about CDK11’s relative affinities for P-SF3B1 vs. SF3B1 would require a better-controlled experiment, e.g. -/+ ATP, or the use of a catalytically dead mutant CDK11 to generate the non-phosphorylated sample. The authors should probably point this out limitation.\n7. Lines 256-258: Here is the missed opportunity. Having established the dTAG-SNIP1 system and achieving rescue of splicing defects by ectopic SNIP expression (albeit partial, likely because of low expression), I would like to have seen tests of the various mutant forms of SNIP that abolish or impair binding in vitro to P-SF3B1—K265G, K301G. R365G, E366G—for their ability to rescue.\n8. Relatedly, Gly substitutions are notoriously disruptive of protein conformation; is there a rationale behind changing basic residues to Gly rather than, say, Ala (or acidic residues)?\n9. Lines 291-298 and Fig. 5c: The intent here was to determine the specificity (or exclusivity) of SF3B1 as a CDK11 substrate, given previous reports that other transcriptional CDKs phosphorylate this protein; the conclusion that an OTS964-sensitive kinase is required for the bulk of SF3B1 phosphorylation in cells is probably correct, but the analysis is both incomplete and flawed. Among the flaws or omissions: 1) although CDK7 is named as a candidate SF3B1 kinase (based on ref. 70), no CDK7-selective inhibitor is tested; 2) the CDK9 inhibitor NVP-2 is tested at a very low dose (30 nM) and with a substrate (RNAPII CTD) that is probably not the best (SPT5 is more sensitive and specific: see, for example: doi: 10.1038/s41467-020-18173-6); 3) even so, NVP-2 treatment does decrease the upshifted (i.e., phosphorylated) SF3B1 detected with the pan-specific antibody at the 3-hour time point, but this is not commented upon; 4) the “slightly increased” phospho-Ser2 signal seen after treatment with the CDK12/13 inhibitor THZ531 does not agree with at least one previous study using this compound, which showed a decrease (doi: 10.1038/nchembio.2166); and 5) none of the effects are quantified.\n10. Lines 380-382 and Fig. 7b: In both the cartoon model and the indicated text, the authors seem to imply that the phosphorylated residues of the SF3B1 IDR are interacting directly with the ISL of U6 snRNA, but, besides being electrostatically unfavorable (as the authors seem to acknowledge), this is not actually supported by any data. What the iCLIP data with the phospho-specific antibodies show is that phosphorylation confers specificity on SF3B1 for the ISL, but this is as likely (or more likely) to be due to a conformational change in the protein upon phosphorylation. For one thing, for this method to work, the antibody needs access to the phosphorylated residues but can’t be disrupting the interaction with RNA. The authors should clarify this in the text and modify Fig. 7b to avoid confusion about this point or inaccuracy in their model.\n11. Line 410-411 (minor): This sentence is ambiguous and potentially confusing; it would be clearer if “in yeast” were moved to immediately precede “SF3B1” (since SF3B1 in humans obviously does contain an IDR).\n12. There is no Supplemental Figure 5 (minor).", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "The manuscript from Gajduskova et. al., describes the role of phosphorylation of SF3B1 (P-SF3B1) in defining a novel intermediate spliceosome B complex (called the B-OTS964), how p-SF3B1 interacts with U6 snRNA, and the role of SNIP1,a component of the RES complex in spliceosome activation. The work follows the groups recent published work that showed CDK11 phosphorylation of SF3B1 using OTS964. The study uses biochemical and genome-wide approaches for these. Use of endogenously dTAG-inserted SNIP1 for acute degradation strengthens many of the studies. A major limitation is that many of the results have already been described in more rigorous structural studies. Overall, it is unclear what is novel, and what the precise message is.\nMajor:\n1. The proposed novel intermediate B-OTS964 is entirely based on biochemical data, which is speculated to have NTR incorporated without NTC. Authors are remiss of intermediate Bact structures reported with cryEM structures have recently been reported with the same configuration of NTR incorporation without NTC (PMID: 39068178). It is unclear why this was not included in references and clarify how the biochemical studies add or refine to this structure.\n2. Similarly, the configuration of SNIP1, a component of RES has been described in other recent solved structures of Bact (PMID: 37165190). This important context is missing and the reference not included – including this help understand what is novel in the findings.\n3. iCLIP showing U6 binding to SF3B1 is presented as novel although the group’s on previous publication (PMID: 36104565) touched upon this. With current known confirmations of Bact, it is not clear why this would be considered novel. Critically, iCLIP for other snRNA is missing ( one would expect strong binding to U2 and minimal interaction with U1).\n4. The claim that CDK11 and SNIP1 act on the same step of spliceosome regulation (line 262) is not supported by the data. In fact analysis of splicing is restricted to global splicing changes at 5’SS and 3’SS. Biochemical analysis of splicing (denaturing gel) and global alternative splicing changes (such as rMATS or SUPPA) are missing. Furthermore, to make a claim that CDK11 and SNIP1 loss have similar effects on splicing, overlap analysis of AS changes in the two conditions should be considered. Otherwise it remains broad failure of splicing.\n5. While the manuscript is heavily reliant on IP and in vitro binding assays, none of these show replicate data or quantification. The critical findings need to be shown as quantified from replicates and statistics.\n6. The assertion that phenotype of E336G homozygosity is due to impaired SNIP1-SF3B1 interaction is not supported by the data presented. Interaction of the mutant with other components of RES or Bact are not shown. Nor is the impact on splicing attempted.\nMinor:\n1. Per structural studies, SF3B1 is not present in B*/C as suggested in the text and Figure 1. This does not affect the conclusions of the study but is important to correct.\n2. iCLIP is semi quantitative given lack of input control. To make quantitiave assumptions is problematic and needs to be discussed/\n3. pSer2 changes in Fig 5C appear minimal and not quantitated. As presented, it does not fully support the claim that CDK9 or 12 does not have effects similar to CDK11 on SF3B1. Longer time points and higher concentrations need to be tested with quantitation to make that claim.\n4. it is unclear why some components of NTC like BCAS2 fall while others such as CDC5L are unchanged with SNIP1 loss. This needs to be discussed in the context of sequential addition of RES, NTR and NTC and available structural data. This is yet another reason to include quantification with replicates for such single blot data.", "reviewer_4": "The manuscript by Gajdušková et al. reveals the role that CDK11-mediated phosphorylation of SF3B1 plays in the specific recruitment of the retention and splicing (RES) complex, which enables proficient splicing. Using the CDK11 inhibitor OTS964, the authors stabilise an intermediate spliceosomal complex, enabling them to investigate the precise function of C-terminal SF3B1 phosphorylation. They elegantly demonstrate that inhibitor treatment causes the loss and gain of critical interactors relevant for the transition from the spliceosomal B complex to the Bact complex. They specifically identify the RES components SNIP1 and BUD13 as direct interactors of the phospho-SF3B1 domain, and demonstrate that loss of SNIP1 (dTAG) or phospho-SF3B1 (OTS964) leads to impaired splicing.\nThrough further mutational investigations, the authors were able to determine how SNIP1 interacts with the phospho-SF3B1 domain, leading them to formulate a well-founded hypothesis regarding the relevance of an E366G mutation in SNIP1 in a neurodevelopmental context.\nThe quality of the data and clarity of the manuscript make this a very interesting study for splicing enthusiasts. A few additional suggestions could be considered without the need for major experiments.\n- Supplementary Table 1: It would be helpful to have a table with all proteins detected in complex with SF3B1 (+/-OTS964) and not just the spliceosome-relevant ones.\n- Figure 1D, Supplementary Fig 1A: How much do these complexes rely on the presence of RNA? Have the authors performed experiments with RNase treatment and thus they assume that all of these interactions are mediated by direct interaction with SF3B1 or are they rather mediated by RNA or a mix of both? The authors should comment on the usage of RNase treatment in the text, as likely many of these interactions are not direct interactors. Could the authors also include RNase treatment-based SF3B1 co-IP experiments to cross-validate their in vitro binding assay experiments?