The G-patch protein Cwf28 interacts with the RNA helicase Cdc28 in catalytically active spliceosomes in Schizosaccharomyces pombe

Laura Olivia Karika; Ingrid Cipakova; Miroslava Kretova; Tomas Selicky; Natalia Cmikova; Daniela Nemcekova; Lenka Kohutova; Peter Barath; Lubos Cipak

doi:10.1371/journal.pone.0357276

Abstract

G-patch proteins are emerging as key regulatory cofactors of DEAH-box RNA helicases involved in pre-mRNA splicing, yet the functions of many family members remain poorly understood. Here, we characterize the conserved G-patch domain–containing protein Cwf28 from Schizosaccharomyces pombe, an essential factor with a previously unclear molecular function, and define its role within the spliceosome. Tandem affinity purification coupled with mass spectrometry revealed that Cwf28 predominantly associates with components of the Prp19 complex (NTC) and factors involved in the catalytic activation and progression of the spliceosome, placing it within catalytically active spliceosomal assemblies. Gene ontology analysis showed enrichment of the Cwf28 interactome in factors involved in spliceosome assembly, activation, and catalytic remodeling. Notably, the DEAH-box RNA helicase Cdc28, the ortholog of human DHX16 and Saccharomyces cerevisiae Prp2, was identified as the most abundant interactor. Further analysis demonstrated that Cwf28 interacts with Cdc28 via its conserved G-patch domain, and domain mapping confirmed that this interaction is G-patch domain dependent. Together, these findings identify Cwf28 as a component of catalytically active spliceosomes and suggest a potential role for Cwf28 in modulating the activity of the RNA helicase Cdc28, providing insight into conserved mechanisms underlying RNA helicase regulation during splicing.

Citation: Karika LO, Cipakova I, Kretova M, Selicky T, Cmikova N, Nemcekova D, et al. (2026) The G-patch protein Cwf28 interacts with the RNA helicase Cdc28 in catalytically active spliceosomes in Schizosaccharomyces pombe. PLoS One 21(9): e0357276. https://doi.org/10.1371/journal.pone.0357276

Editor: Claude Prigent, Centre de Recherche en Biologie cellulaire de Montpellier, FRANCE

Received: June 30, 2026; Accepted: August 14, 2026; Published: September 3, 2026

Copyright: © 2026 Karika et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data for this study are publicly available from the ProteomeXchange repository (http://doi.org/10.6019/PXD080271).

Funding: This work was supported by the EU NextGenerationEU through the Recovery and Resilience Plan for Slovakia under the project No. 09I03-03-V04-00445 (LC), by the Slovak Research and Development Agency, Contract No. APVV-23-0205 (LC), and by the Slovak Grant Agency VEGA grant 2/0059/23 (IC) and 2/0056/26 (LC). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Pre-mRNA splicing removes introns from precursor transcripts and ligates exons to form mature messenger RNA (mRNA). This process is catalyzed by the spliceosome, a highly dynamic ribonucleoprotein complex composed of small nuclear ribonucleoproteins (snRNPs) and numerous auxiliary proteins [1]. Splicing is initiated by recognition of the 5′ splice site and branch point by early spliceosomal components, forming an initial assembly complex. ATP-dependent recruitment of additional snRNPs then occurs, promoting progression from the early (E) commitment complex to the assembly of the precatalytic B complex. Extensive rearrangements then drive spliceosome activation, including remodeling of RNA–RNA and RNA–protein interactions that position the splice sites for catalysis. The first transesterification reaction cleaves the 5′ exon and forms a lariat intermediate, while the second reaction ligates the exons and releases the intron. Finally, the spliceosome is disassembled, allowing its components to be recycled for subsequent rounds of splicing [1,2].

In the fission yeast Schizosaccharomyces pombe, a substantial fraction of genes contains multiple introns with splice-site features resembling those of metazoans. Spliceosome assembly in this organism follows an evolutionarily conserved pathway, progressing from E complex formation through recruitment of the U2 snRNP and the U4/U6 × U5 tri-snRNP to yield a catalytically competent spliceosome. Genome-wide analyses in S. pombe suggest relatively rapid and less stringent early spliceosome assembly, with fidelity control mechanisms acting during initial splice-site recognition, as observed in other eukaryotes [3–6]. For example, the Gpl1–Gih35 complex (homologous to the human GPATCH1–DHX35 complex) has been shown to associate with aberrant spliceosomes in S. pombe, where it inhibits splicing progression and promotes their targeting to a discard pathway, thereby contributing to spliceosome quality control [7–9].

