Fip1 is a multivalent interaction scaffold for processing factors in human mRNA 3′ end biogenesis

  1. Lena Maria Muckenfuss
  2. Anabel Carmen Migenda Herranz
  3. Franziska Maria Boneberg
  4. Marcello Clerici
  5. Martin Jinek  Is a corresponding author
  1. Department of Biochemistry, University of Zurich, Switzerland

Abstract

3′ end formation of most eukaryotic mRNAs is dependent on the assembly of a ~1.5 MDa multiprotein complex, that catalyzes the coupled reaction of pre-mRNA cleavage and polyadenylation. In mammals, the cleavage and polyadenylation specificity factor (CPSF) constitutes the core of the 3′ end processing machinery onto which the remaining factors, including cleavage stimulation factor (CstF) and poly(A) polymerase (PAP), assemble. These interactions are mediated by Fip1, a CPSF subunit characterized by high degree of intrinsic disorder. Here, we report two crystal structures revealing the interactions of human Fip1 (hFip1) with CPSF30 and CstF77. We demonstrate that CPSF contains two copies of hFip1, each binding to the zinc finger (ZF) domains 4 and 5 of CPSF30. Using polyadenylation assays we show that the two hFip1 copies are functionally redundant in recruiting one copy of PAP, thereby increasing the processivity of RNA polyadenylation. We further show that the interaction between hFip1 and CstF77 is mediated via a short motif in the N-terminal ‘acidic’ region of hFip1. In turn, CstF77 competitively inhibits CPSF-dependent PAP recruitment and 3′ polyadenylation. Taken together, these results provide a structural basis for the multivalent scaffolding and regulatory functions of hFip1 in 3′ end processing.

Editor's evaluation

This study explores the structural and biochemical basis for Fip1 interactions within the cleavage and polyadenylation machinery – notably with CPSF30 and CstF77. Overall, the significance of the study is that it provides valuable mechanistic insight into the function of Fip1 in the cleavage and polyadenylation machinery. The data presented in the paper are compelling and the authors use a combination of structural biology and biochemistry to present their case. This study will be of interest to those focusing on mRNA biosynthesis and the biophysical properties of RNA binding proteins.

https://doi.org/10.7554/eLife.80332.sa0

Introduction

3′ end polyadenylation is a fundamental process in eukaryotic messenger RNA (mRNA) biogenesis, essential for the maturation of non-histone precursor mRNAs (pre-mRNAs) prior to their export into the cytoplasm. Poly(A) tails possess key functions in mRNA metabolism, governing mRNA export, translational efficiency, and stability (Nicholson and Pasquinelli, 2019; Passmore and Coller, 2022). Furthermore, alternative mRNA polyadenylation (APA) constitutes a key mechanism of gene expression control through dynamic regulation of polyadenylation site selection in pre-mRNA transcripts (Di Giammartino et al., 2011; Tian and Manley, 2016). Accordingly, defects in polyadenylation are linked to human diseases such as cancer, β-thalessemia, diabetes, or systemic lupus (Hollerer et al., 2014; Gruber and Zavolan, 2019; Dharmalingam et al., 2022). mRNA 3′ end biogenesis occurs by a two-step mechanism comprising endonucleolytic cleavage of the pre-mRNA transcript by the cleavage and polyadenylation specificity factor (CPSF) complex and subsequent polyadenylation of the free 3′ end by the poly(A) polymerase (PAP). In human cells, the process is dependent on the controlled assembly of several protein factors on the pre-mRNA, including CPSF, RBBP6, cleavage stimulation factor (CstF), as well as mammalian cleavage factors I and II (CF Im and CFIIm, respectively), and PAP (Zhao et al., 1999; Xiang et al., 2014; Kumar et al., 2019; Boreikaite et al., 2022; Schmidt et al., 2022). Most of these protein factors are highly conserved between mammals and yeast, underlining the fundamental nature of this process (Xiang et al., 2014). The cleavage site, typically downstream of a CA dinucleotide, is defined by the polyadenylation signal (PAS), a conserved hexanucleotide motif (predominantly AAUAAA) located approximately 10–30 nucleotides upstream (Proudfoot and Brownlee, 1976; Proudfoot, 2011).

The PAS is specifically recognized by CPSF (Chan et al., 2014; Schönemann et al., 2014; Clerici et al., 2018; Sun et al., 2018), which consists of two functional modules: the mammalian polyadenylation specificity factor (mPSF) comprising subunits CPSF160, WDR33, CPSF30, and hFip1 (for human factor interacting with poly(A) polymerase 1) (Bienroth et al., 1991; Murthy and Manley, 1992; Kaufmann et al., 2004; Shi et al., 2009), and the mammalian cleavage factor (mCF) containing the endonuclease CPSF73 (Mandel et al., 2006), CPSF100 as well as Symplekin (Sullivan et al., 2009). RBBP6 associates with mCF and is essential for pre-mRNA cleavage (Di Giammartino et al., 2014; Boreikaite et al., 2022; Schmidt et al., 2022). Within mPSF, the CPSF160–WDR33 subcomplex forms a rigid scaffold (Clerici et al., 2017) that interacts with CPSF30 (Clerici et al., 2018; Sun et al., 2018; Zhang et al., 2019) and the CPSF100 subunit of mCF (Zhang et al., 2019). CPSF30 contains five C3H1-type zinc finger (ZF) domains and a C-terminal zinc knuckle domain which is absent in yeast homolog Yth1 (Barabino et al., 1997) and not required for mPSF complex assembly (Clerici et al., 2017). The ZF1 domain is necessary and sufficient for binding to the CPSF160–WDR33 heterodimer, while ZF2 and ZF3 together with WDR33 mediate recognition of the AAUAAA PAS hexamer motif (Clerici et al., 2018; Sun et al., 2018). ZF4 and ZF5 domains interact with hFip1 (Barabino et al., 2000; Hamilton and Tong, 2020). hFip1 is an important regulator of APA that contributes to cleavage site selection through its interaction with CFIm via its C-terminal arginine/serine-rich (RS) domain (Zhu et al., 2018) and additionally by binding to U-rich regions in the pre-mRNA via an arginine-rich C-terminal region (Kaufmann et al., 2004), thereby specifically promoting polyadenylation of mRNA substrates with U-rich sequences preceding the AAUAAA hexanucleotide (Lackford et al., 2014). Both the RS domain and the arginine-rich region are absent in yeast Fip1 and the alternatively spliced isoform 4 of hFip1.

In previously determined cryo-EM structures of the yeast CPF and human mPSF complexes, ZF4 and ZF5 remained unresolved (Casañal et al., 2017; Clerici et al., 2018; Sun et al., 2018), indicating conformational flexibility with respect to the rigid mPSF core. Recently, a crystal structure of human CPSF30 ZF4–5 domains in complex with hFip1 has been determined (Hamilton and Tong, 2020) and complementary NMR studies of the yeast Fip1 homolog Kumar et al., 2021 have shed light on the molecular details of the CPSF30–Fip1 interaction and revealed considerable structural dynamics of Fip1 in the context of the 3′ processing machinery.

Mammalian CstF is a dimer of trimers comprising CstF77, CstF64, and CstF50 subunits (Takagaki et al., 1990; Yang et al., 2018). It is recruited to the pre-mRNA by U- and G/U-rich sequences located downstream of the cleavage site (Takagaki and Manley, 1997) that are recognized by CstF64 (Takagaki et al., 1992; MacDonald et al., 1994). Through stabilization of CPSF on the pre-mRNA, CstF plays an important role in PAS recognition and is essential for pre-mRNA cleavage (Takagaki et al., 1990; Boreikaite et al., 2022; Schmidt et al., 2022). Dimerization of CstF is mediated by the CstF77 HAT (half-a-tetratricopeptide repeat) domain homodimer (Bai et al., 2007), and further stabilized by CstF50 (Yang et al., 2018). The CstF77 homodimer has an arch-like shape and interacts asymmetrically with CPSF, contacting the CPSF160–WDR33 mPSF scaffold via only one side of the arch (Zhang et al., 2019).

Fip1 interacts with PAP and tethers it to CPSF bound near the nascent 3′ end of the cleaved pre-mRNA, which is required for its processive polyadenylation (Preker et al., 1995; Helmling et al., 2001; Kaufmann et al., 2004; Meinke et al., 2008; Ezeokonkwo et al., 2011). Besides CPSF30 and PAP, biochemical and cellular studies have implicated Fip1 in interactions with other proteins including CPSF160, CstF77 (Preker et al., 1995; Kaufmann et al., 2004), WDR33 (Ohnacker et al., 2000; Clerici et al., 2017), Symplekin (Ghazy et al., 2009), and CF Im (Venkataraman et al., 2005). However, the molecular details of these interactions have not yet been revealed.

Here, we report structural and biochemical analysis of the interactions of hFip1 with CPSF30, PAP, and CstF77 within the human 3′ polyadenylation machinery. While confirming previous structural data (Hamilton and Tong, 2020), we notably show that mPSF contains two hFip1 copies, yet recruits only one PAP molecule at a time. The presence of two PAP-binding sites in mPSF contributes to the processivity of 3′ polyadenylation. Furthermore, we show that hFip1 interacts with CstF77 through a conserved helix in its N-terminal ‘acidic’ region and reveal that CstF77 competes with PAP for hFip1 binding, which attenuates polyadenylation efficiency. These results deepen our understanding of hFip1 as a key interaction partner for 3′ end processing factors, facilitating or regulating their spatiotemporal assembly on the pre-mRNA, and establish a framework for further mechanistic studies of hFip1 interactions and CstF-mediated regulation of mRNA 3′ end biogenesis.

Results

Structural basis for the human hFip1–CPSF30 interaction

The ZF4 and ZF5 domains of CPSF30 are necessary and sufficient for the interaction with the conserved central domain of hFip1 (hereafter referred to as hFip1CD) (Clerici et al., 2017; Hamilton and Tong, 2020). Yet these domains could not be resolved in previously determined cryo-EM reconstructions of the human mPSF (Clerici et al., 2018; Sun et al., 2018), indicating that they are likely flexibly tethered to the mPSF core. To gain insights into the CPSF30–hFip1 interaction, we determined a crystal structure of a CPSF30 fragment spanning ZF4 and ZF5 domains (CPSF30ZF4–ZF5, residues 118–178) in complex with hFip1CD (residues 138–180 of hFip1 isoform 4) at a resolution of 2.2 Å (Figure 1B). The structure reveals that hFip1CD binds CPSF30 in a 2:1 stoichiometry, with one hFip1CD molecule (hFip1CDa) binding predominantly to ZF4 and the other (hFip1CDb) to ZF5. While the overall conformation of the hFip1–CPSF30 complex is highly similar to that of a recent crystal structure of human hFip1–CPSF30 (Hamilton and Tong, 2020), with a root-mean-square deviation of 0.82 Å over the whole complex, in this study, 21 additional residues of hFip1 could be resolved in hFip1CDb due to extended crystallization construct boundaries, with 16 additional residues at the N-terminus (residues 130–145, isoform 4) and 5 additional residues at the C-terminus (residues 177–181, isoform 4). Structural superpositions reveal that hFip1CDa and hFip1CDb bind to the same surfaces of ZF4 and ZF5 domains with a root-mean-square deviation of 0.87 Å over 54 aligned residues (Figure 1C). Moreover, superpositions with CPSF30 ZF2 and ZF3 domains reveal that the interaction surfaces on ZF4 and ZF5 are located on the opposite side of the ZF fold relative to the PAS RNA-binding surfaces of ZF2 and ZF3 (Figure 1C). ZF2 and ZF3 interactions with the RNA are mainly mediated by π–π stacking of aromatic side chains with nucleobases and supplemented by protein mainchain hydrogen bond interactions (Clerici et al., 2018; Sun et al., 2018). Although the aromatic residues are conserved in ZF4 and ZF5 (Figure 1—figure supplement 1), RNA binding is likely precluded by the presence of proline residues at key mainchain hydrogen-binding positions. The hFip1CDa–CPSF30 interaction surface (803 Å2) is almost twice as large as the hFip1CDb–CPSF30 interface (478 Å2) because hFip1CDa binds at the ZF4–ZF5 junction and has additional contacts with ZF5. hFip1CDa and hFip1CDb also contact each other directly (215 Å2). ZF4 interaction with hFip1CDa is mediated by a hydrophobic interface centered on Lys127CPSF30 and Phe131CPSF30 and supported by additional salt-bridge contact involving Arg144CPSF30 and Asn159hFip1 (Figure 1D). In turn, the interaction of ZF5 with hFip1CDb is mainly mediated by Tyr151CPSF30 and Phe155CPSF30 and supported by a salt-bridge contact between Arg168CPSF30 and Asp159hFip1 (Figure 1E). As Fip1 is conformationally dynamic in isolation (Meinke et al., 2008; Ezeokonkwo et al., 2011; Kumar et al., 2021), CPSF30-binding results in structural ordering of the CD region. Interactions with both ZF4 and ZF5 are mediated by a hydrophobic patch in hFip1CD comprising the aromatic side chains of Trp150hFip1, Phe161hFip1, and Trp170hFip1 (Figure 1D, E).

Figure 1 with 2 supplements see all
hFip1 interacts with CPSF30 with 2:1 stoichiometry.