\n- If indeed the recruitment of different subcomplexes is reliant on RNA and/or protein–protein interactions and the IBC complex does not appear to be direct binders of (P-)SF3B1 (Figure 3B), what is the rationale for the lack of this complex from the SF3B1-enriched chromatin fraction after OTS964 treatment? (Steric hindrance with U4 snRNP as well or loss of RES recruitment?)\n- Supplementary Figure 2A: “…resulting in a weaker radioactive signal for RNA-P-SF3B1 complexes…” – Does the autoradiograph really indicate reduced RNA interaction or just reduced IP efficiency because less of the phosphorylated protein is IPed? The authors should provide a WB with their radioactive gels in order to assess IP efficiency versus reduced RNA binding. Otherwise, this statement should be revised. Statements related to understimating the effect of the CDK11 inhibitor on the phospho-SF3B1 iCLIP signal (lines 186–189): The unspecific coverage around the U6 ISL binding site appear to be similar between the libraries. Would this not mean that the general enrichment of U6 sites is similar despite differences in the iCLIP library size? Also, again one needs to ensure that this is a reflection of decreased binding and not of decreased IP efficiency.\n- Figure 2A/B: The transition in the text from the BP CLIP signal to the U6 snRNA focus is not clear. Why are the authors looking at the U6 snRNA? Could the authors include the interaction surface with the U2 snRNA as this should form a complex with the branchpoint. This would be further validation of the U2 snRNP being intake. This paper might be helpful here to infer interaction sites: 10.1261/rna.065664.118\n- Supplementary Figure 2F: Do the authors not find the lack of U6-binding of SFB31 (iCLIP) somewhat concerning? Should the U6 binding not be recovered in SF3B1 data as well, if this was a functionally relevant complex? How sure can the authors be that in the phospho-SF3B1 IP conditions they are only enriching P-SF3B1 protein and nothing else that is part of the complex (have the authors performed proteomics of their IPs?)? Could it be that the U6 binding peak originates from a co-precipitated component? Further experiments should be performed to show interaction of P-SF3B1 to U6. However, seeing as the authors do not go back or use this novel discovery for any further arguments, it might be sufficient to just limit the claim here a little bit.\n- Given that the authors have prepared iCLIP data for P-SF3B1 and RNAseq data for the CDK11 inhibitor, could the authors generate RNA maps to visualize whether the P-SF3B1 binding is preferentially located at CDK11 inhibitor-sensitive sites (similar to 10.1261/rna.069237.118). Could the identity of the targets (GO terms might help here) somehow explain why neurons cannot deal with the SNIP1 mutation?\n- Figure 4D: SNIP1 is known as a transcriptional regulator and acts as an inhibitor of the coactivators CBP and p300 and acts on TGF-beta (J. Biol. Chem. 276:46297-46304 (2001)) and NF-kB signalling. Do the authors see changes in the expression specifically of these pathways as well? Does this play a role here? Previous publications for SNIP1 should be discussed and considered here as well.\n- Figure 4A/D: Given that there are cryoEM structures available, could the authors include a figure to indicate where on the structure SF3B1 and SNIP1 lie and whether they can actually directly interact in the Bact complex (10.1038/cr.2018.14).\n- Figure 5B: Given that BUD13 is also a direct interactor of the phosphorylated-CTD of SF3B1 as per MS and WB, why is this protein not sufficient to assemble the RES to promote productive splicing? Can this be rationalized by looking at the structure of the whole complex or further assembly? Why do all three components need to be there? Would loss of BUD13 result in the same effects?\n- Figure 5A/B: Why does loss of SNIP1 lead to CDK11-dependent hyperphosphorylation? Could the authors include some further discussion of this point?"}, "author_response": "We thank the reviewer for suggestions and positive assessment of the work.\nSpecific concerns:\n1. Line 2: The title is generally a bit acronym-heavy, and in particular I’m afraid readers outside the field will not know what “RES” is. I would recommend giving them at least a clue by saying “RES complex” (minor).\nResponse: We have changed the title, reflecting the suggestion of the reviewer: “SF3B1 phosphorylation by CDK11 orchestrates spliceosome activation via SNIP1-dependent RES complex recruitment”.\n2. In Figure 1c, there are a couple of extreme outliers whose association with SF3B1 drops more drastically than the others in the same complexes (NTC and IBC). What are these (they should probably be labeled)? Are they known to be more loosely associated with their respective complexes, or more labile?\nResponse: The reviewer meant probably Fig. 1d. Outliers correspond to proteins CTNNBL1 (in the NTC complex) and ZNF830 (in the IBC complex). CTNNBL1 is believed not to be a core component of the NTC complex (doi: 10.1093/nar/gkv643) and ZNF830 is a very small protein (29 kDa) and this may result in their loose association with spliceosome intermediate after OTS964 treatment. Order of proteins in each column corresponds to their fold changes after the inhibition-now we have included this information in the figure legend to Fig. 1d.\n3. Lines 172-173: Looking at the correlation analysis in Fig S2b, it is an overstatement to say that the iCLIP replicates for the two anti-P-SF3B1 antibodies “correlated well” (Pearson correlation coefficients for the DMSO treated samples are mostly around 0.5, whereas the OTS964-treated samples show lower correlation, presumably because of lower yield). Compare this to the strong correlations reported in ref. 8 for the total SF3B1 antibody (ED Fig. 9d).\nResponse: We re-ran the correlation analyses for the two phospho-specific antibodies on full gene transcripts using a shorter nucleotide window, which provides higher precision (though at greater computational cost). The updated Pearson correlation coefficients are now included in the manuscript in Supp. Fig. 2a. For the DMSO-treated samples, correlations range from 0.63 to 0.82, while for the OTS964-treated samples they range from 0.31 to 0.52.\nThe somewhat lower correlations observed for P-SF3B1 iCLIP-seq in the DMSO-treated samples, compared to the total SF3B1 iCLIP-seq reported in ref. 8 (doi: 10.1038/s41586-022-05204-z; Extended Data Fig. 9d), are likely due to the use of phospho-specific antibodies and the fact that the phosphorylated form of SF3B1 represents only a minor fraction (10–15%) of total SF3B1 in cells. The reduced correlation in OTS964-treated samples is expected and reflects lower iCLIP yield resulting from SF3B1 dephosphorylation and spliceosome blockage at the B-like intermediate.\n4. Lines 186-189: I am not sure I follow the logic that leads the authors to state that the reduction of P-SF3B1 binding to the U6 internal stem-loop (ISL) “is likely an underestimate.” As I understand the normalization scheme, the OTS964-treated sample was overrepresented (in terms of cell equivalents) to account for decreased protein-RNA binding, but shouldn’t this boost both ISL and non-ISL read counts equally? It is the relative enrichment of ISL in the two conditions (DMSO vs. OTS964) that is being measured here, so I’m not really sure what is being underestimated.\nResponse: We have removed the sentence containing the phrase “is likely an underestimate,” as the reviewer is correct that, given our normalization to an equal number of sequencing reads, and in the absence of input or spike-in controls, the comparison can only be made on a relative basis anyway.\n5. Relatedly, I was unable to deduce from the data shown what the quantitative effect of CDK11 inhibition on U6 binding was (as opposed to the qualitative effect on sequence specificity depicted in Fig. 2). Can this be estimated based on the iCLIP data with the pan-specific anti-SF3B1 antibody?\nResponse: A precise quantitative effect cannot be determined without input material and, ideally, spike-in controls, which are not trivial to implement for iCLIP-seq. However, because the total SF3B1 iCLIP peak over the U6 ISL remains unchanged between DMSO- and OTS964-treated samples after normalization to read counts (Fig. 2d), we can estimate that the reduction in P-SF3B1 binding to the U6 ISL upon OTS964 treatment is approximately 2–3-fold (Fig. 2a, b). This estimate is consistent with our previously published RIP-qPCR data (doi: 10.1038/s41586-022-05204-z, Extended Data Fig. 10), which were normalized to percent of input and decrease of P-SF3B1 binding upon OTS964 is also observed.