RNA helicases play central roles in regulating pre-mRNA splicing by driving the dynamic rearrangements required for spliceosome assembly, activation, and fidelity control [10]. In human cells, helicases such as DHX16, SNRNP200, and DHX38 act in a coordinated manner to promote structural transitions within the spliceosome, including the release of proofreading factors, repositioning of spliceosomal components, and formation of the catalytically active RNA network [2,11,12]. Similarly, in Saccharomyces cerevisiae, the helicase Prp2 (orthologous to DHX16) remodels the B^act complex to generate the catalytically competent B* complex required for the first catalytic step [13,14]. In S. pombe, the helicase Cdc28 is predicted to be the ortholog of DHX16/Prp2 [15], although its precise role in the catalytic cycle remains to be fully defined.

G-patch proteins have emerged as critical regulatory cofactors that modulate the activity of RNA helicases involved in pre-mRNA splicing and ribosome biogenesis [10,14,16–19]. In human cells, several DEAH-box RNA helicases interact with G-patch domain–containing proteins, often in a selective manner. For example, DHX16 and DHX35 appear to associate predominantly with single G-patch partners, GPKOW [20] and GPATCH1 [9], respectively, whereas DHX15 exhibits a broader interaction spectrum, forming complexes with multiple G-patch proteins, including GPATCH2, GPATCH3, GPATCH8, RBM5, SUGP1, TFIP11, and ZGPAT [17,18,21–26]. These interaction patterns underscore both the specificity and versatility of RNA helicase regulation and reflect their participation in multiple RNA processing pathways and distinct stages of spliceosome assembly and remodeling.

Consistent with these observations, regulatory interactions between spliceosomal RNA helicases and G-patch proteins are conserved in yeast. In S. cerevisiae, the G-patch proteins Ntr1 and Cmg1 associate with Prp43, the functional ortholog of human DHX15, and contribute to its regulation [27,28]. In contrast, the G-patch protein Spp2 is an important cofactor of the DEAH-box helicase Prp2, binding its C-terminal region through the G-patch domain and stimulating Prp2 ATPase activity. Although Spp2 is not required for Prp2 recruitment or ATP hydrolysis within the B^act spliceosome, it is essential for coupling Prp2 activity to productive spliceosomal remodeling and the transition from the B^act to the catalytically active B* state [13,14,29,30]. In S. pombe, analogous relationships have been described, including the functional coupling of Ntr1 and Gpl1 with the DEAH-box helicases Prp43 and Gih35, respectively, contributing to spliceosome disassembly, remodeling, and proofreading [7,8,31]. More recently, the S. pombe G-patch protein Sap34 (ortholog of human GPATCH11) has been identified in early spliceosomal complexes, where it associates with Prp43 and may regulate its proofreading activity [32]. Together, these findings highlight the evolutionarily conserved role of G-patch proteins as key modulators of spliceosomal RNA helicases.

In humans, the orthologous protein GPKOW has been proposed to function as a cofactor of DHX16 [10,17,18,20], but the mechanistic evidence supporting this interaction remains less extensive than that established for the S. cerevisiae Spp2–Prp2 pair. In S. pombe, the poorly characterized G-patch protein Cwf28 (ortholog of GPKOW/Spp2) has been reported to co-purify with components of the Prp19 complex (NTC), including Cdc5, as well as the G10 protein Cwf14 [33,34], but its molecular function and interaction partners remain largely unknown.