(A) Schematic representation of the domain architecture of CPSF30 and hFip1. CPSF30 consists of five zinc finger (ZF) domains and a zinc knuckle domain. hFip1 isoform 4 comprises acidic, conserved, and proline-rich regions but lacks the RE/D region interacting with CF Im, as well as the R-rich region, which has been shown to bind U-rich RNA in hFip1 isoform 1 (Kaufmann et al., 2004). (B) Cartoon representation of the crystal structure of CPSF30ZF4–ZF5 in complex with two hFip1 fragments comprising the conserved domain (CD). (C) Superposition of CPSF30 ZF2 domain in complex with PAS RNA onto ZF4 and ZF5. (D) Detailed interaction interface of hFip1CD with CPSF30 ZF4. (E) Detailed interaction interface of hFip1CD with CPSF30 ZF5. (F) Size-exclusion chromatography coupled to multiangle static light scattering (SEC-MALS) chromatogram of MBP-CPSF30ZF4–ZF5 selective hFip1-binding mutants for stoichiometry analysis with GFP-hFip1. (G) In vitro pull-down analysis of FLAG-epitope-tagged mPSF comprising wild-type CPSF30 and its selective hFip1-binding mutants with GFP-PAP. Asterisk indicates anti-FLAG M2 antibody light chain. GFP-hFip1 and GFP-PAP are also visualized with in-gel GFP fluorescence (bottom).

To validate our structural observations, we initially mutated ZF4 and ZF5 interaction surface residues in CPSF30ZF4–ZF5 and tested the interactions of the mutant proteins with hFip1CD in a pull-down assay (Figure 1—figure supplement 2A). Individual substitutions of Tyr127CPSF30, Tyr151CPSF30, or Phe155CPSF30 with glutamate resulted in substantial reduction of hFip1CD binding, while simultaneous mutation of both ZF4 and ZF5 residues resulted in loss of hFip1 binding, in agreement with our structural observations. In hFip1CD, substitution of aromatic residues with glutamate in the hydrophobic interaction patch either substantially reduced (Trp150hFip1 and Trp170hFip1) or completely disrupted (Phe161hFip1) the hFip1CD–CPSF30ZF4–ZF5 interaction (Figure 1—figure supplement 2B). We subsequently performed size-exclusion chromatography coupled to multiangle static light scattering (SEC-MALS) to analyze the stoichiometry of hFip1CD–CPSF30ZF4–ZF5 complexes. hFip1CD and wild-type CPSF30ZF4–ZF5 formed a 2:1 complex. In contrast, CPSF30ZF4–ZF5 proteins containing Y127ECPSF30 or F155ECPSF30 mutations formed a 1:1 complex with hFip1CD, while simultaneous mutation of both residues resulted in complete loss of binding (Figure 1F). Together, these results confirm that human CPSF30 has two independently functional hFip1-binding sites, one on ZF4 and the other on ZF5, each recruiting one copy of hFip1.

Functional redundancy of hFip1–CPSF30 interactions in human CPSF

To probe the functional significance of the dual CPSF30–hFip1 interaction interfaces in the context of human CPSF, we coexpressed wild-type or mutant CPSF30 together with hFip1, WDR33 and FLAG epitope-tagged CPSF160 in baculovirus-infected insect cells, and performed tandem affinity purifications during which purified recombinant catalytic domain of human PAP (residues 1–504) was added in trans after the second affinity purification step. hFip1 copurified with mPSF containing wild-type CPSF30, and PAP was efficiently coprecipitated (Figure 1G). Expression of CPSF30 ZF4 or ZF5 mutants (Y127E or Y151E, respectively) resulted in reduced recovery of both hFip1 and PAP relative to the other mPSF components (Figure 1G), consistent with the reduced stoichiometry of the CPSF30–hFip1 interaction observed in vitro (Figure 1F). In turn, expression of a CPSF30 construct containing mutations in both the ZF4- and ZF5-binding sites resulted in the loss of hFip1 from mPSF, which was thus unable to interact with PAP (Figure 1G). Together, these results indicate that either hFip1-binding site in CPSF30 can contribute to the integrity of mPSF in vivo and both sites are capable of recruiting hFip1 and consequently PAP. Notably, the expression levels of mPSF mutant complexes incapable of binding hFip1 (Y127E/Y151CPSF30) were substantially reduced, consistent with the role of hFip1 in stabilizing the CPSF30 ZF fold (Kumar et al., 2021).

We next assessed the requirement of the hFip1–CPSF30 interactions for RNA 3′ polyadenylation using an in vitro polyadenylation assay. Incubation of a model RNA substrate with purified wild-type mPSF (Figure 2—figure supplement 1A) and PAP resulted in processive addition of ~60 adenosine nucleotides, which was dependent on the presence of ATP and the AAUAAA hexameric PAS in the RNA (Figure 2A). The efficiency of 3′ polyadenylation was reduced upon incubation of the substrate with mPSF complexes containing CPSF30 ZF4 or ZF5 mutants capable of binding only one copy of hFip1 (Figure 2A). No polyadenylation was observed upon incubation with mPSF containing the CPSF30 ZF4/ZF5 double mutant (Figure 2A), consistent with the loss of hFip1 and PAP recruitment (Figure 1G), and polyadenylation could not be rescued by the addition of recombinant hFip1 in trans (Figure 2A). Collectively, these observations indicate that both hFip1-binding sites in CPSF30 contribute to the efficiency of RNA 3′ polyadenylation, suggesting that the presence of two hFip1 copies, and thus two PAP recruitment sites, in mPSF is required for highly efficient, processive 3′ polyadenylation. However, neither hFip1-binding site is strictly necessary for RNA 3′ polyadenylation, suggesting their functional redundancy.

Figure 2 with 1 supplement see all
hFip1 directly recruits poly(A) polymerase.

(A) Polyadenylation activity assay of mPSF complexes containing wild-type and mutant CPSF30 proteins as well as hFip1 added in trans (rightmost lane) using a Cy5-labeled PAS-containing RNA substrate. An RNA substrate lacking the canonical AAUAAA PAS hexanucleotide is denoted by its substitute sequence, AGUACA. Polyadenylated RNA products are indicated as RNA-(A)n. (B) Pull-down analysis of immobilized StrepII-tagged mPSF complexes comprising N-terminal truncations of hFip1 with GFP-PAP. GFP-PAP is visualized by in-gel GFP fluorescence (bottom). Asterisk denotes contaminating protein. (C) Polyadenylation activity assay of mPSF complexes containing hFip1 truncations. (D) Size-exclusion chromatography coupled to multiangle static light scattering (SEC-MALS) analysis of reconstituted mPSF:PAP:RNA complexes and in the absence (purple) or presence of excess PAP (yellow). Theoretical molecular masses of 1:1 and 1:2 mPSF:PAP complexes are indicated.

PAP recruitment occurs via hFip1 N-terminal region

In S. cerevisiae, a poorly conserved peptide motif in the N-terminal region of Fip1 directly interacts with the poly(A) polymerase Pap1 (Meinke et al., 2008). Similarly, the N-terminal region of human hFip1, upstream of the CD, is required for PAP interaction (Kaufmann et al., 2004) but the precise PAP interaction site in human hFip1 has not been identified. To this end, we tested the interaction of green fluorescent protein (GFP)-tagged PAP with purified mPSF complexes containing truncated hFip1 fragments in an in vitro pull-down experiment. PAP was detectably, albeit weakly, coprecipitated by mPSF containing a hFip1 fragment spanning both the N-terminal and CD regions (residues 1–195) as well as by mPSF containing an N-terminally truncated hFip1 (residues 36–195) (Figure 2B). However, further N-terminal truncation of hFip1 resulted in the loss of PAP binding, indicating that a region spanning residues 36–80 in human hFip1 is required for PAP interaction (Figure 2B). An additional pull-down experiment using recombinant PAP and glutathione-S-transferase (GST)-fused hFip1 fragments revealed that although the hFip1 region comprising residues 36–80 was required for PAP interaction, it was not sufficient (Figure 2—figure supplement 1B). This suggests that additional parts of hFip1 contribute to PAP binding.

We subsequently tested the activity of mPSF complexes containing N- or C-terminally truncated hFip1 in the polyadenylation assay. In agreement with the interaction data, mPSF complexes containing hFip1 fragments spanning residues 1–195 or 36–190 were able to support efficient RNA 3′ polyadenylation (Figure 2C), whereas mPSF complexes containing hFip1 fragments comprising residues 80–195 or 130–195 were not. Together, these results indicate that hFip1 residues 36–80 are required for the recruitment of PAP to effect mPSF-dependent 3′ polyadenylation. Interestingly, we also observed that polyadenylation levels were reduced with mPSF containing full-length hFip1 (residues 1–378, isoform 4), as compared to mPSF containing C-terminally truncated hFip1 (residues 1–195), suggesting that the C-terminal region of hFip1, which is proline-rich and predicted to be intrinsically disordered, negatively modulates the processivity of mPSF-dependent 3′ polyadenylation.

CPSF recruits only one copy of PAP

Prior studies have indicated that the polymerase module of endogenous yeast CPF comprises up to two copies of Pap1 (Casañal et al., 2017). Furthermore, a complex comprising human CPSF30 ZF4 and ZF5 domains and two hFip1 molecules is capable of simultaneous interaction with two PAP molecules in vitro (Hamilton and Tong, 2020). To determine whether this also occurs in the context of human mPSF, we analyzed the mPSF–PAP interaction by SEC-MALS. Despite only weakly interacting in pull-down analysis, at high PAP concentrations (40 µM), mPSF and PAP formed a stable complex that could be purified by SEC. Analysis of this complex using SEC-MALS revealed an apparent molecular mass of 347 kDa, closely matching the predicted molecular mass of a complex containing two hFip1 molecules and one PAP (337 kDa) (Figure 2D). Addition of excess PAP to the prepurified mPSF–PAP sample did not cause peak broadening; furthermore, no change in the detected apparent molecular mass could be observed. These results suggest that mPSF predominantly associates with only one PAP molecule at a time, despite the presence of two copies of hFip1 in the complex.

The N-terminal region of hFip1 interacts with CstF77

In analogy with the yeast polyadenylation machinery, human Fip1 has previously been shown to interact with CstF77 (Preker et al., 1995; Kaufmann et al., 2004). To validate these observations and identify the interaction determinants in hFip1, we performed a pull-down experiment with GST-tagged hFip1 fragments and maltose-binding protein (MBP)-tagged fragment of CstF77 comprising the HAT domain (residues 21–549). The very N-terminal region of hFip1 spanning residues 1–35 was necessary and sufficient for the interaction with the CstF77 HAT domain (Figure 3A). Notably, this region is dispensable for the interaction of hFip1 with PAP and for RNA 3′ polyadenylation (Figure 2B, C).

Figure 3 with 3 supplements see all
hFip1 interacts with CstF77 through a conserved motif within its N-terminal acidic domain.

(A) Pull-down analysis of immobilized GST-hFip1 fragments with MBP-CstF7721–549. (B) Cartoon representation of the crystal structure of the CstF77241–549–hFip11–35 complex, superimposed onto the structure of murine CstF77 (white, PDB ID: 2OOE). (C) Zoomed-in view of the hFip1–CstF interaction interface. (D) Multiple sequence alignment of the N-terminal region of Fip1 orthologs. (E) Pull-down analysis of immobilized wild-type and mutant GST-hFip11–35 proteins with MBP-CstF7721–549 and MBP-CstF77mut (R395A/R402A/K431A). Asterisk indicates contaminating free GST protein. (F) 3D cryo-EM density map (EMD-20861) of the human CPSF160–WDR33–CPSF30–PAS RNA–CstF77 complex (Zhang et al., 2019), displayed at contour level 0.015 and color coded according to the corresponding atomic protein model (PDB ID 6URO). The hFip1–CstF77 crystal structure from this study was superimposed onto the atomic model of CstF77, and atomic model of hFip1 is shown (cyan). Inset shows a zoomed-in view of unassigned density that matches hFip1.

To shed light on the hFip1–CstF77 interaction, we subsequently reconstituted a complex comprising the hFip11–35 fragment with a truncated construct of the CstF77 HAT domain (residues 241–549) and determined its X-ray crystallographic structure at a resolution of 2.7 Å. The structure reveals that hFip1 binds to a conserved positively charged patch located on the convex surface of the CstF77 HAT domain arch (Figure 3B, Figure 3—figure supplement 1A, B). Within the hFip11–35 fragment, only the evolutionarily conserved residues 20–27 were ordered, adopting an alpha-helical conformation (Figure 3C, D). Interaction of hFip11–35 with CstF77 involves salt-bridge contacts of Glu22hFip1 and Glu23hFip1 with Arg402CstF77, and hydrophobic contacts involving Leu26hFip1 and Tyr27hFip1 with Phe398CstF77, Val428CstF77, Ile432CstF77, and Leu435CstF77. Additionally, the Tyr27hFip1 side chain interacts with Arg395CstF77 via π–π stacking. Corroborating these structural observations, simultaneous substitutions of Glu22hFip1 and Glu23hFip1, or Trp25hFip1, Leu26hFip1 and Tyr27hFip1 with alanine disrupted the hFip11–35–CstF7721–549 interaction in a pull-down experiment, whereas alanine substitution of Trp25hFip1 alone did not have an effect (Figure 3E). In turn, mutation of the positively charged interaction surface in CstF77 (Arg395, Arg402, and Lys431 mutated to alanines) abolished the interaction with hFip11–35 (Figure 3E).