\nImportantly, OTS964 treatment, as well as the use of two independent phospho-specific SF3B1 antibodies, provides an internal control for the specificity of the P-SF3B1 iCLIP signal. The reproducible P-SF3B1 crosslink patterns on U6, as well as on U1 and U2 snRNAs (now included in the manuscript as Supp. Fig. 2b, c), support the conclusion that phosphorylated SF3B1, that is present only in the activated spliceosome, can indeed contact the U6 ISL.\n6. Lines 213-216: The enrichment of CDK11 and its binding partners (cyclin L, SAP30BP) in the pulldown of P-SF3B1 seems likely to be a trivial consequence of sample preparation differences, i.e., pre-treating only this sample with purified, FLAG-tagged CDK11 complexes. Concluding anything about CDK11’s relative affinities for P-SF3B1 vs. SF3B1 would require a better-controlled experiment, e.g. -/+ ATP, or the use of a catalytically dead mutant CDK11 to generate the non-phosphorylated sample. The authors should probably point this out limitation.\nResponse: In this experiment we did not wish to make any conclusion regarding relative affinities of the CDK11 complex for P-SF3B1 versus SF3B1 and we agree that the presence of the complex in P-SF3B1 sample is most likely a consequence of the sample preparation. We have corrected the relevant section of the text so this is clear.\n7. Lines 256-258: Here is the missed opportunity. Having established the dTAG-SNIP1 system and achieving rescue of splicing defects by ectopic SNIP expression (albeit partial, likely because of low expression), I would like to have seen tests of the various mutant forms of SNIP that abolish or impair binding in vitro to P-SF3B1—K265G, K301G. R365G, E366G—for their ability to rescue.\nResponse: We thank the reviewer for this suggestion. For technical reasons we initially had a problem to prepare flag-tagged SNIP1 (F-SNIP1) proteins (including E366G mutant) that would have similar expression levels with WT F-SNIP1. Comparable amounts of expression are important for the follow up RT(q)-PCR experiments and proper functional comparison between the SNIP1 proteins. In the revised manuscript we were have been able to prepare F-SNIP1 E366G (patient mutation), K265G and double mutant K265G, K301G cell lines, that have similar protein expression levels to the cell line carrying F-SNIP1 WT (Supp. Fig.7 d, e). We performed comparison of splicing by RT-PCR (on agarose gel) and by qPCR on several selected genes and can now observe defect in splicing in all the SNIP1 mutants when compared to F-SNIP1 WT. The data are now presented in Fig. 7e, f and Supp. Fig. 7f.\nAdditionally, we performed proliferation assays with these SNIP1 cell lines after depletion of dTAG-SNIP1 and the mutations affected cellular proliferation correspondingly to their splicing defects. Specifically, patient mutation SNIP1 E366G slowed but did not stop the cellular proliferation, while the other SNIP1 mutations stopped the cellular proliferation more severely. These data are now included in the Fig. 7c, d and support the hypothesis that the mutations affect core splicing and correspondingly also the cellular proliferation.\n8. Relatedly, Gly substitutions are notoriously disruptive of protein conformation; is there a rationale behind changing basic residues to Gly rather than, say, Ala (or acidic residues)?\nResponse: We thank the reviewer for this comment. We originally mutated K301, R365 and K265 to Gly, the same residue present in patients (E366G). As suggested we performed mutation of these residues to Ala and tested incorporation of these SNIP1 proteins to the spliceosome by transient transfection in cells. All tested mutations (i.e. K301A, R365A, K265A and K265A K301A) were less efficiently incorporated to the spliceosome when compared to WT F-SNIP1. This experiment suggests that the observed phenotypes are not just effect of replacement of the tested residues with potentially disruptive Gly mutations on the FHA structure of SNIP1. The results of these experiments are included now in the Supp Fig. 7a.\n9. Lines 291-298 and Fig. 5c: The intent here was to determine the specificity (or exclusivity) of SF3B1 as a CDK11 substrate, given previous reports that other transcriptional CDKs phosphorylate this protein; the conclusion that an OTS964-sensitive kinase is required for the bulk of SF3B1 phosphorylation in cells is probably correct, but the analysis is both incomplete and flawed. Among the flaws or omissions: 1) although CDK7 is named as a candidate SF3B1 kinase (based on ref. 70), no CDK7-selective inhibitor is tested; 2) the CDK9 inhibitor NVP-2 is tested at a very low dose (30 nM) and with a substrate (RNAPII CTD) that is probably not the best (SPT5 is more sensitive and specific: see, for example: doi: 10.1038/s41467-020-18173-6); 3) even so, NVP-2 treatment does decrease the upshifted (i.e., phosphorylated) SF3B1 detected with the pan-specific antibody at the 3-hour time point, but this is not commented upon; 4) the “slightly increased” phospho-Ser2 signal seen after treatment with the CDK12/13 inhibitor THZ531 does not agree with at least one previous study using this compound, which showed a decrease (doi: 10.1038/nchembio.2166); and 5) none of the effects are quantified.\nResponse: We included all these suggestions (and suggestions of the reviewer 3) and in the new version of the Fig. 5c and new Supp. Fig. 5a we show effects of 1) CDK7 inhibition with inhibitor SY-351, 2) use increased concentration of NVP-2 inhibitor (100 nM) and include SPT5 as an additional control substrate and 3) comment on the results in the Result section. Additionally, 4) we cite papers where CDK12 was inhibited (either by selective inhibition of its analog-sensitive version or THZ531) and comment on inhibition by THZ531 in the context of the CDK12/CDK13 literature and 5) given the high number of samples, we chose to present two replicates in the paper and an additional replicate 3 in Raw Data File, instead of quantification.\nOf note, CDK7 or CDK9 inhibition do not result in changes in vitro splicing assays using an RNA reporter and nuclear extract (see doi: 10.1093/nar/gkaf1343; Supp. Fig. 6e and doi: 10.1038/s41586-022-05204-z; Extended Data Fig. 7h). This is in contrast to CDK11 inhibition by OTS964 that affects splicing in vitro (see doi: 10.1038/s41586-022-05204-z; Fig. 3a, b). This provides an additional strong argument that the inhibition of CDK7 and CDK9 do not affect splicing (and SF3B1 phosphorylation) directly, but just indirectly.\n10. Lines 380-382 and Fig. 7b: In both the cartoon model and the indicated text, the authors seem to imply that the phosphorylated residues of the SF3B1 IDR are interacting directly with the ISL of U6 snRNA, but, besides being electrostatically unfavorable (as the authors seem to acknowledge), this is not actually supported by any data. What the iCLIP data with the phospho-specific antibodies show is that phosphorylation confers specificity on SF3B1 for the ISL, but this is as likely (or more likely) to be due to a conformational change in the protein upon phosphorylation. For one thing, for this method to work, the antibody needs access to the phosphorylated residues but can’t be disrupting the interaction with RNA. The authors should clarify this in the text and modify Fig. 7b to avoid confusion about this point or inaccuracy in their model.\nResponse: iCLIP-seq relies on UV crosslinking, which captures only direct RNA–protein contacts before immunoprecipitation under stringent conditions (doi: 10.1016/j.ymeth.2013.10.011). Thus, antibody binding should not interfere with crosslinked RNA-protein interaction. However, we agree that the P-SF3B1 contacts detected on U6 snRNA in the iCLIP data could arise not only from direct interaction of the phosphorylated residues themselves, but also—as the reviewer notes—from phosphorylation-dependent conformational changes within SF3B1 or the spliceosome. Notably, electrostatically unfavorable “interaction” between P-SF3B1 peptides and an RNA was also independently detected in reference 28 (doi: 10.1016/j.molcel.2022.03.024).