To elucidate the molecular function of the conserved and essential G-patch domain–containing protein Cwf28 in S. pombe, we performed affinity purification of Cwf28-containing complexes followed by comprehensive interactome analysis. This analysis revealed the RNA helicase Cdc28 as a major interactor of Cwf28, along with components of the NTC and factors involved in the catalytic activation and progression of the spliceosome. Subsequent protein–protein and domain interaction analyses demonstrated that Cwf28 associates with Cdc28 via its conserved G-patch domain, a motif commonly implicated in modulating RNA helicase activity. Collectively, these findings support a model in which Cwf28 associates with Cdc28 during pre-mRNA splicing and suggest a potential role for this interaction in regulating Cdc28 function.

Materials and methods

Strains and media

The S. pombe strain expressing TAP-tagged Cwf28 protein (h^- cwf28-TAP::KanMX4) was generated as described previously [35]. The primers used to construct the tagging plasmid are listed in S1 Table. Cells were grown in complete YE + 5S medium at 32 °C (5 g/L yeast extract, 3% (w/v) glucose, 0.1 g/L L-leucine, 0.1 g/L L-lysine, 0.1 g/L L-histidine, 0.1 g/L uracil, and 0.15 g/L adenine). For a solid medium, 2% (w/v) agar was added. During selection, geneticin was added to the YE + 5S medium at 150 µg/mL. The E. coli strain BTH101 was cultured at 30 °C in Luria–Bertani (LB) medium, composed of 10 g/L tryptone, 5 g/L yeast extract, and 10 g/L NaCl.

Tandem affinity purification

Cells expressing Cwf28-TAP protein were grown in 14 L of YE + 5S medium at 32 °C to mid-log phase (OD₅₉₅ = 0.7–0.8) and harvested by centrifugation (4,000 × g, 10 min, 4 °C). Cells were frozen in liquid nitrogen, and yeast cell powder (30 g) was generated using a SPEX SamplePrep 6770 Freezer/Mill (SPEX SamplePrep). Proteins were extracted in IPP150 buffer (50 mM Tris-HCl, pH 8.0; 150 mM NaCl; 10% glycerol; 0.1% NP-40). The buffer was supplemented with 1 mM PMSF, protease inhibitors (cOmplete, EDTA-free; Roche), and phosphatase inhibitor mix (20 mM NaF; 5 mM Na-pyrophosphate; 10 mM β-glycerophosphate; 1 mM Na₃VO₄). Yeast cell powder was resuspended at a ratio of 1 g per 4 mL of IPP150 buffer. The protein extract was clarified by centrifugation (40,000 × g, 3 × 10 min, 4 °C) and subjected to tandem affinity purification as described previously [36]. Briefly, 400 µL of IgG Sepharose 6 Fast Flow beads were washed in IPP150 buffer and incubated with protein extract for 2 h at 4 °C with gentle rotation. Subsequently, the beads with bound proteins were washed with 20 bead volumes of IPP150 buffer and equilibrated with five bead volumes of TEV cleavage buffer (TCB: 10 mM Tris-HCl, pH 8; 150 mM NaCl; 10% glycerol; 0.1% NP-40; 0.5 mM EDTA; 1 mM DTT). On-bead cleavage was performed in 2 mL of TCB supplemented with 400 U of Turbo TEV protease for 2 h at 16 °C. The resulting eluate was supplemented with 6 µL of 1 M CaCl₂ and diluted with 6 mL of Calmodulin Binding Buffer 1 (CBB1: 10 mM Tris-HCl, pH 8; 150 mM NaCl; 10% glycerol; 0.1% NP-40; 1 mM imidazole; 1 mM Mg(OAc)₂; 2 mM CaCl₂; 10 mM β-mercaptoethanol). Subsequently, 100 µL of Calmodulin Sepharose 4B beads washed in CBB1 were added, and binding was allowed for 2 h at 4 °C with gentle rotation. Beads were washed with 10 bead volumes of CBB1 followed by five bead volumes of Calmodulin Binding Buffer 2 (10 mM Tris-HCl, pH 8; 150 mM NaCl; 1 mM Mg(OAc)₂; 2 mM CaCl₂; 1 mM β-mercaptoethanol). Proteins were eluted stepwise using one bead volume of elution buffer (10 mM Tris-HCl, pH 8; 150 mM NaCl; 1 mM Mg(OAc)₂; 2 mM EGTA; 1 mM β-mercaptoethanol). Protein eluates were analyzed by 8% SDS–PAGE and visualized by silver staining [37]. Protein-containing eluates were pooled and subjected to LC–MS/MS analysis.