A previously determined cryo-EM reconstruction of the human mPSF–CstF77 complex revealed that the interaction of the CstF77 HAT domain dimer with mPSF is primarily mediated by extensive contacts with WDR33 and CPSF160 (Zhang et al., 2019). Upon close inspection, the cryo-EM map from this dataset (EMDB entry EMD-20861) exhibits residual densities on both CstF77 protomers that could be attributed to the binding of two hFip1 molecules via their N-terminal regions (Figure 3H). This observation indicates that CstF77 is capable of binding two hFip1 copies when bound to mPSF. We subsequently tested the contribution of hFip11–35 to the mPSF–CstF77 interaction in a pull-down experiment using MBP-tagged CstF77 and mPSF complexes containing truncated hFip1 fragments. Although all mPSF complexes were capable of binding CstF77, reduced levels of CstF77 coprecipitation were observed with mPSF containing N-terminally truncated hFip1 that lacked the CstF77 interacting region (Figure 3—figure supplement 2A). Furthermore, to exclude that the hFip1-binding site is obstructed upon assembly of CstF77 into a holo-CstF complex, we tested interaction of GST-tagged hFip11–35 in a pull-down experiment using with purified holo-CstF comprising full-length CstF77, CstF50, and CstF641–198. Comparable amounts of holo-CstF and MBP-CstF77 were precipitated by GST-hFip11–35 indicating that the Fip1 interaction interface in CstF77 remains exposed upon CstF complex assembly (Figure 3—figure supplement 3A). Taken together, these results suggest that direct interactions between hFip1 and CstF77 contribute to the assembly of the CPSF–CstF complex during mRNA 3′ end biogenesis.

CstF inhibits polyadenylation

As CstF77 and PAP bind to nonoverlapping, yet adjacent, sites in hFip1, CstF77 binding could nevertheless preclude PAP recruitment due to steric crowding. To probe this, we carried out a pull-down experiment with GST-tagged hFip1 and mixtures of MBP-tagged CstF77 and GFP-tagged PAP at varying molar ratios. In the presence of excess CstF77, PAP binding was considerably reduced, indicating that CstF77 competes with PAP for binding to hFip1 (Figure 4A). Consistent with this, CPSF-dependent RNA 3′ polyadenylation was substantially reduced in the presence CstF77, suggesting that CstF77 inhibits 3′ polyadenylation via interaction with hFip1 (Figure 4B). The inhibitory effect of CstF77 was reduced either when mPSF lacked the N-terminal CstF77 interaction site in hFip1 (Figure 4B) or when CstF77 (CstFmut) was incapable of interaction with the N-terminal region of hFip1 (Figure 4—figure supplement 1A), indicating that the inhibitory effect of CstF77 is in part dependent on its interaction with hFip1. In both cases, addition of excess CstF77 led to a reduction of RNA 3′ polyadenylation rate, although not to the same extent (Figure 4B, Figure 4—figure supplement 1A). Furthermore, inhibition of 3′ polyadenylation was also observed upon addition of a holo-CstF complex, suggesting that the inhibitory effect of CstF77 is maintained when it is assembled within CStF (Figure 4C). To exclude the possibility that the observed CstF77-dependent reduction of RNA 3′ polyadenylation rate might be due to the close proximity of the PAS and the 3′ terminal CA dinucleotide in the SV40 mRNA-based substrate (11 nt), we tested an alternative RNA substrate (adenoviral L3 PAS-containing mRNA) in which the PAS and the cleavage site are separated by 20 nt and observed a similar degree of inhibition (Figure 4—figure supplement 1B), suggesting that the inhibition is likely not a consequence of steric crowding due to the proximity of the PAS and the 3′ end. Altogether, these results suggest that CstF inhibits RNA 3′ polyadenylation via CstF77, and that CstF77-mediated inhibition is in part dependent on its interaction with hFip1.

Figure 4 with 2 supplements see all
CstF77 competitively inhibits 3′ polyadenylation.

(A) Pull-down analysis of immobilized GST-hFip11–195 with varying molar ratios of GFP-PAP and MBP-CstF7721–549. GFP-PAP is visualized by in-gel GFP fluorescence (bottom). Asterisk denotes contaminating protein. (B) Polyadenylation activity assay of mPSF complexes containing full-length hFip1 and N-terminally truncated hFip1 (hFip136–195) in the presence of varying molar ratios of CstF77. Polyadenylated RNA products are indicated as RNA-(A)n. (C) Polyadenylation activity assay of mPSF in the presence of varying molar ratios of holo-CstF complex.

Discussion

Despite extensive efforts to obtain structural insights into the molecular organization and regulation of the eukaryotic mRNA 3′ end processing machinery, high-resolution structural information has so far only been obtained for stable subassemblies composed of structurally rigid subunits (Casañal et al., 2017; Clerici et al., 2017; Clerici et al., 2018; Sun et al., 2018; Hill et al., 2019; Zhang et al., 2019; Hamilton and Tong, 2020; Kumar et al., 2021). Although hFip1 is an integral component of the CSPF complex, specifically its mPSF module, it has not been structurally visualized in this context owing to its intrinsically disordered nature (Meinke et al., 2008).

In our study, we reveal the molecular basis for the interactions of human hFip1 with both CPSF30, PAP, and CstF77. While confirming the 2:1 binding stoichiometry of the hFip1:CPSF30 interaction in isolation (Hamilton and Tong, 2020; Kumar et al., 2021), we expand this finding to the CPSF complex, confirming that its mPSF module assembles with two hFip1 copies in cells and demonstrating that both the ZF4 and ZF5 domains in CPSF30 are capable of binding hFip1 independently. Using polyadenylation assays we show that the two hFip1 copies are functionally redundant in recruiting PAP to the mPSF, which increases the processivity of RNA 3′ polyadenylation. As the recruitment of PAP to the 3′ end of the cleaved pre-mRNA is prerequisite for its processivity (Ezeokonkwo et al., 2011), while PAP only weakly associates with the mPSF, the presence of two hFip1 copies thus likely increases the 3′ polyadenylation efficiency polyadenylation by increasing the local PAP concentration. While recent studies of human CPSF30 and its yeast homolog Yth1 reported higher binding affinity for the Fip1:ZF4 interaction as compared to Fip1:ZF5 (Hamilton and Tong, 2020; Kumar et al., 2021), we show that polyadenylation efficiency is reduced equally independent of which hFip1 interaction site (ZF4 or ZF5) is impaired. This indicates that PAP recruitment by mPSF is the limiting factor in 3′ polyadenylation.

Although the yeast Fip1–Pap1 interaction has been extensively characterized, the Pap1 interaction motif in Fip1, as observed in the crystal structure of the complex (Meinke et al., 2008), is poorly conserved in human Fip1 (Figure 4—figure supplement 2A), only partially mapping to hFip1 residues 80–86 (hFip1 isoform 4). Recent analysis of the yeast Fip1–Pap1 interaction using nuclear magnetic resonance, however, showed that additional residues located N-terminally of the Pap1-binding motif are involved in Pap1 interaction (Kumar et al., 2021). This aspartate-rich acidic region is well conserved in hFip1, corresponding to residues 58–79 (hFip1 isoform 4). In agreement with this, we show that an additional N-terminal segment in hFip1 spanning residues 36–80 and including the aspartate-rich acidic motif is required but not sufficient for PAP binding. This suggests that the hFip1–PAP interaction mode closely resembles that of yeast Fip1–Pap1, despite low sequence conservation of the respective interaction motifs in yeast Fip1 and hFip1.

Notably, our biophysical analysis of the human mPSF–PAP interaction reveals that despite the presence of two hFip1 copies, only one copy of PAP is stably recruited by mPSF. This finding is unexpected and in contrast to the previous observation that two copies of the PAP catalytic domain are stably bound by the CPSF30–hFip1 subcomplex in isolation (Hamilton and Tong, 2020). Moreover, up to two Pap1 copies have been detected in the polymerase module of yeast CPF by native mass spectrometry (Casañal et al., 2017). It is important to note that although our experimental data indicate that only one PAP associates with mPSF, this does not exclude the possibility that mPSF binds a second copy of PAP with a low affinity. Nevertheless, we speculate that the observed 1:1 mPSF–PAP complex likely reflects the predominant state under physiological conditions. As the efficiency of the 3′ polyadenylation reaction depends on PAP recruitment, the presence of two PAP-binding sites in mPSF thus might not serve to simultaneously populate both sites with one PAP molecule each but to increase the probability of PAP recruitment to enhance polyadenylation efficiency. Furthermore, the presence of two hFip1 copies might instead be required for mPSF integrity and its interactions with CstF. In this context, it is conceivable that the binding of two PAP molecules to mPSF bound to a substrate RNA is precluded due to molecular crowding or steric hindrance, particularly considering that the two Fip1 molecules make nonidentical interactions with mPSF, and the two Fip1-binding sites in the CstF77 homodimer are not equivalent when CstF is bound to mPSF.

The interaction between CPSF and CstF has previously been shown to involve direct contacts between the CstF77 homodimer and an extensive interface provided by the CPSF160 and WDR33 subunits of CPSF (Zhang et al., 2019), yet CstF also interacts with CPSF via hFip1 (Kaufmann et al., 2004). Our crystal structure of the hFip1–CstF77 subcomplex reveals that a hFip1 binds via conserved motif within the N-terminal ‘acidic’ region to the convex arch of the CstF77 HAT domain on both protomers in the CstF77 homodimer, resulting in a 2:2 stoichiometry. By reanalysis of previously reported cryo-EM data (Zhang et al., 2019), we reveal that this interaction mode is preserved in the context of the mPSF–CstF complex. Strikingly, both CstF77 and holo-CstF inhibit 3′ polyadenylation in vitro. Moreover, CstF77-mediated inhibition is partially dependent of its interaction with hFip1, which likely reflects the dual interaction mode of CStF with mPSF. Accordingly, the hFip1–PAP and hFip1–CstF77 interactions appear to be competitive, possibly as a result of the proximity of the PAP and CstF77 interaction sites within the hFip1 N-terminus. As CStF is strictly required for CPSF73-dependent pre-mRNA cleavage while PAP might not be (Boreikaite et al., 2022), these results imply that the CPSF–CstF interaction is disrupted or undergoes a remodeling after cleavage to enable PAP recruitment to the cleaved pre-mRNA and subsequent 3′ polyadenylation.

To perform the coupled reaction steps of cleavage and polyadenylation, the 3′ end processing machinery likely undergoes a sequence of conformational and compositional rearrangements as polyadenylation site recognition by the mPSF module of CPSF and activation by RBBP6 triggers CstF-dependent cleavage by the mCF, after which the nascent 3′ end needs to be made accessible to PAP for subsequent poly(A) synthesis (Boreikaite et al., 2022; Schmidt et al., 2022). Based on our structural and biochemical findings, we propose a model in which hFip1 helps coordinate the two steps of 3′ end processing. Initially, the two hFip1 molecules present in mPSF facilitate the assembly of CPSF and CstF on the pre-mRNA via the interactions of their N-terminal motifs with CstF77 (Figure 5A). In part, these interactions also preclude PAP recruitment until the pre-mRNA has been cleaved and a free 3′ end has been generated. Upon endonucleolytic cleavage of the pre-mRNA by CPSF73, remodeling of the 3′ end processing machinery, possibly enabled by the dissociation of the downstream pre-RNA cleavage product and concomitant release of CstF, reduces steric constraints around the nascent 3′ end and exposes PAP interaction sites in hFip1, enabling PAP recruitment to initiate processive 3′ polyadenylation of the cleaved pre-mRNA (Figure 5B). The conformational and compositional transitions required for accessing the nascent 3′ end are orchestrated by hFip1 and facilitated by its flexible attachment to mPSF via CPSF30, as well as by its intrinsic conformational dynamics (Meinke et al., 2008; Ezeokonkwo et al., 2011; Kumar et al., 2021). The presence of two hFip1 molecules in the 3′ end processing complex promotes efficient PAP recruitment and contributes to the processivity of 3′ end polyadenylation. Our model implies that PAP might be recruited to the pre-mRNA only after the cleavage step, which is supported by recent findings reported by Boreikaite et al., demonstrating that the presence of PAP is not required for endonucleolytic cleavage by mCF, but is contradicted by the study of Schmidt et al., which reported that PAP is required for pre-mRNA cleavage. The functional role of PAP in pre-mRNA cleavage thus remains unclear, necessitating further studies. Finally, the functional role of hFip1 as a major interaction platform for 3′ end processing factors is also important in the context of the its well-documented role in regulating alternative polyadenylation (Lackford et al., 2014) as it suggests that the interactions with processing factors might be further modulated by direct interactions of Fip1 with U-rich sequences in the pre-mRNA.

Model of CPSF-mediated pre-mRNA cleavage and polyadenylation and CstF77-dependent inhibition of polyadenylation.

(A) Prior to pre-mRNA cleavage, PAP recruitment is inhibited by CStF, in part due to competitive interactions of CstF77 and hFip1 (left). (B) Upon pre-mRNA cleavage, structural remodeling of the CPSF–CstF complex enables hFip1 to recruit PAP to the nascent 3′ end of the mRNA and consequently stimulates polyadenylation. Figure 1—figure supplement 1: sequence alignment of CPSF30 zinc finger domains. (A) Sequence alignment of human CPSF30 zinc finger domains. Residues responsible for RNA interactions (in ZF2/ZF3) or hFip1 interaction (in ZF4/ZF5) are highlighted and the nature of their interaction color coded. ZF4/ZF5 domains contain proline residues (yellow) at positions corresponding to critical main chain hydrogen bonding interactions in ZF2/ZF3.