\nOTS964 treatment affects both, SF3B1 phosphorylation status (thereby reducing recovery with phospho-specific antibodies) and the conformational state of the SF3B1 N-terminus. These two effects cannot be separated in our current dataset. We now acknowledge this explicitly in the Discussion. In the Results section, we have already stated that “OTS964 treatment significantly weakened this interaction, underscoring the importance of both SF3B1 phosphorylation and RNA structure formation in Bact assembly,”. Additionally, we have expanded the Discussion, adjusted the wording throughout the manuscript and text to Fig. 8 legend to clarify that the schematic model and iCLIP results do not explicitly imply that the phosphorylated residues directly bind to the U6 ISL. Instead, the figure legend has been revised to clarify the phosphorylation of SF3B1 as enabling or stabilizing the conformation that allows SF3B1 to engage the U6 ISL.\nThe observed relative decrease in P-SF3B1 binding to the U6 ISL with OTS964 treatment, and the consistent results obtained with two independent phospho-specific antibodies, both serve as controls confirming the specificity of the P-SF3B1 iCLIP signal. We have clarified these points in the revised result text and figure legend.\n11. Line 410-411 (minor): This sentence is ambiguous and potentially confusing; it would be clearer if “in yeast” were moved to immediately precede “SF3B1” (since SF3B1 in humans obviously does contain an IDR).\nResponse: This was corrected\n12. There is no Supplemental Figure 5 (minor).\nResponse: Supplemental Fig. 5 is now included.\nReviewer #2 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.\nResponse: We thank the reviewer for the input and suggestions to improve the manuscript\nReviewer #3 (Remarks to the Author):\nThe manuscript from Gajduskova et. al., describes the role of phosphorylation of SF3B1 (P-SF3B1) in defining a novel intermediate spliceosome B complex (called the B-OTS964), how p-SF3B1 interacts with U6 snRNA, and the role of SNIP1, a component of the RES complex in spliceosome activation. The work follows the groups recent published work that showed CDK11 phosphorylation of SF3B1 using OTS964. The study uses biochemical and genome-wide approaches for these. Use of endogenously dTAG-inserted SNIP1 for acute degradation strengthens many of the studies. A major limitation is that many of the results have already been described in more rigorous structural studies. Overall, it is unclear what is novel, and what the precise message is.\nWe would like to thank the reviewer for suggestions to improve the manuscript and for pointing out the value of the previous structural studies. In short, our work demonstrates the mechanistic role of SF3B1 phosphorylation in spliceosome activation. This is partly due to the phospho-mediated interaction with the FHA domain of SNIP1. Based on our biochemical data and AF3 predictions, we were able to refine the interpretation of the previous cryoEM data (PDB: 8I0R; 10.1038/s41467-024-50785-0 and PDB: 5Z57; 10.1038/cr.2018.14). As now explained in more detail below, the Bact complexes have non-interpreted density interacting with the loops of the SNIP1 FHA domain. Our mutational analysis guided by AF3 prediction provides an interpretation for this additional density. We show that the spatial position of the SF3B1 derived phosphopeptide (repetitive sequence present in SF3B1 between residues 176 and 394) as predicted in complex with the SNIP1 FHA domain matches exactly the region of the uninterpreted cryo-EM density. The predicted SF3B1 phosphopeptide fits very well into the corresponding density (CCC=0.85 at 4 Å map resolution for the AF3 prediction of SNIP1 FHA domain in complex with the SF3B1 phosphopeptide) (Fig. 6d, e). This observation independently validates our in vitro (Supp. Fig. 6) and newly added cellular in vivo data (Fig. 7, Supp. Fig. 7) analysis of SNIP1-loop mutants (K265G, K301G and R365G) and the neurodevelopmental disorder-associated SNIP1 mutation E366G which are all (partially) defective in P-SF3B1 binding, and supports our conclusions that RES assembly on the spliceosome is dependent on CDK11-mediated phosphorylation of SF3B1. In agreement, mutations K265G, K265GK301G and E366G impair pre-mRNA splicing and also cellular proliferation (Fig. 7c, d, e, f and Supp. Fig. 7f).\nMajor:\n1. The proposed novel intermediate B-OTS964 is entirely based on biochemical data, which is speculated to have NTR incorporated without NTC. Authors are remiss of intermediate Bact structures reported with cryEM structures have recently been reported with the same configuration of NTR incorporation without NTC (PMID: 39068178). It is unclear why this was not included in references and clarify how the biochemical studies add or refine to this structure.\nResponse: We thank the reviewer for drawing our attention to this important recent study (PMID: 39068178), which we regret omitting in the initial submission. We have now included and discussed these Bact structures in the context of our biochemical data obtained following CDK11 inhibition with OTS964.\nIn this study (PMID: 39068178), six intermediates between the B and B* complexes were captured using N-palmitoyl-L-leucine (NPL), an inhibitor proposed to interfere with enzymatic activity within the spliceosome and known to affect late-stage spliceosome assembly. A careful examination of the reported structures and Supp. Table 2 shows that three intermediates (pre-Bact, Bact-I, and Bact-II) contain the NTR complex but also include three components of the NTC (CDC5L, CWC15, and PLRG1). The remaining intermediates (Bact-III, Bact-IV, and Post-Bact) contain the full NTR and NTC complexes associated.\nThus, the composition of all reported cryo-EM intermediates differs from our biochemical and quantitative mass spectrometry results (Fig. 1d and Supp. Fig. 1a), in which we observe robust association of all NTR components but a complete absence of NTC components. This distinction suggests that the spliceosome intermediate formed upon selective CDK11 inhibition by OTS964 is compositionally distinct from the intermediates described in the NPL-based structural study.\nWe agree with the reviewer that a cryo-EM structure of the spliceosome following OTS964 treatment would be of great interest; however, such analyses are beyond the scope of the current manuscript. We believe that our quantitative mass spectrometry data (Fig. 1d), independently corroborated by immunoprecipitation-free isolation of chromatin-associated spliceosomes from cells (Supp. Fig. 1a), provide new biochemical insights into a spliceosome intermediate formed upon CDK11 inhibition. This intermediate is characterized by differential recruitment of the complete NTR complex in the absence of the NTC, refining current models of spliceosome assembly and regulation.\n2. Similarly, the configuration of SNIP1, a component of RES has been described in other recent solved structures of Bact (PMID: 37165190). This important context is missing and the reference not included – including this help understand what is novel in the findings.\nResponse: We have now included the discussion of our results in the context of existing cryo-EM models and also cite the reference PMID: 39068178. Notably, our data help to interpret a previously unassigned density in the cryo-EM maps of Bact structures as suggested by the reviewer (PDB: 8I0R; 10.1038/s41467-024-50785-0). Combining our biochemical data with AF3 prediction, we now show in Fig. 6e that the additional density very likely corresponds to the P-ThrPro[X] P-ThrPro motif of SF3B1 (28 such motifs are present between SF3B1 residues 176 and 394) which binds firmly to SNIP1 FHA domain through two phosphorylated Thr residues and anchors (or recruits) SNIP1 to Bact complex. Our biochemical data further show that this interaction is not unique to particular P-ThrPro[X] P-ThrPro motif, as mutations in Thr357Ala and Thr369Ala in SF3B1 did not change the SNIP1 binding affinity (Supp. Fig. 6d).\n3. iCLIP showing U6 binding to SF3B1 is presented as novel although the group’s on previous publication (PMID: 36104565) touched upon this. With current known confirmations of Bact, it is not clear why this would be considered novel. Critically, iCLIP for other snRNA is missing (one would expect strong binding to U2 and minimal interaction with U1).\nResponse: We mention in the introduction and result sections that P-SF3B1 was found by RIP-qPCR to associate with U6 snRNA (published in PMID: 36104565) and limitations of the RIP-qPCR experiment (which did not use UV crosslinking). Importantly, our previous study did not provide any clues whether these associations are direct or indirect and single nucleotide-resolution association of (P)-SF3B1 to U6 snRNA (and other snRNAs) are unknown. We now also include single-nucleotide resolution binding profiles of (P)-SF3B1 to U1 snRNA (no binding as expected) and U2 snRNAs (predominantly binding of SF3B1 by iCLIP is observed) (Supp. Fig. 2b, c). We believe that the presented iCLIP data will be of interest for future studies of roles of (P)-SF3B1-RNA interactions in the U2/U6 snRNA catalytic core structure of the activated spliceosome.