Mass spectrometry analysis

The samples were reduced with 5 mM DTT, then alkylated with 15 mM iodoacetamide, and the reaction was quenched with an additional 5 mM DTT. Digestion with trypsin (1:25 w/w) was performed overnight (16 h) in the presence of phosphatase inhibitors at 37 °C. The microtip C18 SPE was used to purify the peptides. For liquid chromatography-coupled mass spectrometry on a Dionex UltiMate 3000 RSLC nano system (Thermo Scientific) and Orbitrap Elite mass spectrometer (Thermo Scientific) peptides were loaded onto a PepMap100 C18 trap column (300 μm × 5 mm, 5-μm particle size, Thermo Scientific), separated with the EASY-Spray PepMap RLSC C18 analytical column with an integrated nanospray emitter (75 μm × 500 mm, 2-μm particle size, Thermo Scientific) using a 60-min gradient (2.4%–34.4% acetonitrile) at a flow rate 250 nL/min and sprayed directly into the mass spectrometer. Precursors were measured in the mass range 300–1,700 m/z with a resolution of 120,000 and selected for fragmentation in a data-dependent mode using the Top15 strategy. The fragmentation was performed by the HCD mechanism with a normalized collision energy of 25%, and MS/MS scans were acquired with a resolution of 15,000. Each sample was measured in technical duplicates. The datasets were processed by MaxQuant [38] version 2.7.0.0. Carbamidomethylation (C) was set as a permanent modification, and oxidation (M), acetylation (protein N-terminus), and phosphorylation (STY) were set as variable modifications. The search was performed against S. pombe proteomes from UniProt (downloaded 9.3.2026, isoforms included) and PomBase (downloaded 11.3.2026). Identified proteins and phosphosites with localization probability > 0.75 are listed in the S1 Table. Proteins are arranged in descending order from the highest iBAQ value (intensity normalized in order to be proportional to copy number and not to protein mass). To assess whether the identified phosphosites may represent potential CK2 target sites, phosphorylation site predictions were performed using NetPhos 3.1 [39].

Functional analysis of Cwf28 interactome

Gene Ontology (GO) term enrichment was performed using the STRING database [40]. The protein–protein interaction network among proteins co-purified with Cwf28 was constructed with STRING v12.0 (http://string-db.org) using the high-confidence setting (0.7).

Mutagenesis of the cwf28 allele

PCR-based random mutagenesis was used to introduce mutations within the cwf28 allele [41]. Mutagenesis was performed using 50 ng of cwf28-pUT18 plasmid as the template in a 50 μL reaction containing 1 × Standard Taq buffer (10 mM Tris-HCl, pH 8.3, 50 mM KCl, 1.5 mM MgCl₂), supplemented with an additional 7 mM MgCl₂, 1 mM each of dCTP and dTTP, 0.2 mM each of dATP and dGTP, 2 μM of primers (5′-CGACAGGTTTCCCGACTGG-3′ and 5′-GAAGTTACAGAAGTACTCGTACACTTCCCCGGG-3′), 0.05 U Taq DNA polymerase, and 0.5 mM MnCl₂ added immediately before PCR. Thermal cycling conditions were as follows: initial denaturation at 95 °C for 3 min, 12 cycles of 95 °C for 1 min, 56 °C for 1 min, and 68 °C for 3 min, followed by a final extension at 68 °C for 10 min. Following mutagenesis, PCR products were digested with SmaI and PstI and cloned into the pUT18 vector. Plasmids from positive transformants were isolated and sequenced to identify mutations in the cwf28 allele. Subsequently, plasmids encoding single amino acid substitutions within the G-patch domain were used for protein–protein interaction analysis.