In sum, these results advance our understanding of hFip1 as a multivalent interaction hub in mRNA 3′ end processing and unravel a novel aspect of polyadenylation regulation by CstF. Through interspacing binding sites for processing factors with intrinsically disordered, low-complexity sequences hFip1 can achieve the required degree of conformational freedom to accommodate the remodeling of the 3′ end processing machinery and ensure correct spatiotemporal regulation of the processing factors at the nascent mRNA 3′ end. The molecular basis of these transitions, however, awaits further structural and functional investigations.

Materials and methods

Protein expression and purification

Protein expression vectors

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To generate polypromoter plasmids for E. coli expression using ligation-independent cloning (LIC) analogous to MacroLab Series 438, MacroLab Series 2 vectors were modified by inserting whole expression cassettes as gBlocks (IDT) into 2B-T with Gibson assembly carrying the necessary modifications. The resulting vectors denoted 16-B (for His6-TEV-tagged protein expression), 16 M (for His6-MBP-TEV-tagged protein expression), and 16M_ΔHis (for MBP-TEV-tagged protein expression) have two PmeI restriction sites flanking the T7 expression cassette, an internal SspI site for target gene insertion, and a SwaI site downstream of the T7 terminator for biobricks-type assembly using LIC. Gene assembly proceeds following the Series 438 vectors assembly protocol (Gradia et al., 2017).

Cloning for expression in E. coli

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Constructs encoding for CPSF30 isoform 3 (Uniprot O95639-3), hFip1 isoform 4 (Uniprot Q6UN15-4), poly(A) polymerase alpha (Uniprot P51003-1), and CstF77 (Uniprot Q12996-1) were cloned into LIC expression vectors 1B (gift from Scott Gradia, Addgene plasmid #29653), 1 M (Addgene plasmid #29656), 2 G-T (Addgene plasmid #29707), 2GFP-T (Addgene plasmid #29716), and cotransformation vector 13 S-A (Addgene plasmid 48323), respectively. DNA encoding for hFip1130–195 was first cloned into 2 G-T, PCR amplified starting from the GST-tag and inserted into 13 S-A using LIC cloning. Point mutations in CPSF30, hFip1, PAP, and CstF77 were introduced by obtaining linear DNA fragments (GeneArt Strings, Thermo Fisher) encoding for the desired construct with LIC overhangs and cloned into the respective expression vectors according to Supplementary file 1.

Cloning for expression in Sf9 cells

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DNA encoding human CPSF160 (Uniprot Q10570), WDR33 (Uniprot Q9C0J8-1), CPSF30 isoform 3, and hFip1 isoform 4 were cloned into MacroBac Series 438 cloning system vectors (Gradia et al., 2017) according to Supplementary file 1. Subcloning of three- or four-subunit mPSF complexes into a single baculovirus transfer plasmid was performed following the MacroBac protocol (Gradia et al., 2017). For FLAG-tagged mPSF complexes, subcloning was performed using the biGBac protocol (Weissmann et al., 2016).

Expression and purification of CPSF30 and hFip1 for SEC-MALS

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His6-MBP-TEV-CPSF301–243 wt and mutants (Y127E, F155E, and Y127E/F155E) were expressed overnight in E. coli BL21 star (DE3) cells and His6-GFP-TEV-hFip11–195 in E. coli Rosetta2 (DE3) cells, respectively, at 18°C by addition of isopropyl thio-β-galactoside (IPTG) to a final concentration of 0.5 mM at OD600 of about 0.6–0.8. Cells were resuspended in buffer A (25 mM Tris–HCl pH 7.5, 200 mM NaCl) supplemented with 0.5 mM Tris (2-carboxyethyl)phosphine (TCEP), 1 µM Pepstatin, and 400 µM 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride (AEBSF) protease inhibitor followed by lysis via sonication. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and clarified lysate was purified on Ni-NTA agarose resin (QIAGEN) eluted with buffer A supplemented with 0.5 mM TCEP and 200 mM imidazole. The protein was further purified by size-exclusion chromatography on a Superdex 75 (Cytiva) column, eluting with buffer A supplemented with 1 mM dithiothreitol (DTT). Eluting peak fractions were concentrated in centrifugal filter (Amicon Ultra-15, MWCO 30 kDa, Merck Millipore), flash frozen, and stored at −80°C.

MBP-CPSF30 proteins for pull-down analysis

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His6-MBP-TEV-CPSF301-243 mutants were expressed and purified as described for SEC-MALS. For CPSF30 point mutants Y151E, Y127E/Y151E, Y127E/Y155E, a high salt wash (25 mM Tris–HCl pH 7.5, 1 M NaCl) during Ni-IMAC purification was included prior to elution with buffer A buffer supplemented with 200 mM imidazole.

PAP expression and purification

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His6-MBP-TEV-PAP1–504 was expressed in E. coli Rosetta2 (DE3) cells overnight at 18°C by induction with 0.5 mM IPTG at OD600 of about 0.6–0.8. Cells were lysed by sonication in 20 mM Tris–HCl pH 8.0, 500 mM NaCl, 5 mM imidazole, 0.5 mM TCEP supplemented with 0.1 µM Pepstatin, and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and cleared lysate was subjected to Ni-NTA resin (QIAGEN), washed and protein eluted with buffer B (20 mM Tris–HCl pH 8.0, 200 mM NaCl) supplemented with 200 mM imidazole. Eluted protein was further purified on MBPTrap HP (Cytiva), eluting in buffer B supplemented with 10 mM maltose. The eluted protein fractions were injected onto a Superdex 200 column (Cytiva) equilibrated in buffer B supplemented with 1 mM DTT. Tag was cleaved off the protein with His6-MBP-TEV protease, and the cleavged tags including protease removed from protein sample using a MBPTrap HP (Cytiva). For use in pull-down analysis, the tag was not cleaved from His6-MBP-TEV-PAP1–504 wt and mutant (R395A, R402A, and K431A) after size-exclusion chromatography. Purified protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 50 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C.

Expression and purification of GFP-PAP for pull-down analysis

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His6-GFP-TEV-PAP1–504 was expressed overnight in E. coli Rosetta2 (DE3) cells at 18°C by induction with 0.5 mM IPTG at OD600 of about 0.6–0.8. Cells were lysed by high-pressure cell disruption at 25 kpsi in buffer B supplemented with 5 mM imidazole, 0.5 mM TCEP, 0.1 µM Pepstatin, and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and clarified lysate was subjected to Ni-NTA resin (QIAGEN), washed and protein eluted with buffer B supplemented with 250 mM imidazole. Eluted protein was further purified on Superdex 200 column (Cytiva) equilibrated in 20 mM Tris–HCl pH 8.0, 150 mM NaCl, and 0.5 mM TCEP. Purified protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 50 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C.

Expression of mPSF complexes

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For expression of mPSF complexes in Sf9 cells (Thermo Fisher Scientific, cat. no. 11496015; cell line was authenticated and tested for mycoplasma contamination by the manufacturer, no further validation was done by the authors), recombinant baculoviruses were generated according to the Bac-to-Bac Baculovirus expression system (Invitrogen). 2 ml of P3 virus were used to infect 1 l of Sf9 insect cells at a density of 1.1 × 106 ml−1. Cells were harvested 72 hr postinfection.

Purification of mPSF complexes for polyadenylation assays and pull-down analysis

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Cells were resuspended in buffer C (25 mM Tris–HCl pH 7.5, 200 mM NaCl, 10% glycerol, and 0.5 mM TCEP) supplemented with 5 mM imidazole, 0.05% Tween-20, and cOmplete Protease-Inhibitor-Cocktail (Roche). Cells were lysed by sonication, cleared by centrifugation (20 min, 20,000 × g, 4°C), and the clarified lysate was purified on Ni-NTA resin (QIAGEN) eluting in buffer C supplemented with 200 mM imidazole. The eluted protein was incubated with Strep-Tactin Sepharose (IBA Lifesciences) beads, washed with 10 column volumes of buffer B, and eluted with buffer C supplemented with 5 mM Desthiobiotin. Strep-Tactin purified mPSF complexes were concentrated in centrifugal filter (Amicon Ultra-15, MWCO 300 kDa, Merck Millipore) to approximately 0.5 mg ml−1. To account for impurities, mPSF complex concentrations were assessed on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and adjusted accordingly (Figure S1C), aliquoted, flash frozen, and stored at −80°C. For use in pull-down analysis, CPSF complexes were used directly after Strep-Tactin purification.

Purification of mPSF–PAP complex for SEC-MALS analysis

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mPSF complexes comprising CPSF160–WDR331–410–CPSF30–hFip11–198 for SEC-MALS analysis were produced as described above with subsequent tag removal by incubation with His6-TEV protease. mPSF (assuming to comprise two hFip1) was supplemented with untagged PAP1–504 in 2.5-fold molar excess and 1.2-fold molar excess of a 27-nt RNA substrate based on the SV40 pre-mRNA containing a PAS and a 3′ penta-A tail (CUGCAAUAAACAACUUAACAACAAAAA). The complex was purified on a Superose 6 column (Cytiva) in 20 mM HEPES pH 8.0, 150 mM KCl, 0.5 mM TCEP. The mPSF–PAP complex was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 100 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C.

Expression and purification of GST-hFip1 proteins for pull-down analysis

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His6-GST-TEV-hFip11–195 and His6-GST-TEV-hFip11–35 were expressed overnight in E. coli BL21 star (DE3) cells at 18°C by induction with 0.5 mM IPTG at OD600 of about 0.6–0.8. Cells were lysed by sonication in buffer A supplemented with 5 mM imidazole, 1 µM Peptsatin A, and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and clarified lysate was purified on Ni-NTA resin (QIAGEN) eluting in buffer A supplemented with 200 mM imidazole in gravity flow. Eluted protein was loaded on a HiTrap Q FF (Cytiva) anion exchange chromatography column and eluted with a linear gradient from 200 mM to 1 M NaCl over 15 CV in 25 mM Tris–HCl pH 7.5 and 1 mM DTT. Eluting peak fractions were further purified on a Superdex 75 (Cytiva) column equilibrated in 25 mM Tris–HCl pH 7.5, 500 mM NaCl, 1 mM DTT. Protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 30 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C. His6-GST-TEV-hFip136–195 was expressed in E. coli BL21 star (DE3) cells overnight at 18°C by induction with 0.5 mM IPTG at OD600 of about 0.6–0.8. Cells were lysed by sonication in buffer A supplemented with 1 mM DTT, 1 µM Peptsatin A, and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C). Clarified lysate was subjected to a GSTrap Fast Flow (Cytiva) column and eluted in buffer A supplemented with 1 mM DTT and 10 mM GSH. His6-GST-TEV-hFip11–35 mutants (E22A, E23A; W25A, L26A, Y27A; W25A) were expressed and purified analogously to His6-GST-TEV-hFip136–195 but with buffers containing 500 mM NaCl. All proteins were further purified on a Superdex 200 (Cytiva) column equilibrated in buffer A supplemented with 1 mM DTT. Protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 30 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C. His6-GST-TEV-hFip136–80 was expressed in E. coli BL21 star (DE3) cells overnight at 18°C by induction with 0.5 mM IPTG at OD600 of about 0.6–0.8. Cells were lysed by sonication in buffer A supplemented with 1 mM DTT, 1 µM Peptsatin A, and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C). Clarified lysate was subjected to a GSTrap Fast Flow (Cytiva) column, washed with buffer A supplemented with 1 mM DTT prior to elution in buffer A supplemented with 1 mM DTT and 10 mM GSH. Protein was further purified on a Superdex 200 (Cytiva) column equilibrated in buffer A supplemented with 1 mM DTT. Protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 30 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C. His6-GST-TEV-hFip180–195 was expressed and purified using the same protocol, but changing to buffer B. His6-GFP-TEV-hFip11–195 mutants (W150E, F161E, and W170E) were expressed and purified following the same purification strategy as for the His6-GFP-TEV-hFip11–195 wt protein.

Expression and purification of His6-CstF77 proteins for polyadenylation assays

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His6-TEV-CstF7721–549 wt and mutant (R395A, R402A, K431A) were expressed overnight in E. coli BL21 star (DE3) cells at 18°C by induction with 0.5 mM IPTG at OD600 of about 0.6–0.8. Cells were lysed by sonication in 40 mM Tris–HCl pH 7.5, 500 mM NaCl, 5 mM imidazole, supplemented with 1 µM Peptsatin A and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and clarified lysate was purified on Ni-NTA resin (QIAGEN) eluting with buffer A supplemented with 250 mM imidazole in gravity flow. Salt concentration and pH of protein sample were reduced to 60 mM NaCl and pH 7.0 by dilution and purified on a HiTrap SP FF (Cytiva) cation exchange chromatography column. Protein was eluted from column with a linear gradient from 60 mM to 1 M NaCl over 10 CV in 20 mM Tris–HCl pH 7.0 and 1 mM DTT. Eluting peak fractions were further purified on a Superdex 200 (Cytiva) column equilibrated in 20 mM Tris–HCl pH 7.5, 200 mM NaCl, 1 mM DTT. Protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 100 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C.

Expression and purification of MBP-CstF77 proteins for pull-down analysis

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For use in pull-down analysis, His6-MBP-TEV-CstF7721–549 wt and mutant (R395A, R402A, and K431A) were expressed and purified analogous to CstF77 for cocrystallization with hFip1, omitting tag cleavage with His6-TEV protease prior to size-exclusion chromatography.