\n4. The claim that CDK11 and SNIP1 act on the same step of spliceosome regulation (line 262) is not supported by the data. In fact analysis of splicing is restricted to global splicing changes at 5’SS and 3’SS. Biochemical analysis of splicing (denaturing gel) and global alternative splicing changes (such as rMATS or SUPPA) are missing. Furthermore, to make a claim that CDK11 and SNIP1 loss have similar effects on splicing, overlap analysis of AS changes in the two conditions should be considered. Otherwise it remains broad failure of splicing.\nResponse: To demonstrate that loss of CDK11 kinase activity and SNIP1 depletion have similar effects on splicing, we analyzed fold changes in intron ratios (i.e. ratios of intronic to exonic reads for each gene) across 12,727 genes (Fig. 4c) using the RNA-seq datasets after OTS964 inhibition (doi: 10.1038/s41586-022-05204-z, Fig. 2a) and degradation of SNIP1 (this manuscript, Fig. 4b). This analysis (presented in Fig. 4c now) revealed a strong correlation between fold changes in intron ratios in the dTAG-SNIP1/DMSO and OTS964/DMSO conditions (Spearman correlation coefficient = 0.79), indicating highly similar global splicing defects. These results more rigorously support a shared role of SNIP1 and CDK11 kinase activity in regulating constitutive splicing, particularly global intron retention.\nWe did not perform in vitro splicing assays using denaturing gels because these assays require HeLa nuclear extracts (Prof. Chris Smith, University of Cambridge, personal communication), whereas our dTAG-SNIP1 system was engineered in HCT116 cells (to be able to compare to OTS964 treatment performed also in HCT116 cells). Generating a degron-tagged SNIP1 HeLa cell line for such assays would require substantial time and resources, with uncertain feasibility. To further support our statement that “Highly similar splicing defects strongly suggest that SNIP1 and CDK11 act on the same step of spliceosome regulation” we have also added new data characterizing the functional consequences of SNIP1 mutations in cells (Fig. 7; Supp. Fig. 7), that complement biochemical characterization of SNIP1 binding to P-SF3B1 (Supp. Fig. 6). Together with comparative analysis of intron retention fold changes (Fig. 4c) and CDK11-mediated hyperphosphorylation of SF3B1 after SNIP1 depletion (Fig. 5 b, c), these complementary approaches collectively provide strong support for the conclusion that SNIP1 and CDK11 act at the same step (IDR of SF3B1) of spliceosome regulation and that SNIP1 is required for global constitutive splicing in cells.\nFinally, tools such as rMATS or SUPPA were developed to detect and quantify alternative splicing events (e.g., exon skipping or alternative splice site usage; doi:10.1093/nar/gkr1291) and are not designed to analyze defects in constitutive splicing. Consistent with this limitation, our analyses of the dTAG SNIP1 and OTS964 RNA-seq datasets using rMATS did not yield interpretable or meaningful results. Because the primary splicing defect observed in our study is global intron retention (Fig. 4a, b, c; Supp. Fig. 4d, e), we therefore did not include analyses based on these tools in the manuscript.\n5. While the manuscript is heavily reliant on IP and in vitro binding assays, none of these show replicate data or quantification. The critical findings need to be shown as quantified from replicates and statistics.\nResponse: All experiments (including confirmatory ones) are now presented with at least two independent replicates, and in many cases three (or more). Replicate data are shown in the manuscript, with additional replicates provided in the Resource Data File. Quantification and/or statistics were performed for characterization of SNIP1 mutations by immunoprecipitation (Fig. 7a, b) and for RT-(q)PCR splicing assays (Fig. 7e, f, Supp. Fig. 7f) and for all mass spectrometry data (Fig. 1c, d, Fig. 3b)\n6. The assertion that phenotype of E336G homozygosity is due to impaired SNIP1-SF3B1 interaction is not supported by the data presented. Interaction of the mutant with other components of RES or Bact are not shown. Nor is the impact on splicing attempted.\nResponse: We thank the reviewer for this question and helpful suggestions. In the revised manuscript, we performed additional immunoprecipitation experiments and show that the E366G mutant (as well as other SNIP1 mutants) interact less efficiently with components of the RES and Bact complexes when compared with wild-type (WT) F-SNIP1. These results are now presented in Fig. 7a, b and Supp. Fig. 7a, b, c.\nWith respect to testing the SNIP1 E366G mutant in splicing assays, we initially encountered technical difficulties in generating flag-tagged SNIP1 (F-SNIP1) cell lines, including the E366G mutant, that exhibited expression levels lower than WT F-SNIP1. Comparable and sufficiently high expression levels of F-SNIP1 proteins are essential for subsequent RT–(q)PCR experiments and for accurate functional comparison among SNIP1 variants. In the revised manuscript, we were able to generate flag-tagged F-SNIP1 WT, SNIP1 E366G (patient mutation), SNIP1 K265G, and SNIP1 K265G/K301G cell lines expressing comparable levels of F-SNIP1 proteins (Supp. Fig. 7d, e). We then assessed splicing by RT–PCR (agarose gel analysis) and by qPCR for several selected genes. These experiments revealed splicing defects in all SNIP1 mutants compared with WT SNIP1. These data are now presented in Fig. 7e, f and Supp. Fig. 7f.\nAdditionally, we performed proliferation assays following depletion of endogenous dTAG-SNIP1 and parallel expression of the SNIP1 variants. The mutations affected cellular proliferation in a manner corresponding to their splicing defects. Specifically, the patient mutation SNIP1 E366G slowed, but did not completely block, cellular proliferation, whereas the other SNIP1 mutations inhibited cellular proliferation more severely. These results are now presented in Fig. 7c, d and support the conclusion that these mutations impair core splicing and consequently affect cellular proliferation.\nMinor:\n1. Per structural studies, SF3B1 is not present in B*/C as suggested in the text and Figure 1. This does not affect the conclusions of the study but is important to correct.\nResponse: We have corrected Fig. 1 and the corresponding figure legend to clarify when SF3B1 is phosphorylated (from the Bact to C intermediate) and when, based on structural studies, SF3B1 is present in the spliceosome (from the A to B* intermediate).\n2. iCLIP is semi quantitative given lack of input control. To make quantitiave assumptions is problematic and needs to be discussed/\nResponse: In line with this suggestion (and that of other reviewers) we discuss now this issue in the Discussion.\n3. pSer2 changes in Fig 5C appear minimal and not quantitated. As presented, it does not fully support the claim that CDK9 or 12 does not have effects similar to CDK11 on SF3B1. Longer time points and higher concentrations need to be tested with quantitation to make that claim.\nResponse: In line with this suggestion (and that of reviewer 1), we have expanded the analysis and now present revised Fig. 5c and Supp. Fig. 5a. Specifically, we include longer inhibitor treatment time points (30 min to 4 h), the selective CDK7 inhibitor SY-351 (doi: 10.1101/gad.341545.120), and an increased concentration of the CDK9 inhibitor NVP-2 (from 30 to 100 nM; IC₅₀ ~1 nM; doi: 10.1038/nchembio.2538).\nWe did not increase the concentration of the covalent CDK12/CDK13 inhibitor THZ531, as concentrations above 100 nM are known to inhibit additional kinases, including CDK7, which is required for CDK11 activation (doi: 10.1038/nsmb1028; doi: 10.1101/gad.341545.120; doi: 10.1093/nar/gkaf1343). Thus, higher concentrations would be expected to produce non-specific effects. The activity of THZ531 at 100 nM is supported by previously reported subtle changes in RNAPII CTD phosphorylation and SPT5 Thr806 phosphorylation at longer time points, as well as reduced expression of CDK12-dependent DNA damage response genes (doi: 10.1038/nchembio.2166; see Fig. 6b). Consistent with this, similarly subtle and heterogeneous CTD phosphorylation changes accompanied by decreased expression of long DNA damage response genes have been documented using selective CDK12 inhibition in cells carrying an analog-sensitive CDK12 allele (doi: 10.1093/nar/gkaa514; doi: 10.15252/embr.201847592; doi: 10.1126/sciadv.aaz5041; doi: 10.1016/j.bbagrm.2015.07.010; doi: 10.1093/narcan/zcaa003). We also add a Fig. R1 to document comparable effects of treatment of cells on expression of CDK12-dependent DNA damage response genes when treated with 100 nM THZ531 and when CDK12 is selectively inhibited in HCT116 cells carrying engineered endogenous analog-sensitive (AS) variants of either CDK12 or CDK13 (control).