Protein–protein interaction analysis

The sequences encoding the full-length ORFs of Cdc28, Cwf28, specific Cwf28 constructs, or G-patch domains engineered to encode defined single amino acid substitutions were created using specific primers (S1 Table) and subcloned or assembled in-frame with the T18 and T25 fragments of adenylate cyclase into the pUT18 and pKNT25 BACTH vectors using the NEBuilder HiFi DNA Assembly Cloning Kit [42]. The prepared plasmids were propagated in E. coli DH5α, purified, and co-transformed into E. coli BTH101 (10 ng of each plasmid). Positive transformants were screened on LB plates supplemented with 40 µg/mL X-gal, 0.5 mM IPTG, 100 µg/mL ampicillin, and 50 µg/mL kanamycin (at 30 °C). Transformants were then spotted onto fresh LB plates supplemented with 40 µg/mL X-gal, 0.5 mM IPTG, 100 µg/mL ampicillin, and 50 µg/mL kanamycin, and incubated for 24 h at 30 °C.

β-galactosidase activity assay

β-galactosidase activity assay was measured at 30 °C on stationary-phase aliquots of cultures in 3 mL of LB medium in the presence of 0.5 mM IPTG and antibiotics as described previously [43]. The optical density (OD₆₀₀) of each culture was recorded. Cells were permeabilized by adding 15 µL of toluene and 15 µL of 0.1% SDS solution to 1.25 mL of culture, vortexed, plugged with cotton, and vigorously agitated for 40 min at 37 °C. Aliquots (0.1 mL) of permeabilized cells were mixed with 0.9 mL of PM2 assay buffer (60 mM Na₂HPO₄, 40 mM NaH₂PO₄, 0.14 mM MnCl₂, and 1 mM MgSO₄, pH 7.0) supplemented with 7 µL of β-mercaptoethanol (final concentration 100 mM) and preincubated for 5 min at 30 °C. PM2 assay buffer served as a blank. The reaction was initiated by adding 200 µL of 0.4% (w/v) ONPG, followed by incubation at 30 °C. The reaction was allowed to proceed until a visible yellow color developed, and the reaction time was recorded. The reaction was then terminated by adding 200 µL of 1 M Na₂CO₃. OD₄₂₀ and OD₅₅₀ were measured with an S-200 Spectrophotometer (Boeco, Germany). β-galactosidase activity was expressed as Miller units using the following equation: Miller Units = 1000 × [(OD₄₂₀–1.75 × OD₅₅₀)]/(t × V × OD₆₀₀), where OD₄₂₀ is the absorbance of the yellow o-nitrophenol, OD₅₅₀ is the scatter from cell debris, which, when multiplied by 1.75 approximates the scatter observed at 420 nm, OD₆₀₀ reflects cell density, t is reaction time in min and V is a volume of culture assayed in mL. The Miller Units give the change in OD₄₂₀/min/mL of cells/OD₆₀₀.

Statistical analysis

Statistical significance of interactions between Cdc28, Cwf28, and Cwf28 G-patch domain constructs was evaluated relative to the full-length Cwf28 construct using an unpaired t-test in Microsoft Excel. p-values <0.05 were considered statistically significant.

AlphaFold modeling

The amino acid sequences of Cdc28 and Cwf28 were used as input for AlphaFold-Multimer to predict the structure of their protein complex [44,45]. The predicted model was subsequently visualized and analyzed in PyMOL (v3.1) to assess structural features and potential interaction interfaces between the proteins [46].

Results and discussion

Despite the well-established importance of G-patch proteins in modulating RNA helicase activity, the precise molecular functions of many members of this family remain incompletely understood. We therefore sought to determine whether the poorly characterized and essential fission yeast G-patch protein Cwf28 operates within a regulatory framework analogous to that described for its orthologs, S. cerevisiae Spp2 and human GPKOW [13,14,20,29,30].

To address this, we purified native Cwf28-containing complexes by tandem affinity purification and conducted comprehensive interactome analysis to identify associated factors (Fig 1A). Our results show that Cwf28 predominantly co-purifies with components of the NTC as well as proteins involved in the catalytic activation and progression of the spliceosome. Notably, the DEAH-box RNA helicase Cdc28, the ortholog of human DHX16 and S. cerevisiae Prp2, was identified as the most abundant and prominent interactor of Cwf28 (Fig 1B and S1 Table).

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Fig 1. Identification of proteins co-purifying with G-patch protein Cwf28.