Expression and purification of CstF for polyadenylation assay

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For expression of CstF complex comprising CstF77, His6-TEV-2xStrepII-TEV-2xStrepII-TEV-CstF641–198, and CstF50 in Sf9 cells (Thermo Fisher Scientific, cat. no. 11496015; cell line was authenticated and tested for mycoplasma contamination by the manufacturer, no further validation was done by the authors), recombinant baculoviruses were generated according to the Bac-to-Bac Baculovirus expression system (Invitrogen). 2 ml of each P3 virus were used to coinfect 1 l of Sf9 insect cells at a density of 1.1 × 106 ml−1. Cells were harvested 72 hr postinfection. Cells were resuspended in buffer B supplemented with 10% glycerol, 5 mM imidazole, 0.4% Triton X-100, and cOmplete Protease-Inhibitor-Cocktail (Roche). Cells were lysed by sonication, cleared by centrifugation (20 min, 20,000 × g, 4°C) and the clarified lysate was purified on Ni-NTA resin (QIAGEN) eluting in buffer B supplemented with 10% glycerol, 150 mM imidazole, 0.4% Triton X-100. The eluted protein was incubated with Strep-Tactin Sepharose (IBA Lifesciences) beads, washed with 10 column volumes of 25 mM Tris pH 7.5, 120 mM KCl, 10% glycerol, 2 mM MgCl2, 0.5 mM TCEP, 0.05% Tween-20, and eluted with wash buffer supplemented with 5 mM Desthiobiotin. The protein was further purified by size-exclusion chromatography on a Superdex 200 (Cytiva) column eluting with 25 mM Tris pH 7.5, 120 mM KCl, 10% glycerol, 2 mM MgCl2, and 0.5 mM TCEP. Protein was diluted with 2× dilution buffer (25 mM Tris pH 7.5, 10% glycerol, 2 mM MgCl2, 0.5 mM TCEP, 10% Tween-20) to reduce KCl concentration to 50 mM, concentrated in centrifugal filter (Amicon Ultra-15, MWCO 100 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C.

Expression and purification of CstF complexes for pull-down analysis

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CstF complexes comprising CstF77, His6-TEV-2xStrepII-TEV-2xStrepII-TEV-CstF641–198, and CstF50 were expressed as described above. Cells were resuspended in buffer B supplemented with 5 mM imidazole and cOmplete Protease-Inhibitor-Cocktail (Roche). Cells were lysed by sonication, cleared by centrifugation (20 min, 20,000 × g, 4°C) and the clarified lysate was purified on Ni-NTA resin (QIAGEN) eluting in buffer B supplemented with 150 mM imidazole. The eluted protein was incubated with Strep-Tactin Sepharose (IBA Lifesciences) beads, washed with 10 column volumes of buffer B and eluted with buffer B supplemented with 5 mM Desthiobiotin. After overnight cleavage with His6-TEV protease, the protein was further purified by size-exclusion chromatography on a Superdex 200 (Cytiva) column eluting with 20 mM Tris pH 7.5, 150 mM KCl, 1 mM DTT. Protein was concentrated in centrifugal filter (Amicon Ultra-15, MWCO 100 kDa, Merck Millipore), aliquoted, flash frozen, and stored at −80°C.

Preparation of CPSF30–hFip1 complex for crystallization

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Plasmids encoding for His6-TEV-CPSF30118–178 and GST-TEV-hFip1130–195 was cotransformed and proteins were expressed overnight in E. coli BL21 star (DE3) cells at 18°C by addition of IPTG to a final concentration of 0.5 mM at OD600 of about 0.6–0.8. Cells were resuspended in buffer A supplemented with 1 µM Pepstatin and 400 µM AEBSF, and lysed by sonication. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and protein was purified on Glutathione Sepharose 4 Fast Flow resin (Cytiva), eluting with buffer A supplemented with 10 mM reduced L-glutathione (GSH). After overnight cleavage with His6-TEV protease, the protein was further purified by size-exclusion chromatography on a Superdex 75 (Cytiva) column with a GSTrap Fast Flow (Cytiva) column in tandem to capture any residual GST tags, eluting with 25 mM Tris–HCl pH 7.5, 150 mM NaCl, and 1 mM DTT. Eluting peak fractions were concentrated in centrifugal filter (Amicon Ultra-15, MWCO 10 kDa, Merck Millipore) to 6.14 mg ml−1, flash frozen, and stored at −80°C.

Preparation of CstF77–hFip1 complex for crystallization

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His6-MBP-TEV-CstF77241–549 was expressed in E. coli BL21 star (DE3) cells at 18°C overnight by addition of IPTG to a final concentration of 0.5 mM at OD600 of about 0.6–0.8. Cells were resuspended in buffer containing buffer A supplemented with 1 mM DTT, 1 µM Pepstatin, and 400 µM AEBSF, and lysed by sonication. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and protein was purified on amylose resin (New England Biolabs) including a high salt wash with buffer containing 25 mM Tris–HCl pH 7.5, 500 mM NaCl, and 1 mM DTT prior to elution with buffer A supplemented with 1 mM DTT and 10 mM maltose. After digestion with His6-TEV protease, the tags and protease were removed from the protein by passage through a Ni-NTA superflow cartridge (QIAGEN). The protein was further purified by size-exclusion chromatography on a Superdex 200 Increase (Cytiva) column, eluting with 20 mM HEPES pH 7.5, 150 mM KCl, 1 mM TCEP. His6-GST-TEV-hFip11–35 was expressed in E. coli BL21-AI (Invitrogen) cells overnight at 18°C by induction with 0.2% arabinose at OD600 of 0.8. Cells were lysed by high-pressure cell disruption at 25 kpsi in buffer A supplemented with 1 mM DTT, 1 µM Pepstatin, and 400 µM AEBSF protease inhibitor. Lysate was cleared by centrifugation (20 min, 20,000 × g, 4°C) and clarified lysate was subjected to a GSTrap Fast Flow (Cytiva) column, washed with 25 mM Tris–HCl pH 7.5, 500 mM NaCl, and 1 mM DTT prior to elution in buffer A supplemented with 1 mM DTT and 10 mM GSH. Affinity tag was cleaved from protein using His6-MBP-TEV protease while dialyzing into buffer A supplemented with 1 mM DTT and hFip11–35 was further purified by size-exclusion chromatography on a Superdex 75 (Cytiva) column into 25 mM HEPES pH 7.5, 150 mM KCl, 1 mM DTT. The absolute mass of hFip11–35 (4.1 kDa) was confirmed with ESI-MS analysis. Peak fractions of both CstF77 and hFip1 were pooled individually, concentrated, flash frozen, and stored at −80°C.

CPSF30–hFip1 complex crystallization and structure determination

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The CPSF30–hFip1 complex was crystallized at 20°C using the hanging drop vapour diffusion method by mixing 0.5 μl of protein at 6.14 mg ml−1 with 0.5 μl of reservoir solution containing either 1.8 M (NH4)SO4, 0.1 M Bis-Tris pH 6.5 (native dataset) or 1.626 M (NH4)SO4, 0.1 M Bis-Tris pH 6.5 (zinc SAD dataset). Crystals were transferred into reservoir solution supplemented with 20% (vol/vol) glycerol for cryo-protection prior to flash-cooling by plunging into liquid nitrogen. X-ray diffraction data were recorded at beam line X06DA (PXIII) at Swiss Light Source (Paul Scherrer Institute, Villigen, Switzerland) on a PILATUS 2 M-F (Dectris) detector, at a wavelength of 1.28095 Å using an oscillation range of 0.1° and an exposure time of 0.1 s per image while rotating the crystal through 360°. Detailed data collection statistics are listed in Table 1. Diffraction data were processed with XDS (Kabsch, 2010) in space group P21, with four complex copies in the asymmetric unit and the presence of pseudmerohedral twinning. Twin law h, -k, -h-l was determined using phenix.xtriage (Zwart and Grosse-Kunstleve, 2005) comprising a twin fraction of approximately 48%. Exploiting the presence of zinc ions bound to CPSF30, phase determination was performed by single-wavelength anomalous diffraction (SAD) followed by phasing and density modification with autoSHARP (Vonrhein et al., 2007). A homology model based on CPSF30 ZF2 (PDB ID: 6FUW) was fitted into the electron density in Coot (Emsley and Cowtan, 2004), followed by automated model building using phenix.autobuild (Terwilliger et al., 2008). The structure was completed by iterative cycles of manual model building in Coot and refinement with phenix.refine (Adams et al., 2010). Molecular models were visualized using PyMOL (Schrödinger LLC, 2021).

Table 1
Crystallographic data collection and refinement statistics.
hFip1–CPSF30hFip1–CstF77
Data collection
Space groupP21P6122
Cell dimensions
a, b, c (Å)60.127, 115.125, 66.444157.612, 157.612, 161.005
α, β, γ (°)90, 116.781, 9090, 90, 120
Wavelength (Å)1.28091.0000
Resolution (Å)48.65–2.201 (2.28–2.201)56.31–2.55 (2.641–2.55)
Total reflections226,720 (15,294)1,577,004 (162,259)
Unique reflections37,698 (3244)38,981 (3836)
Rmerge (%)7.5 (95.9)9.2 (186.1)
Rpim (%)3.2 (46.9)1.5 (28.8)
I/σI13.5 (1.1)36.0 (2.6)
Cc(1/2)0.998 (0.557)1 (0.836)
Completeness (%)92.3 (80.22)99.96 (100.00)
Redundancy6.0 (4.7)40.5 (42.3)
Refinement
Resolution (Å)48.65–2.20156.31–2.55
No. reflections37,69838,975
Rwork / Rfree0.2406/0.26220.2410/0.2647
No. non-hydrogen atoms
 Protein46075188
 Ligand/ion898
 Water6725
B-factors (Å2)
 Protein56.5365.34
 Ligand/ion63.6969.46
 Water49.8355.9
R.m.s. deviations
 Bond lengths (Å)0.0080.009
 Bond angles (°)1.031.1
Ramachandran plot
 % favored95.8397.9
 % allowed4.172.1
 % outliers00
  1. Values in parentheses are for highest resolution shell.

CstF77–hFip1 complex structure determination

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A 1.5-fold molar excess of hFip1 was added per CstF77 molecule (corresponding to a threefold molar excess to a dimer of CstF77) and concentrated to 13.7 mg ml−1 (A280 = 23.36) using a centrifugal filter (Amicon Ultra-0.5, MWCO 3 kDa, Merck Millipore) prior to crystallization. The CstF77–hFip1 complex was crystallized using the sitting drop vapor diffusion method by mixing 0.1 µl protein with 0.1 µl reservoir solution containing 0.1 M Bicine pH 9.0, 10% (wt/vol) PEG 20 k, 2% (vol/vol) Dioxane. Crystals were cryo-protected by transfer into reservoir solution supplemented with 24% (vol/vol) glycerol prior to flash-cooling with liquid nitrogen. X-ray diffraction data were recorded at beam line X06SA (PXI) at Swiss Light Source (Paul Scherrer Institute, Villigen, Switzerland) on an EIGER 16 M (Dectris) detector, using an oscillation range of 0.2° and an exposure time of 0.1 s per image while rotating the crystal through 360°. Detailed data collection statistics are listed in Table 1. Diffraction data were processed with Autoproc (Vonrhein et al., 2011) in space group P6122. The structure was solved using residues 241–549 of murine CstF77 (PDB ID: 2OOE) as search model for phasing with molecular replacement (MR) in phenix.phaser (McCoy et al., 2007). A total of two CstF77 molecules could be placed into the electron density, corresponding to a dimer. After rigid-body refinement of the molecular replacement solution, the structure was completed by iterative cycles of manual model building in Coot, including the placement of the hFip1 peptides into the electron density unoccupied by CstF77, and refinement with phenix.refine (Adams et al., 2010). Molecular models were visualized using PyMOL (Schrödinger LLC, 2021).

Pull-down assays

For all pull-down assays, bound proteins were eluted with 1× SDS–PAGE loading buffer on ice and analyzed by SDS–PAGE on 4–20% Mini-PROTEAN TGX Precast Protein Gels (Bio-Rad) without prior heating to preserve the GFP fluorescence. GFP fluorescence was visualized on a Typhoon FLA 9500 laser scanner (Cytiva) at 473 nm and subsequently stained with Coomassie brilliant blue R250.

Pull-down analysis of mPSF–PAP interaction

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Strep-Tactin purified mPSF complexes were incubated with 30 µl Anti-FLAG M2 magnetic beads (Sigma-Aldrich) equilibrated in FLAG wash buffer (25 mM Tris–HCl pH 8.0, 200 mM NaCl, 0.1% Tween-20) and gently agitated at 4°C for 1 hr. The beads were washed three times with 0.5 ml of FLAG wash buffer supplemented with 2 mM MgCl2 and 165 µg PAP and 17 µg 27-nt substrate based on the SV40 pre-mRNA containing a PAS and a 3′ penta-A tail (CUGCAAUAAACAACUUAACAACAAAAA) were added to the mixture and gently agitated at 4°C for 1 hr. The beads were washed three times with 0.5 ml of FLAG wash buffer supplemented with 2 mM MgCl2 and the bound protein was eluted with 1× SDS–PAGE loading buffer supplemented with 100 µg ml−1 3× FLAG peptide (Sigma-Aldrich) on ice. FLAG elutions were analyzed by SDS–PAGE on 4–20% Mini-PROTEAN TGX Precast Protein Gels (Bio-Rad) without prior heating to preserve the GFP fluorescence. GFP fluorescence was visualized on a Typhoon FLA 9500 laser scanner (Cytiva) at 473 nm and subsequently stained with Coomassie brilliant blue R250. For equal mPSF complex concentrations to compare the corresponding GFP-hFip1 and GFP-PAP fluorescence intensities, loading volumes were adjusted according to CPSF160 band intensities. Beads control loading volume corresponds to the maximum mPSF sample loading volume.