\nFig. R1: 100 nM THZ531 reduces expression of CDK12-dependent DNA damage response genes. Comparison of expression of CDK12-dependent DNA damage response genes after 6 h treatment of maternal HCT116 cells with 100 nM THZ531 and 6 h inhibition of either CDK12 or CDK13 by 10 µM ATP analog 3MB-PP1 in HCT116 cells carrying engineered endogenous analog-sensitive (AS) variants of either CDK12 or CDK13 (control).\nRather than quantifying effects of all inhibitors and time points, which can be technically challenging and potentially misleading given the subtle and heterogeneous nature of some phosphorylation changes and high number of samples and conditions, we now present two independent biological replicates in the manuscript (Fig. 5c, Supp. Fig. 5a), and the third replicate is provided in the Resource Data File. We believe that the distinct dynamics and magnitude of the effect of 50 nM OTS964 on SF3B1 phosphorylation, compared with the other inhibitors tested, are clearly evident.\nFinally, treatments with the inhibitors were performed following complete degradation of SNIP1 protein. Under these conditions, differential effects of the inhibitors on their canonical substrates (such as the RNAPII CTD or SPT5), which are typically characterized in SNIP1-proficient cells, cannot be fully excluded. Consistent with this, phosphorylation of SPT5 Thr806 upon SNIP1 depletion is visibly increased compared with control cells. Additionally, we have expanded the discussion of these results and their interpretation in the Results section of the revised manuscript.\n4. it is unclear why some components of NTC like BCAS2 fall while others such as CDC5L are unchanged with SNIP1 loss. This needs to be discussed in the context of sequential addition of RES, NTR and NTC and available structural data. This is yet another reason to include quantification with replicates for such single blot data.\nResponse: We thank the reviewer for raising this point. We repeated the experiment with two additional independent replicates, which are now included in the Resource Data File. These experiments faithfully reproduced the levels of all proteins and complexes shown in Fig. 4d.\nDuring this process, we identified a technical error in the original CDC5L blot: the membrane corresponding to CDC5L in the initial replicate was inadvertently swapped. This was confirmed by rerunning the CDC5L western blot from the original replicate, which yielded results consistent with the additional replicates. The corrected data are now shown in the revised Fig. 4d and supported by the additional replicates provided in the Resource Data File.\nReviewer #4 (Remarks to the Author):\nThe manuscript by Gajdušková et al. reveals the role that CDK11-mediated phosphorylation of SF3B1 plays in the specific recruitment of the retention and splicing (RES) complex, which enables proficient splicing. Using the CDK11 inhibitor OTS964, the authors stabilise an intermediate spliceosomal complex, enabling them to investigate the precise function of C-terminal SF3B1 phosphorylation. They elegantly demonstrate that inhibitor treatment causes the loss and gain of critical interactors relevant for the transition from the spliceosomal B complex to the Bact complex. They specifically identify the RES components SNIP1 and BUD13 as direct interactors of the phospho-SF3B1 domain, and demonstrate that loss of SNIP1 (dTAG) or phospho-SF3B1 (OTS964) leads to impaired splicing.\nThrough further mutational investigations, the authors were able to determine how SNIP1 interacts with the phospho-SF3B1 domain, leading them to formulate a well-founded hypothesis regarding the relevance of an E366G mutation in SNIP1 in a neurodevelopmental context.\nThe quality of the data and clarity of the manuscript make this a very interesting study for splicing enthusiasts. A few additional suggestions could be considered without the need for major experiments.\nResponse: We thank the reviewer for suggestions and positive assessment of the work.\n-Supplementary Table 1: It would be helpful to have a table with all proteins detected in complex with SF3B1 (+/-OTS964) and not just the spliceosome-relevant ones.\nResponse: All detected proteins are part of the Supp. Data 1, sheet “IP SF3B1 (Chromatin) all data”.\n-Figure 1D, Supplementary Fig 1A: How much do these complexes rely on the presence of RNA? Have the authors performed experiments with RNase treatment and thus they assume that all of these interactions are mediated by direct interaction with SF3B1 or are they rather mediated by RNA or a mix of both? The authors should comment on the usage of RNase treatment in the text, as likely many of these interactions are not direct interactors. Could the authors also include RNase treatment-based SF3B1 co-IP experiments to cross-validate their in vitro binding assay experiments?\nResponse: We thank the reviewer for raising this point. The purpose of the experiments shown in Fig. 1d and Supp. Fig. 1a was to isolate the chromatin-associated spliceosome, as previously described (doi: 10.1038/s41586-022-05204-z; doi: 10.1038/nature14512; doi: 10.1038/ncomms1998) and characterize its composition (with or without the OTS964 treatment).\n[REDACTED]\n11\n[REDACTED]\n-If indeed the recruitment of different subcomplexes is reliant on RNA and/or protein–protein interactions and the IBC complex does not appear to be direct binders of (P-)SF3B1 (Figure 3B), what is the rationale for the lack of this complex from the SF3B1-enriched chromatin fraction after OTS964 treatment? (Steric hindrance with U4 snRNP as well or loss of RES recruitment?)\nResponse: The experiment shown in Fig. 3b was designed to identify potentially direct interactors of SF3B1 in both its phosphorylated and unphosphorylated states. Cellular lysates were fully solubilized by treatment with RNase A to maximally disrupt the spliceosome and its subcomplexes from various RNA-dependent subcomplexes, thereby facilitating the release of individual splicing factors from their pre-assembled complexes. Candidate direct interactions were subsequently validated using in vitro binding assays with purified proteins, as demonstrated for SNIP1 and GST–SF3B1 (Supp. Fig. 6.)\nThe absence of the IBC complex in OTS964-treated samples (Fig. 1d) reflects arrest of the spliceosome at an intermediate stage prior to IBC recruitment and does not imply any specific mechanism of direct or indirect interaction. The order of assembly events can be inferred from our mass spectrometry data (Fig. 1d) in conjunction with published structural studies. For example, structural work (doi: 10.1126/science.abc3753, Supp. Fig. 6a, b) indicates that the IBC complex is recruited only after the NTR, NTC, and RES complexes have been assembled into the spliceosome (pre-Bact-1 intermediate). Consistent with this model, the absence of RES and NTC in the OTS964-arrested spliceosome prevents IBC recruitment, thereby blocking formation of the subsequent pre-Bact-2 intermediate.\n-Supplementary Figure 2A: “…resulting in a weaker radioactive signal for RNA-P-SF3B1 complexes…” – Does the autoradiograph really indicate reduced RNA interaction or just reduced IP efficiency because less of the phosphorylated protein is IPed? The authors should provide a WB with their radioactive gels in order to assess IP efficiency versus reduced RNA binding. Otherwise, this statement should be revised. Statements related to understimating the effect of the CDK11 inhibitor on the phospho-SF3B1 iCLIP signal (lines 186–189): The unspecific coverage around the U6 ISL binding site appear to be similar between the libraries. Would this not mean that the general enrichment of U6 sites is similar despite differences in the iCLIP library size? Also, again one needs to ensure that this is a reflection of decreased binding and not of decreased IP efficiency.\nResponse: We thank the reviewer for this important question. We agree that, following OTS964 treatment, it is not possible to unambiguously distinguish between (i) reduced immunoprecipitation efficiency of phosphorylated SF3B1 due to its dephosphorylation and (ii) a genuine reduction in RNA binding caused by OTS964-induced stalling of the spliceosome at the BOTS964 intermediate. This is now discussed in the Discussion.\nUnfortunately, we do not have a western blot corresponding directly to the radioactive gel shown in former Supp. Fig. 2a to assess IP efficiency in this specific experiment. However, we have previously published western blot analyses from RNA immunoprecipitation experiments using both total SF3B1 and phospho-Thr235-SF3B1 antibodies (doi: 10.1038/s41586-022-05204-z, Review File, Fig. RR4), which demonstrate reduced immunoprecipitation efficiency following OTS964 treatment. These data support the notion that decreased recovery of P-SF3B1 is, at least in part, due to loss of the phosphorylated epitope also in the iCLIP experiment presented in this manuscript.