(A) Proteins associated with Cwf28 were isolated by tandem affinity purification. The complexity of eluates (E₁–E₈) was analyzed by SDS-PAGE, and proteins were visualized by silver staining. Molecular weight marker (M) is indicated on the left. The positions corresponding to Cdc28 (~121 kDa) and the Cwf28-CBP fusion protein are indicated. (B) Selected proteins co-purifying with Cwf28 are shown. Proteins are ranked by their intensity-based absolute quantification (iBAQ) values from two independent biological replicates, providing an estimate of their relative abundance in the Cwf28 pulldown. Corresponding S. cerevisiae and human orthologs are indicated. For the complete list of identified proteins and quantitative information, see S1 Table.

https://doi.org/10.1371/journal.pone.0357276.g001

Consistent with previous characterization and annotation of spliceosomal components in S. pombe [15], only a limited subset of spliceosome-associated factors, including NTC-associated proteins and U2-type catalytic spliceosome-associated factors (Cwf16, Cwf19, Cwf25, and Mug161), were not detected among the proteins co-purifying with Cwf28. In addition, components of the casein kinase 2 (CK2) complex, including the catalytic subunit Cka1 and the regulatory subunits Ckb1 and Ckb2 [47], were identified in the Cwf28 interactome (S1 Table). The presence of the CK2 complex agrees with earlier studies demonstrating its association with spliceosomal assemblies in S. pombe, where it is thought to regulate the activity of spliceosome-associated factors through phosphorylation [48–51].

Gene Ontology (GO) analysis revealed significant enrichment of proteins associated with biological processes related to pre-mRNA splicing, including spliceosomal conformational rearrangements leading to catalytic activation, spliceosomal complex assembly, mRNA splice site selection, 5′ splice site recognition, and mRNA splicing, among others. In terms of cellular components, the Cwf28 interactome was enriched for proteins associated with the NTC, U2-type catalytic step 1 and step 2 spliceosomes, the protein kinase CK2 complex, and others (Fig 2). Collectively, these results strongly indicate that the G-patch protein Cwf28 is an integral component of catalytically active spliceosomal assemblies and likely functions in coordinating spliceosome activation and remodeling, potentially through regulation of RNA helicase Cdc28.

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Fig 2. GO enrichment analysis of proteins co-purifying with Cwf28.

The scatter plot includes the top 7 most enriched biological process and cellular component categories. The X-axis represents the strength (the enrichment effect), and the Y-axis shows the specific GO terms. The color of the dots represents the FDR values, with light green indicating lower FDR values (higher significance) and blue indicating higher FDR values (lower significance). Dot size corresponds to the number of proteins in the respective GO term.

https://doi.org/10.1371/journal.pone.0357276.g002

Interestingly, multiple phosphorylation sites were identified among proteins co-purifying with Cwf28, including S95, S96, and S138 of Cdc28; S535 of Cip2; S385 of Cwf2; T62, S721, and S853 of Cwf22; S11, S13, S31, S101, S102, S152, S277 of Cwf28; S160 of Cwf29; S51 of Prp10; S39 of Ctnnbl1; and S228 and S236 of Prp45 (S1 Table). The phosphorylation events detected within Cwf28-associated complexes indicate that multiple spliceosomal factors associated with Cwf28 are phosphorylated. Notably, the CK2 complex was identified among the Cwf28-associated proteins, and several of the identified phosphosites correspond to residues predicted to be compatible with CK2 substrate preferences, including S51 of Prp10, S39 of Ctnnbl1, S385 of Cwf2, S721 of Cwf22, S96 and S138 of Cdc28, and S11, S13, and S102 of Cwf28. These observations suggest a potential association between the CK2 complex and phosphorylation of Cwf28-associated spliceosomal components. However, the relative abundance and phosphorylation occupancy of these sites were not determined, and whether these modifications are directly mediated by the CK2 complex or have functional consequences for spliceosome function remains to be established. Future studies will be needed to define the contribution of individual phosphorylation events to spliceosome dynamics, RNA helicase activity, and pre-mRNA splicing efficiency and fidelity.