Pull-down analysis of hFip1–CstF77 interaction

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For pull-down analysis with purified hFip1 and CstF77 proteins (wt and mutants), 10 µg of purified His6-GST-hFip1 protein was immobilized on 15 µl Glutathione Sepharose 4 Fast Flow beads (Cytiva) and washed three times with 0.5 ml pull-down wash buffer (20 mM Tris–HCl pH 7.5, 200 mM NaCl, 0.05% Tween-20, 0.5 mM TCEP). His6-MBP-CstF77 protein was added to the immobilized protein at fourfold molar excess and incubated gently agitating at 4°C for 1 hr followed by washing three times with 0.5 ml of pull-down wash buffer. The bound protein was eluted at room temperature by adding 1× SDS–PAGE loading buffer and analyzed by SDS–PAGE on 4–20% Mini-PROTEAN TGX Precast Protein Gels (Bio-Rad) stained with Coomassie brilliant blue R250.

Pull-down analysis of hFip1–CstF77–PAP interaction

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For competitive pull-down analysis of both CstF77 and PAP with hFip11–195, 5 µg of purified His6-GST-hFip11–195 protein was immobilized on 15 µl Glutathione Sepharose 4 Fast Flow beads (Cytiva) equilibrated in pull-down wash buffer, gently agitated at 4°C for 1 hr, and washed three times with 0.5 ml pull-down wash buffer. His6-MBP-CstF77 and His6-GFP-PAP were incubated with the bait, either individually or combined (1:1) at fourfold molar excess, as well as adding a 32-fold molar excess of one protein while keeping the other at fourfold molar excess, resulting in an eightfold excess of one protein over the other (8:1, 1:8).

Pull-down analysis of hFip1–CPSF30 interaction

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For pull-down analysis of hFip1 mutants binding to CPSF30, 120 µg of purified His6-MBP-TEV-CPSF301–243 wt protein was incubated with 120 µl amylose resin (NEB) equilibrated in pull-down wash buffer and gently agitated at 4°C for 1 hr. The beads were washed three times with 0.5 ml of pull-down wash buffer and equally distributed in four tubes. His6-GFP-TEV-hFip11–195 wt and point mutants (W150E, F161E, and W170E) were added in fivefold molar excess to the beads. After incubation at 4°C for 1 hr, gently agitated, unbound protein was washed off by adding three times 0.5 ml pull-down wash buffer. For pull-down analysis of the hFip1 interaction with the ZF of CPSF30, 15 µg of His6-MBP-TEV-CPSF301–243 wt and ZF mutants were incubated each with 30 µl amylose resin (NEB) equilibrated in pull-down wash buffer and gently agitated at 4°C for 1 hr. Unbound protein was washed off three times with 0.5 ml of pull-down wash buffer and His6-GFP-TEV-hFip11–195 wt was added in fourfold molar excess to the resin and incubated at 4°C for 1 hr, gently agitated. Beads were washed three times with 0.5 ml pull-down wash buffer.

Pull-down analysis of hFip1–PAP interaction

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For pull-down analysis of the hFip1:PAP interaction, 20 µg of His6-GST-TEV-hFip1 truncation constructs (hFip11–195, hFip136–195, hFip180–195, hFip136–80, and hFip11–35) were incubated each with 15 µl Glutathione Sepharose 4 Fast Flow beads (Cytiva) equilibrated in pull-down wash buffer and gently agitated at 4°C for 1 hr. Unbound protein was washed off three times with 0.5 ml of pull-down wash buffer and His6-GFP-TEV-PAP1–504 was added in fourfold molar excess to the resin and incubated at 4°C for 1 hr, gently agitated. Beads were washed three times with 0.5 ml pull-down wash buffer.

Pull-down analysis of mPSF–CstF77 interaction

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For pull-down analysis of the mPSF:CstF77 interaction, Ni-IMAC purified mPSF complexes from Sf9 cells containing hFip11–195 and N-terminal truncations hereof (hFip136–195, hFip180–195) were incubated with 20 µl Strep-Tactin (IBA Lifesciences) beads in buffer containing 20 mM HEPES–KOH pH 8.0, 150 mM KCl, 0.05% Tween-20, 0.5 mM TCEP and gently agitated at 4°C for 1 hr. Unbound protein was washed off three times with 0.5 ml of buffer containing 20 mM HEPES–KOH pH 8.0, 150 mM KCl, 0.05% Tween-20, 0.5 mM TCEP, and 10 µg His6-MBP-TEV-CstF7721–549 was added to the resin and incubated at 4°C for 1 hr, gently agitated. Beads were again washed three times with 0.5 ml.

Pull-down analysis of hFip1–CstF interaction

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For pull-down analysis with purified hFip1, CstF complex, and CstF77, 10 µg of purified His6-GST-hFip1 protein was immobilized on 15 µl Glutathione Sepharose 4 Fast Flow beads (Cytiva) and washed three times with 0.5 ml pull-down wash buffer (20 mM Tris–HCl pH 7.5, 200 mM NaCl, 0.05% Tween-20, 0.5 mM TCEP). CstF complex and His6-MBP-CstF77 protein were added to the immobilized protein at fourfold molar excess and incubated gently agitating at 4°C for 1 hr followed by washing three times with 0.5 ml of pull-down wash buffer. The bound protein was eluted at room temperature by adding 1× SDS–PAGE loading buffer and analyzed by SDS–PAGE on 4–20% Mini-PROTEAN TGX Precast Protein Gels (Bio-Rad) stained with Coomassie brilliant blue R250.

In vitro polyadenylation assays

Reaction conditions for pre-mRNA polyadenylation were adjusted for the individual need of each assay and evolved over the course of the project. To account for potential impurities and to ensure equal mPSF complex concentrations, CPSF160 band intensities were assessed on SDS–PAGE (Figure S1C) and concentrations adjusted accordingly. All polyadenylation reactions were performed in polyadenylation buffer (25 mM Tris–HCl pH 7.5, 10% glycerol, 50 mM KCl, 2 mM MgCl2, 0.05% Tween-20, 1 mM DTT) with 20 nM 5′ Cy5-labeled a 27-nt RNA substrate based on the SV40 pre-mRNA containing a PAS and a 3′ penta-A tail (CUGCAAUAAACAACUUAACAACAAAAA). All proteins were first diluted in polyadenylation buffer. Protein–RNA mixes with a total volume of 36 µl were prepared on ice, preheated at 37°C for 1 min and reaction was started by the addition of preheated 12 µl ATP at 37°C. Reaction mix for polyadenylation assay with CPSF30 ZF4/ZF5 mutants (Figure 2A) contained 80 nM mPSF complexes, 1.46 µM PAP, and a final concentration of 4 µM ATP. Reaction mix for polyadenylation assay with hFip1 truncations (Figure 2C) contained 40 nM mPSF complexes, 120 nM PAP, and a final concentration of 500 µM ATP. Reaction mix for polyadenylation assay with CstF77 (Figure 4B) at 80 (denoted 0.5), 160 (denoted +), 320, 640, or 1280 nM (denoted 2, 4, and 8, respectively) contained 80 nM mPSF complexes, 160 nM PAP, and a final concentration of 500 µM ATP. Reaction mix for polyadenylation assay with holo-CstF at 80 (denoted +), 160, 320, and 640 nM (denoted 2, 4, and 8, respectively) contained 80 nM mPSF complexes, 80 nM PAP, and a final concentration of 500 µM ATP. Time points were taken at indicated times (1 and 10 min) and polyadenylation stopped by the addition of Ethylenediaminetetraacetic acid (EDTA) with final concentration of 166 mM and incubation with 20 µg Proteinase K at 37°C for 10 min. The reactions were mixed with 2× denaturing PAGE loading dye (90% formamide, 5% glycerol, 25 mM EDTA, bromophenol blue), incubated at 95°C for 10 min and analyzed on a 15% denaturing PAGE gel containing 8 M urea and 0.5× Tris–borate–EDTA (TBE) buffer with low range ssRNA ladder (NEB). Gel was subsequently stained with SYBR Gold nucleic acid stain (Invitrogen). In-gel fluorescence of 5′ Cy5-labeled RNA and SYBR Gold-stained ssRNA ladder was visualized with Typhoon FLA 9500 laser scanner (Cytiva) at 635 and 473 nm, respectively.

Polyadenylation assay with titration of CstF77

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Polyadenylation reactions were performed according to standard procedure described above. Reaction mix for polyadenylation assay with varying CstF77 or CstF77mut concentrations (Figure 4—figure supplement 1A) contained 80 nM of mPSF complex (CPSF160-WDR331–410-CPSF30-hFip11–198), 160 nM PAP1–504, and a final concentration of 500 µM ATP. His6-TEV-CstF7721–549 or His6-TEV-CstF77mut,21–549 were added at 160 (denoted as +) or 1280 nM (denoted 8).

Polyadenylation assay with L3 mRNA and titration of CstF77

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Polyadenylation reactions were performed according to the standard procedure described above with 20 nM 5′ Cy5-labeled 38-nucleotide L3 mRNA-based substrate. Reaction mix for polyadenylation assay with varying CstF77 concentrations (Figure 4—figure supplement 1B) contained 80 nM of mPSF complex (CPSF160–WDR331–410–CPSF30–hFip11–198), 160 nM PAP1–504, and a final concentration of 500 µM ATP. His6-TEV-CstF7721–549 was added at 160 (denoted as +) or 1280 nM (denoted 8).

SEC-MALS analysis

SEC-MALS was carried out on an HPLC system (Agilent LC1100, Agilent Technologies) coupled to an Optilab rEX refractometer and a miniDAWN three-angle light-scattering detector (Wyatt Technology). Data analysis was performed using the ASTRA software (version 7.3.2; Wyatt Technology).

SEC-MALS analysis of hFip1–CPSF30 complex

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For unambigous determination of the stoichiometry of the respective hFip1–CPSF30 complexes, tagged proteins were used to increase the molecular weight difference between the 2:1 and 1:1 complexes of hFip1–CPSF30. Stoichiometry of the complexes was determined injecting 33 µg His6-MBP-CPSF301–243 (wt and mutants) and fourfold molar excess of His6-GFP-hFip11–195 premixed in a total injection volume of 100 µl. Proteins were separated on a Superdex 200 10/300 GL column (Cytiva) run at 0.5 ml/min at room temperature in 20 mM Tris–HCl pH 7.5, 200 mM NaCl, 0.5 mM TCEP (pH was adjusted at room temperature).

SEC-MALS analysis of mPSF–PAP complex

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Stoichiometry of the mPSF–PAP complex was determined injecting 50 µg of prepurified mPSF-PAP comprising CPSF160–WDR331–410–CPSF30–2xhFip11–198, PAP1–504, and 27-nt SV40 PAS-containing mRNA in a total injection volume of 100 µl. In a second run, prepurified 50 µg of mPSF:PAP was spiked with additional 41.6 µg PAP1–504 (fivefold molar excess) in a total injection volume of 100 µl to test whether excess PAP can lead to a stable 1:2 complex of mPSF and PAP. Proteins were separated on a Superose 6 10/300 GL column (Cytiva) run at 0.5 ml/min at room temperature in 20 mM HEPES pH 8.0, 150 mM KCl, 0.5 mM TCEP (pH was adjusted at room temperature).

Multiple sequence alignment

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The multiple sequence alignment of hFip1 orthologs was produced with MAFFT version 7 (Katoh et al., 2018) and visualized using Jalview (Waterhouse et al., 2009). Input sequences are listed in Supplementary file 3.

Analysis of interaction interfaces

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Buried surface area of the hFip1–CPSF30 interaction interface was calculated using the PDBe PISA (Proteins, Interfaces, Structures and Assemblies) tool (Krissinel and Henrick, 2007).

3D density map analysis

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Visualization and analysis of the 3D density map for CPSF160–WDR33–CPSF30–PAS RNA–CstF77 complex (EMD-20861) were performed with UCSF Chimera (Pettersen et al., 2004), developed by the Resource for Biocomputing, Visualization, and Informatics at the University of California, San Francisco. The 3D density map was segmented and color coded based on the corresponding atomic model (PDB ID: 6URO). The CstF77–hFip1 crystal structure from this study was superimposed onto the atomic model of CstF77.

Bioinformatic analysis of CstF77

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Color-coded electrostatic surface representation of CstF77 was generated for the biological assembly of murine CstF7720–549 (PDB ID: 2OOE) in PyMOL 2.5.0 (Schrödinger LLC, 2021) using the protein contact potential option. The CstF77241–549–hFip11–35 structure from this study and cryo-EM structure of human CPSF160–WDR33–CPSF30–PAS RNA–CstF77 complex (PDB ID: 6URO) were superimposed onto murine CstF77 using PyMOL’s align command to identify and visualize the hFip1- and mPSF-binding regions, respectively. Analysis of evolutionary conservation of CstF77 was carried out using the ConSurf web server (Ashkenazy et al., 2016) with murine CstF7720–549 (PDB ID: 2OOE) as input and applying standard settings (sequence alignment with MAFFT, homologs taken from UniRef90). The degree of conservation was visualized in PyMOL by color coding (green: variable, violet: conserved) the protein surface according to the conservation scores which are written into the tempFactor column of the ConSurf web server output PDB file.