\nIn response to this and related comments from other reviewers, we have substantially revised the Results section entitled “P-SF3B1 interacts with the U6 snRNA ISL in the activated spliceosome” and expanded the Discussion. In particular, we removed above mentioned statement and we now emphasize that OTS964 treatment, together with the use of two independent phospho-specific SF3B1 antibodies, serve as internal controls for the specificity of the P-SF3B1 iCLIP signal at the U6 snRNA ISL. Moreover, the reproducible P-SF3B1 crosslinking patterns observed not only on U6, but also on U1 and U2 snRNAs (now included as Supp. Fig. 2b, c), support the conclusion that phosphorylated SF3B1, present specifically in the activated spliceosome, can contact the U6 ISL or help elicitate contact with U6 ISL.\nWe further clarify in the Discussion that iCLIP-seq is inherently semi-quantitative in the absence of input material or spike-in controls. Importantly, the total SF3B1 iCLIP signal over the U6 ISL remains comparable between DMSO- and OTS964-treated samples after normalization to total read counts (Fig. 2d). Based on this normalization, we estimate that P-SF3B1 associating with the U6 ISL is reduced approximately 2–3-fold upon OTS964 treatment. This estimate is consistent with our previously published RIP-qPCR data (doi: 10.1038/s41586-022-05204-z, Extended Data Fig. 10), which were normalized to percent of input and likewise show a reduction in P-SF3B1 binding following OTS964 treatment\n- Figure 2A/B: The transition in the text from the BP CLIP signal to the U6 snRNA focus is not clear. Why are the authors looking at the U6 snRNA? Could the authors include the interaction surface with the U2 snRNA as this should form a complex with the branchpoint. This would be further validation of the U2 snRNP being intake. This paper might be helpful here to infer interaction sites: 10.1261/rna.065664.118\nResponse: We thank the reviewer for this comment and suggestions. As noted in our response above, we have substantially rewritten the corresponding Results section and expanded the Discussion to clarify the main focus of the manuscript, which is the characterization of the contact between phosphorylated SF3B1 (P-SF3B1) and U6 snRNA.\nTo avoid confusion and to improve narrative clarity, we have removed the section describing branchpoint (BP) and pre-mRNA interactions with (P-)SF3B1. While these interactions are important, a detailed analysis of BP- or pre-mRNA-associated interactions is beyond the scope of the present study and would detract from the main message focused on U6 snRNA binding.\nWe focus on U6 snRNA because it is a core component of the spliceosome RNA catalytic center that is present in the activated spliceosome (when SF3B1 is in its phosphorylated form). In our previous work, RNA immunoprecipitation (RIP) followed by qPCR showed that U6 snRNA is preferentially associated with phosphorylated SF3B1 (doi: 10.1038/s41586-022-05204-z, Ext. Data Fig. 10). Given that iCLIP provides single-nucleotide resolution, our goal here was to precisely map where P-SF3B1 contacts U6 snRNA within the activated spliceosome, defined by SF3B1 phosphorylation. The mapping suggests how the P-SF3B1 engagement may contribute to the formation and/or stabilization of the U2/U6 snRNA catalytic core in activated and catalytically competent spliceosomes.\nWe have now also included mapped iCLIP interactions of both SF3B1 and P-SF3B1 on U2 snRNA. These data show that U2 snRNA is bound predominantly by total SF3B1, with substantially weaker binding by P-SF3B1. In addition, binding to U1 snRNA, included as a negative control, has now been incorporated into the manuscript (Supp. Fig. 2b, c).\n- Supplementary Figure 2F: Do the authors not find the lack of U6-binding of SFB31 (iCLIP) somewhat concerning? Should the U6 binding not be recovered in SF3B1 data as well, if this was a functionally relevant complex? How sure can the authors be that in the phospho-SF3B1 IP conditions they are only enriching P-SF3B1 protein and nothing else that is part of the complex (have the authors performed proteomics of their IPs?)? Could it be that the U6 binding peak originates from a co-precipitated component? Further experiments should be performed to show interaction of P-SF3B1 to U6. However, seeing as the authors do not go back or use this novel discovery for any further arguments, it might be sufficient to just limit the claim here a little bit.\nResponse: We thank the reviewer for raising this concern. This comment prompted us to re-examine the SF3B1 and P-SF3B1 iCLIP binding profiles on U6 snRNA in detail. Upon closer inspection, we noted that total SF3B1 does in fact display a binding peak at the U6 ISL that precisely overlaps with the P-SF3B1 binding site identified using both phospho-Thr235– and phospho-Thr211–specific antibodies. To make this clearer to a reader, we now present the binding profiles aligned directly beneath each other (Fig. 2a, b, d), and we have revised the corresponding Results text accordingly.\nThe use of two independent phospho-specific SF3B1 antibodies, together with an antibody recognizing total SF3B1, yields highly similar binding profiles on U6 snRNA, as well as on U1 and U2 snRNAs (Supp. Fig. 2b, c). This reproducibility strongly supports the conclusion that the observed associations are specific. Nevertheless, we agree that we cannot formally exclude the possibility that additional spliceosomal component(s) are co-enriched in the immunoprecipitations and could contribute to the observed signal. We now explicitly acknowledge this limitation in the Discussion.\nWe have not performed proteomic analyses of the immunoprecipitated material, and we agree that additional biochemical or in vitro approaches would be required to definitively demonstrate a direct interaction between P-SF3B1 and U6 snRNA (and RNA catalytic core). Such experiments, however, are beyond the scope of the present study.\nIn line with the suggestion of this and other reviewers, we have also moderated and better explained our claims in the Results and Discussion to reflect limitations of the iCLIP.\n- Given that the authors have prepared iCLIP data for P-SF3B1 and RNAseq data for the CDK11 inhibitor, could the authors generate RNA maps to visualize whether the P-SF3B1 binding is preferentially located at CDK11 inhibitor-sensitive sites (similar to 10.1261/rna.069237.118). Could the identity of the targets (GO terms might help here) somehow explain why neurons cannot deal with the SNIP1 mutation?\nResponse: We thank the reviewer for this interesting suggestion. Although we generated iCLIP data for (P)-SF3B1, we chose to remove the data depicting (P)-SF3B1 binding to pre-mRNA from the manuscript (former Supp. Fig. 2a, c, d, e), as these data detracted from the main focus of the study, namely the mechanistic function of (P)-SF3B1 within the spliceosome.\nIn addition, CDK11 inhibition affects global (constitutive) splicing (doi: 10.1038/s41586-022-05204-z; Fig. 2), impacting the splicing of the majority of introns. iCLIP binding profiles of (P)-SF3B1, and, in our experience, of other constitutive splicing factors, do not provide sufficient coverage across most introns. As a result, a direct comparison between (P)-SF3B1 iCLIP binding and CDK11 inhibitor–sensitive splicing events could be misleading. For this reason, we did not attempt to generate RNA maps or perform a comprehensive overlap or gene ontology (GO) analysis of (P)-SF3B1–bound targets versus CDK11 inhibitor–sensitive splicing events, either in this study or in our previous work (doi: 10.1038/s41586-022-05204-z). We agree that such analyses would be of interest; however, they would require substantial additional optimization of the iCLIP experimental design to improve coverage of pre-mRNA binding. This is beyond the scope of the present study.\nRegarding the neuronal sensitivity to SNIP1 mutations (and to mutations in many other splicing factors), we believe that this vulnerability is best explained by well-established features of neuronal gene architecture. Many genes that are specifically expressed in neurons and neuronal tissues are exceptionally long and intron-rich, rendering them particularly dependent on efficient and tightly regulated spliceosome function. Consequently, perturbations in spliceosome components or their regulatory pathways, including SNIP1, disproportionately affect neuronal cells. This concept is supported by numerous previous studies (e.g., doi:10.1186/s12864-020-6760-4; doi:10.1002/wrna.1532).