The identification of the RNA helicase Cdc28 as the most abundant interactor within the Cwf28 complex (Figs 1A and 1B, S1 Table) prompted us to test whether these two proteins physically interact. Previous studies in S. cerevisiae demonstrated that the G-patch protein Spp2 interacts with and regulates the spliceosomal helicase Prp2 [13,14,29,30,52]. This interaction is mediated by the G-patch domain of Spp2 and involves the C-terminal region of Prp2, including the OB-fold motif [30]. To gain structural insight into the interaction between Cwf28 and Cdc28 in S. pombe, we employed AlphaFold-Multimer to predict the architecture of their complex. The resulting model positioned the G-patch domain of Cwf28 at the interface with Cdc28, where it adopts an extended conformation that engages a defined surface within the C-terminal region of the helicase, forming a putative interaction interface enriched in both hydrophobic and polar contacts (Fig 3A). This predicted arrangement resembles the interaction mode described for the S. cerevisiae Spp2–Prp2 complex, in which the G-patch domain contacts the C-terminal region of the helicase [30].

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Fig 3. Analysis of interactions between the G-patch protein Cwf28 and the RNA helicase Cdc28.

(A) AlphaFold-Multimer predicted the three-dimensional structure of the Cwf28–Cdc28 complex. Cwf28 is shown in blue, with the G-patch domain highlighted in red, while Cdc28 is depicted in green. The predicted Cwf28–Cdc28 interaction interface is shown in an enlarged view, with substituted amino acids within the G-patch domain labelled and colour-coded according to their effects on the interaction observed in the BACTH assay. The G-patch domain is shown in red, residues whose substitution abolished the interaction are indicated in cyan, and the residue whose substitution reduced the interaction is indicated in yellow. This model represents a predicted interaction surface that is supported by the BACTH results. (B) Schematic representation of specific Cwf28 constructs. (C) Alleles encoding Cdc28, Cwf28, and specific Cwf28 constructs were cloned into the pUT18 and pKNT25 plasmids of the BACTH assay. Empty pUT18 and pKNT25 vectors were used as negative controls, while pUT18C-zip and pKT25-zip plasmids served as positive controls. (D) Interaction strengths between the studied proteins and constructs were assessed by β-galactosidase activity assay and quantified in Miller units. Data represent mean ± S.D. from six independent biological replicates. (E) Schematic representation of specific Cwf28 G-patch domain mutants. Introduced single amino acid substitutions in the G-patch domain are indicated in red. Conserved glycine residues are highlighted in grey. (F) Alleles encoding Cdc28, Cwf28, and Cwf28 constructs containing specific single amino acid substitutions within the G-patch domain were assembled into the pUT18 and pKNT25 plasmids of the BACTH assay. Empty pUT18 and pKNT25 vectors were used as negative controls. (G) Interaction strengths between the studied proteins and constructs were assessed by β-galactosidase activity assay and quantified in Miller units. Data represent mean ± S.D. from six independent biological replicates. Note: Statistical significance of interactions between Cdc28 and Cwf28 G-patch domain constructs was evaluated relative to the full-length Cwf28 construct using an unpaired t-test (***p < 0.001).

https://doi.org/10.1371/journal.pone.0357276.g003

To experimentally validate this predicted interaction, we employed a bacterial adenylate cyclase two-hybrid (BACTH) assay. Consistent with the structural model, the BACTH results confirmed an interaction between Cwf28 and Cdc28 and further demonstrated that the G-patch domain is necessary and sufficient for this association. Notably, Cdc28 interaction was retained with all Cwf28-derived constructs containing the G-patch domain, including the N-terminal region encompassing the G-patch, the G-patch fused to the C-terminal region, and the G-patch domain alone. In contrast, no interaction was detected with Cwf28 constructs comprising only the N-terminal or C-terminal regions lacking the G-patch domain. These findings support the conclusion that the G-patch domain of Cwf28 is necessary for mediating interaction with Cdc28 (Figs 3B, 3C, and 3D).