Appendix 1

Appendix 1—key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Escherichia coli)BL21 star (DE3)Thermo Fisher scientificBL21 star (DE3)Chemically competent cells
Strain, strain background (Escherichia coli)BL21(DE3)-AIThermo Fisher scientificBL21(DE3)-AIChemically competent cells
Strain, strain background (Escherichia coli)Rosetta2 (DE3)NovagenRosetta2 (DE3)Chemically competent cells
Cell line (Spodoptera frugiperda)Sf9Thermo Fisher ScientificCat. #11496015
Recombinant DNA reagentpLM B042; pLM B043 (plasmid)This paperHolo-CstF
Recombinant DNA reagentpLM B092
(plasmid)
This paperMBP-CstF77
Recombinant DNA reagentpLM B123
(plasmid)
This paperCstF77
Recombinant DNA reagentpLM B142
(plasmid)
This paperPAP
Recombinant DNA reagentpLM B156
(plasmid)
This paperMBP-PAP
Recombinant DNA reagentpLM B157
(plasmid)
This paperGFP-PAP
Recombinant DNA reagentpLM B164
(plasmid)
This paperMBP-CstF7721–549
or MBP-CstF77
Recombinant DNA reagentpLM B168
(plasmid)
This paperMBP-CstF77mut
Recombinant DNA reagentpLM B170
(plasmid)
This paperCstF77mut
Recombinant DNA reagentpMC B051
(plasmid)
This paperCPSF30ZF4–ZF5
Recombinant DNA reagentpMC B054
(plasmid)
This paperMBP-CPSF30ZF4–ZF5
Recombinant DNA reagentpMC B055
(plasmid)
This paperCPSF30 ZF4 mutant
Recombinant DNA reagentpMC B056
(plasmid)
This paperCPSF30 ZF5 mutant
Recombinant DNA reagentpMC B057
(plasmid)
This paperCPSF30 ZF4 and ZF5 mutant
Recombinant DNA reagentpMC B058
(plasmid)
This paperCPSF30 ZF4 mutant
Recombinant DNA reagentpMC B059
(plasmid)
This paperCPSF30 ZF4 mutant
Recombinant DNA reagentpMC B060
(plasmid)
This paperCPSF30 ZF5 mutant
Recombinant DNA reagentpMC B061
(plasmid)
This paperCPSF30 ZF5 mutant
Recombinant DNA reagentpMC B062
(plasmid)
This paperCPSF30 ZF4 and ZF5 mutant
Recombinant DNA reagentpMC B063
(plasmid)
This paperCPSF30 ZF4 and ZF5 mutant
Recombinant DNA reagentpMC C011
(plasmid)
This paperhFip1CD
Recombinant DNA reagentpMC C015
(plasmid)
This paperGST-hFip1 fragment or hFip180–195
Recombinant DNA reagentpMC C030
(plasmid)
This paperhFip1CD
Recombinant DNA reagentpMC C049
(plasmid)
This paperGFP-hFip1
Recombinant DNA reagentpMC C050
(plasmid)
This paperGST-hFip1 fragment or hFip136–80
Recombinant DNA reagentpMC C059
(plasmid)
This paperGST-hFip1 fragment, GST-hFip11–35, or hFip11–35
Recombinant DNA reagentpMC C060
(plasmid)
This paperGST-hFip1 fragment, GST-hFip11–195, or hFip11–195
Recombinant DNA reagentpMC C066
(plasmid)
This paperHis6-GFP-TEV-hFip11–195 point mutant
Recombinant DNA reagentpMC C067
(plasmid)
This paperHis6-GFP-TEV-hFip11–195 point mutant
Recombinant DNA reagentpMC C068
(plasmid)
This paperHis6-GFP-TEV-hFip11–195 point mutant
Recombinant DNA reagentpMC C073
(plasmid)
This paperGST-hFip1 fragment, GST-hFip136–195, or hFip136–195
Recombinant DNA reagentpMC C093
(plasmid)
This papermutant GST-hFip11–35
Recombinant DNA reagentpMC C094
(plasmid)
This papermutant GST-hFip11–35
Recombinant DNA reagentpMC C096
(plasmid)
This papermutant GST-hFip11–35
Recombinant DNA reagentpMC N015
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018A
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018C-2
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-0
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-8
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-10
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-12
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-14
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-15
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018G-21
(plasmid)
This paperFLAG-epitope-tagged mPSF
Recombinant DNA reagentpMC N018G-22
(plasmid)
This paperFLAG-epitope-tagged mPSF
Recombinant DNA reagentpMC N018G-23
(plasmid)
This paperFLAG-epitope-tagged mPSF
Recombinant DNA reagentpMC N018G-24
(plasmid)
This paperFLAG-epitope-tagged mPSF
Recombinant DNA reagentpMC N018H
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018I
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018J
(plasmid)
This papermPSF
Recombinant DNA reagentpMC N018K
(plasmid)
This papermPSF
Sequence-based reagentrLM 011This paper27 nt RNA substrate based on SV40 pre-mRNA; CUGCAAUAAACAACUUAACAACAAAAA
Sequence-based reagentrLM 015This paper5′ Cy5-labeled 27 nt RNA substrate based on SV40 pre-mRNA; CUGCAAUAAACAACUUAACGUCAAAAA
Sequence-based reagentrLM 016This paper5′ Cy5-labeled 27 nt RNA substrate based on SV40 pre-mRNA; CUGCAGUACACAACUUAACGUCAAAAA
Sequence-based reagentrLM 031This paper5′ Cy5-labeled 38 nt RNA substrate based on adenoviral L3 pre-mRNA; ACUUUCAAUAAAGGCAAAUGUUUUUAUUUGUACAAAAA

Data availability

X-ray diffraction data (atomic coordinates and structure factors) have been submitted to the PDB and will be released upon publication.

The following data sets were generated
    1. Muckenfuss L
    2. Jinek M
    (2022) RCSB Protein Data Bank
    ID 7ZYH. Crystal structure of human CPSF30 in complex with hFip1.
    1. Muckenfuss L
    2. Jinek M
    (2022) RCSB Protein Data Bank
    ID 7ZY4. Crystal structure of human CstF77 in complex with hFip1.
The following previously published data sets were used
    1. Sun Y
    2. Zhang Y
    3. Walz T
    4. Tong L
    (2019) RCSB Protein Data Bank
    ID 6URO. Cryo-EM structure of human CPSF160-WDR33-CPSF30-PAS RNA-CstF77 complex.
    1. Sun Y
    2. Zhang Y
    3. Walz T
    4. Tong L
    (2019) Electron Microscopy Data Bank
    ID EMD-20861. Cryo-EM structure of human CPSF160-WDR33-CPSF30-PAS RNA-CstF77 complex.

References

    1. Emsley P
    2. Cowtan K
    (2004) Coot: model-building tools for molecular graphics
    Acta Crystallographica. Section D, Biological Crystallography 60:2126–2132.
    https://doi.org/10.1107/S0907444904019158
    1. Kabsch W
    (2010) XDS
    Acta Crystallographica. Section D, Biological Crystallography 66:125–132.
    https://doi.org/10.1107/S0907444909047337
  1. Software
    1. Schrödinger LLC
    (2021)
    The pymol molecular graphics system, version 2.5.0
    PyMOL.
    1. Zwart PH
    2. Grosse-Kunstleve R
    (2005)
    Xtriage and fest: automatic assessment of X-ray data and substructure structure factor estimation
    CCP4 Newsl 43:27–35.

Decision letter

  1. Eric J Wagner
    Reviewing Editor; University of Rochester Medical Center, United States
  2. James L Manley
    Senior Editor; Columbia University, United States
  3. William Marzluff
    Reviewer; University of North Carolina, United States

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Fip1 is a multivalent interaction scaffold for processing factors in human mRNA 3' end biogenesis" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by James Manley as the Senior Editor. The following individual involved in the review of your submission has agreed to reveal their identity: William Marzluff (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission. We note that no further experimentation is requested. Rather, textual changes are suggested to further strengthen the manuscript. The two critical components of these suggested changes are highlighted in the 'essential revisions' but we encourage the authors to consider response/changes in accordance with other points raised by the Reviewers as described below.

Essential revisions:

1) Through thoughtful discussion, please address differences in approaches used to support the idea that a single copy of PAP exists in the complex in contrast to a previous publication. There are caveats to both studies and distinct experimental differences that should be highlighted to temper their overall narrative.

2) Reviewers were particularly interested in the CstF77 data, specifically the interpretation. This is potentially important data to the field and should be discussed more in this study.

Reviewer #1 (Recommendations for the authors):

In the study by Muckenfuss et al., the authors provide two independent crystal structures of Fip1 in complex with CPSF30 or CstF77 as a basis for biochemical experiments exploring the function of Fip1 in polyadenylation. The structure of CPSF30/Fip1 is validated using polyadenylation assays and the authors conclude that while two molecules of Fip1 interact with a single molecule of CPSF30 there is likely only one copy of PAP brought to this subcomplex. Secondary to this, the authors present the first structure of Fip1 associating with CstF77 and provide biochemical validation of this model. Interestingly, the authors show that excess CstF77 leads to inhibition of in vitro polyadenylation of mPSF through the likely mutually exclusive association of Fip1 with 77 and PAP. Overall, the strengths of the study lie in high-resolution structural biology coupled with careful biochemical assays. My main concerns lie in the reduced novelty of the first half of this paper with the Hamilton and Tong study previously published. The main additional detail is whether one or two PAP molecules can associate with CPSF30/Fip1 and this seems somewhat incremental. There is a novelty to the observations that CstF77 can inhibit polyadenylation but this portion of the study is not as developed as it could be. If this portion of the paper were modestly expanded, I could be more convinced that it is worthy of publication. Several specific comments and suggestions are listed below.

1. The authors contend that their data support a model whereby a single copy of PAP is recruited to Fip1/CPSF30 in contrast to what was observed by Hamilton and Tong. It is important to note the experimental differences between the two systems: in this study, a GFP-PAP was used for in vitro binding, whereas Hamilton and Tong used only the catalytic module (1-524) of PAP; in this study, the authors use mPSF-PAP factors whereas in Hamilton and Tong full-length CPSF30 was used and Fip1 (79-200) was used. While I appreciate that experiments presented here appear to be done in a more 'complete' biochemical context, I am equally concerned that the use of GFP-PAP can somehow inhibit the ability of PAP to associate with Fip1/mPSF.

2. Does the inhibitory effect of CstF77 apply to other mRNA substrates beyond the one tested in this study?

3. Would supplementing the polyadenylation assays with the CstF complex (50/64/77) also cause inhibition, or is it unique to supplementation with isolated CstF77? If this were to be the case, how would this be interpreted?

Reviewer #3 (Recommendations for the authors):

1. The authors need to mention the role of FIP1 in alternative polyadenylation, and that it specifically promotes polyadenylation of mRNAs with U-rich stretches preceding the polyadenylation signal. The experiments of Shi et al. (Lackford et al., 2014), show that knockdown of Fip1 has a dramatic effect on alternative polyadenylation, resulting in skipping upstream polyA sites that have U-rich regions before the AAUAAA that bind Fip1. One potential interpretation of these experiments is that if FIp1 is limiting you can form an active polyadenylation complex that lacks Fip1, and complexes that lack Fip1 fail to cleave and polyadenylate at a subset of sites that require the U-rich elements for cleavage.

2. In interpreting the overall data, the authors should discuss possibilities of what actually happens in the cell when the cleavage complex forms on the nascent pre-mRNA. Their (and others) in vitro data on the properties of complexes not bound to substrate RNA describe the properties of complexes that may not be relevant to the functional complex that is bound to the pre-mRNA just prior to cleavage.

For example, they show CstF77 inhibits polyadenylation by interfering with the binding to Fip1. Since CstF and CPSF must interact during cleavage, this suggests that before cleavage (when all factors are bound to the substrate and RPPP6 is being recruited), PAP is likely not bound to the complex. After cleavage, either CstF may dissociate from the complex (since the 3' fragment of the RNA is bound to CstF) or alter its conformation to allow PAP to bind for polyadenylation. The authors might want to discuss these possibilities. This may be part of the reason there is a discrepancy in the literature on whether or not PAP is required for cleavage. If their model is correct PAP is likely not in the active cleavage complex (which contains CstF77), but might on the mPSF (or CPSF) that binds initially to the polyadenylation signal, but is lost when the active cleavage complex, which includes CstF forms.

3. They need to explicitly mention on page 5 (description of Figure 1) whether there are any differences between their structure and the structure of Tong and Hamilton of what seems to be the same complex.

4. They say (l. 138-139) "both Fip1 binding sites contribute to the integrity of stability of mPSF. Might be better to say that "either FIP1 binding site can contribute to the integrity of mPSF." The complexes with one FIp11 binding site look similar to those with 2 sites (wt) in Figure 1G.

5. In Figure 2A, it looks as if in mutant ZF4 or ZF5 polyadenylation is no longer processive. In the wild-type how long are the A-tails that accumulate? They should mention that. In the mutant AGUACA (I presume that is a mutant of the AAUAAA?) there is an addition of 3-6 A tails. It looks like the Y127E mutant is a little more active than the Y151E mutant (based on the length of the A tails). If that is reproducible that they could mention that.

6. The evidence that only one PAP is present in mPSF is convincing, and either FIP1 binding site seems by itself to be able to recruit PAP.