\n- Figure 4D: SNIP1 is known as a transcriptional regulator and acts as an inhibitor of the coactivators CBP and p300 and acts on TGF-beta (J. Biol. Chem. 276:46297-46304 (2001)) and NF-kB signalling. Do the authors see changes in the expression specifically of these pathways as well? Does this play a role here? Previous publications for SNIP1 should be discussed and considered here as well.\nResponse: Our new analyses of RNA-seq data following dTAC-mediated degradation of SNIP1, and their comparison with CDK11 inhibition, reveal a constitutive (broad) splicing defect (Fig. 4c), without obvious enrichment for effects on specific gene groups or signaling pathways. In addition, we observe that single-point mutations in the SNIP1 FHA domain lead to defects in splicing and cellular proliferation (Fig. 7c–f; Supplementary Fig. 7f), supporting a direct role for SNIP1 in spliceosome function.\nWhether TGF-β or NF-κB signaling pathways contribute to the observed phenotypes remains unclear. Most previous studies linking SNIP1 to transcriptional regulation of these pathways relied on long-term siRNA-mediated knockdown, protein overexpression, and reporter assays, which were the standard technologies available at the time but have clear limitations in distinguishing primary from secondary effects. Importantly, such approaches cannot readily discriminate between direct transcriptional effects of SNIP1 and secondary transcriptional changes resulting from SNIP1-mediated perturbations of splicing. It is now well-established that disruption of spliceosome function can rapidly feedback on transcriptional regulation (doi: 10.1016/j.molcel.2021.02.016; doi: 10.1038/s41586-022-05204-z).\nWe now cite these earlier publications describing transcriptional roles of SNIP1 and links to TGF-β or NF-κB signaling pathways and discuss some of these limitations in the Discussion. Taken together, our data support a primary role for SNIP1 in constitutive splicing, while not excluding the possibility that additional transcription-related effects may arise indirectly as a consequence of splicing perturbation or that SNIP1 also affects transcription directly.\n- Figure 4A/D: Given that there are cryoEM structures available, could the authors include a figure to indicate where on the structure SF3B1 and SNIP1 lie and whether they can actually directly interact in the Bact complex (10.1038/cr.2018.14).\nResponse: We thank the reviewer for this suggestion. There are two cryo-EM structures that contain information on SF3B1 and SNIP1 proteins and we discuss both in the manuscript now. The first structure, denoted as late Bact, was reported in 2018 at 6.5 Å resolution (PDB: 5Z57; 10.1038/cr.2018.14) and the second one, denoted as Bact-I, in 2024 at 3 Å resolution (PDB: 8I0R; 10.1038/s41467-024-50785-0) by the same group. Both structures were analyzed in light of the interaction between SF3B1 and SNIP1 proteins. In common density was assigned to the FHA domain of SNIP1 (aa 221-374) and the structured C-terminal part of SF3B1 that includes the HEAT repeats (aa 395-2304). Overall the spatial arrangement of SF3B1 and SNIP1 does not differ in the two structures. As shown in Fig. 6d, a small portion of SF3B1 contacts one side of the twisted β sandwich of the FHA domain. This is not a typical interaction for an FHA domain and is very likely the result of SF3B1 and SNIP1 positioning in the Bact complex as a result of contacts with the rest of the spliceosome components (not shown). The uninterpreted density next to SNIP1 is surface exposed, supporting further that the phosphorylated motifs of SF3B1 could control recruitment of SNIP1 to spliceosome. This information is now included and analysed in new Fig. 6d, e and discussed in the Result and Discussion sections.\n- Figure 5B: Given that BUD13 is also a direct interactor of the phosphorylated-CTD of SF3B1 as per MS and WB, why is this protein not sufficient to assemble the RES to promote productive splicing? Can this be rationalized by looking at the structure of the whole complex or further assembly? Why do all three components need to be there? Would loss of BUD13 result in the same effects?\nResponse: We thank the reviewer for this insightful question, which we believe refers to the data presented in Fig. 3b, c. BUD13 (and RBMX2), the remaining components of the RES complex, are not associated with either SF3B1 or phosphorylated SF3B1 (P-SF3B1) in our mass spectrometry analysis (Fig. 3b). To clarify this point, we have now highlighted both proteins in the volcano plot in Fig. 3b.\nWe also revisited our western blot analyses (Fig. 3c) and re-tested binding of BUD13 and RBMX2 to SF3B1 and P-SF3B1. These experiments did not detect a specific interaction. The previously shown BUD13 signal reflected a background interaction with GST–SF3B1 (1–461) that was independent of the phosphorylation status of SF3B1. Accordingly, we have removed the BUD13 western blot from the revised manuscript and updated the text to reflect this correction.\n- Figure 5A/B: Why does loss of SNIP1 lead to CDK11-dependent hyperphosphorylation? Could the authors include some further discussion of this point?\nResponse: We propose that the CDK11-dependent hyperphosphorylation observed upon acute depletion of SNIP1 reflects a rapid compensatory cellular response to the loss of a spliceosome component (SNIP1). This increase in phosphorylation likely indicates, that phosphorylation of SF3B1 by CDK11 represents a rate-limiting step for SNIP1 (RES complex) binding or stable anchoring to the spliceosome. We have now included this explanation in the relevant part of the Results section."}, {"round": 2, "reviewer_comments": {"reviewer_1": "This is a revised version of a manuscript I reviewed previously, The authors have addressed all my concerns and greatly strengthened the study. I believe they have also adequately addressed the other reviewers’ critiques. I have only a few, small comments and suggestions for further revision (which would not necessitate another round of review) listed below:\n1. Line 108: I was tripped up by the odd syntax, “…with the integrated NTR complex but NTC complex not associated.” Rephrasing it as, “…with the NTR complex integrated but the NTC complex not associated” would be much clearer.\n2. Line 112: There should be a comma between “activation” and “and”.\n3. Lines 284-286: An alternative, perhaps more likely explanation for the SF3B1 hyperphosphorylation seen after SNIP1 degradation is a “spliceosome cycle arrest,” with SF3B1 dephosphorylation (presumably by PP1) being blocked while CDK11 activity remains high.\n4. Lines 414-416, Fig. 7e: There is a labeling mistake in the figure, “unspl./spl.” Instead of “spl./unspl.” (the figure legend is correct).", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "The authors have responded to most of my comments satisfactorily and I have no further concerns.", "reviewer_4": "In their revised manuscript, Gajdušková et al. present a rigorous and well-executed study that defines how CDK11-mediated phosphorylation of SF3B1 controls spliceosome activation by recruiting the RES complex via SNIP1. Through the integration of quantitative proteomics, iCLIP, biochemical assays, acute protein degradation, mutational analysis and structural interpretation, the authors present a coherent and mechanistically compelling model.\nThe authors have thoroughly addressed the reviewers’ concerns. The revised manuscript now clearly positions the B(OTS964) intermediate within the context of recent cryo-EM studies, explaining how it is compositionally distinct from previously described activation states. The integration of AlphaFold-based modelling with functional mutational analyses is particularly notable, as it allows the interpretation of previously unassigned density in BAC structures.\nNotably, the newly incorporated cellular experiments that test SNIP1 FHA-domain mutants (including the disease-associated E366G variant) substantially reinforce the study by establishing a direct link between impaired phospho-SF3B1 recognition and defects in spliceosome assembly, global splicing, and cell proliferation. The authors have also appropriately moderated claims related to iCLIP quantification and clarified limitations where necessary.\nOverall, this is a high-quality, well-revised manuscript that provides important mechanistic insights into spliceosome activation relevant to both fundamental biology and disease. I have no further concerns and support publication in Nature Communications."}, "author_response": "We thank the reviewer for suggestions that we have included in the revised manuscript."}]

Decision letter