To further elucidate the molecular basis of the interaction between the G-patch domain of Cwf28 and Cdc28, we generated a series of point mutations targeting specific residues within the G-patch sequence. Specifically, single amino acid substitutions were introduced at positions predicted to contribute to protein–protein interactions: W206R and N207S (immediately following the third conserved glycine), Q209R (following the fourth glycine), and G226S (fifth glycine) and M227K (adjacent to the fifth glycine residue) (Fig 3E). Assessing the impact of these substitutions on the interaction with Cdc28, we found that the G226S mutation resulted in a marked reduction in interaction strength, whereas the W206R, Q209R, and M227K substitutions abolished the interaction. In contrast, the N207S substitution did not produce a significant change in interaction compared to the wild-type G-patch domain (Figs 3F and 3G). The disruptive effects of W206R, Q209R, and M227K substitutions can be explained by alterations in physicochemical properties. Replacement of hydrophobic or aromatic residues (e.g., tryptophan at position 206 and methionine at position 227) with positively charged residues (arginine or lysine) likely disrupts hydrophobic contacts essential for stable binding. Additionally, substitution of glutamine with arginine (Q209R) introduces a charged side chain that may perturb local hydrogen-bonding networks or introduce electrostatic repulsion at the interaction interface. The moderate effect observed for G226S may reflect the importance of glycine in maintaining backbone flexibility. Its substitution with serine could restrict conformational adaptability required for optimal binding. In contrast, the N207S substitution represents a conservative change between polar residues, likely preserving hydrogen-bonding capacity and the overall structural integrity of the interaction interface. Although these results support the importance of specific G-patch residues in mediating the Cwf28–Cdc28 interaction, we cannot completely exclude the possibility that differences in fusion protein abundance may contribute to the observed interaction phenotypes, as a general limitation of the BACTH assay.

Collectively, the BACTH analysis demonstrates that the interaction between Cwf28 and Cdc28 is mediated specifically by the G-patch domain of Cwf28, supporting a model in which Cwf28 directly engages the RNA helicase Cdc28 through a defined G-patch-dependent interface. Consistent with both the mutational analysis and structural predictions, the G-patch domain of Cwf28 provides the essential interaction surface required for formation of the Cwf28–Cdc28 complex. This mode of interaction is consistent with previous studies in S. cerevisiae demonstrating that the G-patch protein Spp2 interacts with and modulates the enzymatic properties of the spliceosomal helicase Prp2 [13,14,29,30,52], similar to the regulation of the helicase Prp43 by the G-patch proteins Ntr1 and Pfa1 [28,53–55].

Given the established role of G-patch proteins as cofactors of DEAH-box helicases, our findings suggest that Cwf28 may contribute to the regulation or modulation of Cdc28 function during spliceosome progression in S. pombe. However, the direct impact of Cwf28 on Cdc28 activity remains to be determined. Thus, while our study positions Cwf28 in close association with catalytically active spliceosomal complexes and supports a functional relationship between Cwf28 and Cdc28, further biochemical and functional studies will be required to establish whether Cwf28 acts as a regulatory cofactor of the Cdc28 helicase. In particular, it will be important to determine whether the Cwf28–Cdc28 interaction is required for efficient and accurate pre-mRNA splicing, and how its disruption affects spliceosome dynamics, RNA processing fidelity, and overall cellular homeostasis. Future studies combining in vivo functional assays with high-resolution structural and biochemical approaches will be essential to further define the mechanistic contribution of Cwf28 to spliceosome function.

Supporting information

S1 Table. Primers, proteins identified in Cwf28-TAP purification, and their associated phosphorylation sites.

https://doi.org/10.1371/journal.pone.0357276.s001

(XLSX)

S1 Fig. Original, uncropped images corresponding to Figs. 1A, 3C, and 3F.

https://doi.org/10.1371/journal.pone.0357276.s002

(TIF)

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Figures

Abstract

Introduction

Materials and methods

Strains and media

Tandem affinity purification

Mass spectrometry analysis

Functional analysis of Cwf28 interactome

Mutagenesis of the cwf28 allele

Protein–protein interaction analysis

β-galactosidase activity assay

Statistical analysis

AlphaFold modeling

Results and discussion

Supporting information

S1 Table. Primers, proteins identified in Cwf28-TAP purification, and their associated phosphorylation sites.

S1 Fig. Original, uncropped images corresponding to Figs. 1A, 3C, and 3F.

References