7. They describe and characterize the structure of a complex of the N-terminal 35 aa of Fip 1 with CstF77, adding a third interaction to the previously described interactions of CstF77 with Wdr33 and CPSF160. Since there are two copies of CstF77 in CstF, CstF can bind two molecules of Fip1, and they provide evidence that happens in mCSF, and is likely part of the mechanism by which mPSF and CstF interact during cleavage.

https://doi.org/10.7554/eLife.80332.sa1

Author response

Essential revisions:

1) Through thoughtful discussion, please address differences in approaches used to support the idea that a single copy of PAP exists in the complex in contrast to a previous publication. There are caveats to both studies and distinct experimental differences that should be highlighted to temper their overall narrative.

2) Reviewers were particularly interested in the CstF77 data, specifically the interpretation. This is potentially important data to the field and should be discussed more in this study.

Reviewer #1 (Recommendations for the authors):

In the study by Muckenfuss et al., the authors provide two independent crystal structures of Fip1 in complex with CPSF30 or CstF77 as a basis for biochemical experiments exploring the function of Fip1 in polyadenylation. The structure of CPSF30/Fip1 is validated using polyadenylation assays and the authors conclude that while two molecules of Fip1 interact with a single molecule of CPSF30 there is likely only one copy of PAP brought to this subcomplex. Secondary to this, the authors present the first structure of Fip1 associating with CstF77 and provide biochemical validation of this model. Interestingly, the authors show that excess CstF77 leads to inhibition of in vitro polyadenylation of mPSF through the likely mutually exclusive association of Fip1 with 77 and PAP. Overall, the strengths of the study lie in high-resolution structural biology coupled with careful biochemical assays. My main concerns lie in the reduced novelty of the first half of this paper with the Hamilton and Tong study previously published. The main additional detail is whether one or two PAP molecules can associate with CPSF30/Fip1 and this seems somewhat incremental. There is a novelty to the observations that CstF77 can inhibit polyadenylation but this portion of the study is not as developed as it could be. If this portion of the paper were modestly expanded, I could be more convinced that it is worthy of publication. Several specific comments and suggestions are listed below.

We thank the Reviewer for the positive feedback and the constructive suggestions.

1. The authors contend that their data support a model whereby a single copy of PAP is recruited to Fip1/CPSF30 in contrast to what was observed by Hamilton and Tong. It is important to note the experimental differences between the two systems: in this study, a GFP-PAP was used for in vitro binding, whereas Hamilton and Tong used only the catalytic module (1-524) of PAP; in this study, the authors use mPSF-PAP factors whereas in Hamilton and Tong full-length CPSF30 was used and Fip1 (79-200) was used. While I appreciate that experiments presented here appear to be done in a more 'complete' biochemical context, I am equally concerned that the use of GFP-PAP can somehow inhibit the ability of PAP to associate with Fip1/mPSF.

We thank the Reviewer for the comment upon which we critically examined our experimental setup. While we indeed used GFP-PAP for our in vitro pull-down assays (Figure 1G, 2B, 4A), it is important to note that an untagged catalytic domain of human PAP (1-504) was used for copurification of a mPSF-PAP complex and its subsequent SEC-MALS analysis in isolation and in the presence of excess untagged PAP (Figure 2D). In fact, both mPSF and PAP are untagged in this analysis which ensures that any interference from affinity/epitope tags is eliminated.

Initially, we used untagged PAP for our in vitro pull-down analysis. However, due to the transient interaction of PAP with mPSF/hFip1 and the resulting weak bands on SDS-PAGE, we decided to use GFP-PAP for improved detection by in-gel fluorescence of the N-terminally fused GFP tag. For reference, a comparative pull-down analysis using untagged and tagged PAP with mPSF complexes is shown in Author response image 1. Equal amounts of PAP are precipitated by Strep-tagged mPSF complexes, independent whether it is fused to a GFP tag or not.

Author response image 1

Furthermore, analysis of the three-dimensional structure of PAP reveals that the hFip1-binding site on PAP is located on the opposite face of the molecule relative to its N-terminus. Together with the fact that PAP is about twice the size (58 kDa) of the N-terminally fused GFP tag (28 kDa), it is unlikely that the tag would negatively affect its interaction with hFip1.Therefore, we believe that the different results observed in the two studies stem from the different compositions of the complexes (mPSF in our study vs. CPSF30-hFip1 in Hamilton and Tong) used to test for their interaction with PAP. Interestingly, Hamilton and Tong observe dimerization of CPSF30-hFip1 (70-200) complexes, which we do not observe with our mPSF complexes, an additional indication that isolated CPSF30-hFip1 complexes are able to establish additional interactions that are obstructed once the complex is integrated into the mPSF.

At present, we cannot exclude the possibility that mPSF complexes assemble with two PAP molecules in vitro at higher PAP concentration; however, we believe that this does not represent the predominant physiological assembly.

We made additional revisions to the manuscript, including the Discussion section, to elaborate on the observation of only one stably bound PAP molecule.

2. Does the inhibitory effect of CstF77 apply to other mRNA substrates beyond the one tested in this study?

We thank the Reviewer for raising this point. It is important to note that CstF64, rather than CstF77, is the principal RNA-binding component of the CstF complex that mediates interactions with G/U-rich downstream sequences through its RNA recognition motif (RRM) (Takagaki et al., 1997). CstF64, however, was not included in our analysis of CstF77-mediated inhibition of polyadenylation (and neither was CstF50) (Figure 4B, Figure 4 —figure supplement 1); furthermore, our use of pre-cleaved RNA substrates precludes the inclusion of downstream elements. As CstF77 does not possess an RRM and is not known to directly bind RNA (Bai et al., 2007; Yang et al., 2018), competition between CstF77 and PAP is expected to be mediated by protein-protein interactions rather than via the RNA substrate.

Following the suggestion of the Reviewer, we probed the inhibitory effect of CstF77 on polyadenylation with an alternative adenoviral L3 PAS-containing mRNA substrate (38 nt), that was previously shown to be processed in vitro by a recombinant human 3’ end processing complex (Boreikaite et al., 2022; Schmidt et al., 2022). Compared to SV40 mRNA, in which the PAS and cleavage site are separated by only 11 nt, these sites are separated by 20 nt in the L3 mRNA, which is within the normal range (15-21 nt) observed for mammalian pre-RNAs (Beaudoing et al., 2000; Gruber et al., 2016). We observe a similar degree of inhibition by CstF77 using the alternative adenoviral L3 mRNA substrate (Figure 4 —figure supplement 1B), suggesting that the inhibition is independent of the RNA substrate.

3. Would supplementing the polyadenylation assays with the CstF complex (50/64/77) also cause inhibition, or is it unique to supplementation with isolated CstF77? If this were to be the case, how would this be interpreted?

We thank the Reviewer for this intriguing question. hFip1 interacts with both CstF77 and CstF in our pull-down analysis (now: Figure 3 —figure supplement 3), indicating that the hFip1 binding site on CstF77 is not obstructed upon assembly of CstF77 into the CstF complex. As suggested by the Reviewer, we carried out polyadenylation assay using a recombinant holoCstF complex comprising CstF77, CstF641-198, and CstF50 (now: Figure 4C) to demonstrate that holo-CstF causes inhibition of the polyadenylation reaction to a similar extent as isolated CstF77, suggesting that the inhibitory effect of CstF77 persists when it is assembled within CStF. As indicated, we added two additional figures to the manuscript (Figure 3 —figure supplement 3; Figure 4C) that further strengthen our findings on CstF-mediated inhibition of polyadenylation.

Reviewer #3 (Recommendations for the authors):

1. The authors need to mention the role of FIP1 in alternative polyadenylation, and that it specifically promotes polyadenylation of mRNAs with U-rich stretches preceding the polyadenylation signal. The experiments of Shi et al. (Lackford et al., 2014), show that knockdown of Fip1 has a dramatic effect on alternative polyadenylation, resulting in skipping upstream polyA sites that have U-rich regions before the AAUAAA that bind Fip1. One potential interpretation of these experiments is that if FIp1 is limiting you can form an active polyadenylation complex that lacks Fip1, and complexes that lack Fip1 fail to cleave and polyadenylate at a subset of sites that require the U-rich elements for cleavage.

We thank the Reviewer for this comment. We now mention the role of hFip1 in alternative polyadenylation through modulating the selection of cleavage sites via its interaction with Urich sequence elements as well as cleavage factor Im in both the Introduction section of the manuscript, as well as in the Discussion.

2. In interpreting the overall data, the authors should discuss possibilities of what actually happens in the cell when the cleavage complex forms on the nascent pre-mRNA. Their (and others) in vitro data on the properties of complexes not bound to substrate RNA describe the properties of complexes that may not be relevant to the functional complex that is bound to the pre-mRNA just prior to cleavage.

For example, they show CstF77 inhibits polyadenylation by interfering with the binding to Fip1. Since CstF and CPSF must interact during cleavage, this suggests that before cleavage (when all factors are bound to the substrate and RPPP6 is being recruited), PAP is likely not bound to the complex. After cleavage, either CstF may dissociate from the complex (since the 3' fragment of the RNA is bound to CstF) or alter its conformation to allow PAP to bind for polyadenylation. The authors might want to discuss these possibilities. This may be part of the reason there is a discrepancy in the literature on whether or not PAP is required for cleavage. If their model is correct PAP is likely not in the active cleavage complex (which contains CstF77), but might on the mPSF (or CPSF) that binds initially to the polyadenylation signal, but is lost when the active cleavage complex, which includes CstF forms.

We thank the Reviewer for raising these points. We have revised our Discussion section to expand on these ideas and discuss the possibilities, also with respect to the conflicting data in the literature on the requirement of PAP for pre-mRNA cleavage.

3. They need to explicitly mention on page 5 (description of Figure 1) whether there are any differences between their structure and the structure of Tong and Hamilton of what seems to be the same complex.

We note that although the structures are highly similar, they are not identical due to the different choices of protein construct boundaries. As a result, we have been able to resolve an additional 21 amino acid residues, mostly in the N- and C-terminal extensions of hFip1CDb. We have revised the relevant Results section to mention the similarities and differences between the two structures.

4. They say (l. 138-139) "both Fip1 binding sites contribute to the integrity of stability of mPSF. Might be better to say that "either FIP1 binding site can contribute to the integrity of mPSF." The complexes with one FIp11 binding site look similar to those with 2 sites (wt) in Figure 1G.

We thank the Reviewer for the comments. We have reworded the section as suggested. However, we respectfully disagree with the comment that the SDS-PAGE bands of GFP-PAP precipitated by single hFip1 and double hFip1 complexes (WT mPSF) are of similar intensities. Densitometric quantitation of the GFP fluorescence using ImageLab 6.1 software (see Author response table 1) reveals a marked reduction of both GFP-hFip1 and GFP-PAP band intensities upon mutation of either of the two hFip1 binding sites in CPSF30 (single hFip1 complexes) that consequently results in reduced recruitment of PAP. Nevertheless, so as not to over-interpret this data, we distinguish only between the presence or absence of interaction.

Author response table 1
Quantitation of co-precipitated GFP-hFip1 and GFP-PAP protein levels (GFP fluorescence detected at 473 nm), normalized against WT mPSF.
wtZF4ZF5
GFP-hFip10.290.42
GFP-PAP10.280.43

5. In Figure 2A, it looks as if in mutant ZF4 or ZF5 polyadenylation is no longer processive. In the wild-type how long are the A-tails that accumulate? They should mention that. In the mutant AGUACA (I presume that is a mutant of the AAUAAA?) there is an addition of 3-6 A tails. It looks like the Y127E mutant is a little more active than the Y151E mutant (based on the length of the A tails). If that is reproducible that they could mention that.

We thank the Reviewer for bringing up these points. We now added molecular weight markers to the gels showing polyadenylation assays. Furthermore, we specified more clearly that AGUACA is a mutant of the AAUAAA polyadenylation signal. As mPSF retains residual low affinity RNA binding activity in the absence of the canonical AAUAAA hexanucleotide (Clerici et al., 2017), residual polyadenylation activity can be observed for this RNA substrate. Although the slight difference in polyadenylation efficiencies of the Y127E and Y151E mutant is reproducible, we cannot exclude that this is due to the Y127E CPSF30 mutant (ZF4) still retaining some affinity for hFip1 (Figure 1E) as it has higher affinity for hFip1 than ZF5 does (Hamilton et al., 2020). Therefore, we refrained from further discussion of the results so as not to overinterpret the data.

6. The evidence that only one PAP is present in mPSF is convincing, and either FIP1 binding site seems by itself to be able to recruit PAP.

We thank the Reviewer for the positive feedback.

https://doi.org/10.7554/eLife.80332.sa2

Article and author information

Author details

  1. Lena Maria Muckenfuss

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Conceptualization, Formal analysis, Investigation, Methodology, Writing - original draft
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3558-7211
  2. Anabel Carmen Migenda Herranz

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  3. Franziska Maria Boneberg

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  4. Marcello Clerici

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Conceptualization, Investigation
    Competing interests
    No competing interests declared
  5. Martin Jinek

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    jinek@bioc.uzh.ch
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7601-210X

Funding

Schweizerischer Nationalfonds zur Förderung der Wissenschaftlichen Forschung (NCCR RNA and Disease)

  • Martin Jinek

Boehringer Ingelheim Fonds

  • Lena Maria Muckenfuss

Howard Hughes Medical Institute (55008735)

  • Martin Jinek

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Birgit Dreyer for assistance with SEC-MALS, Beat Blattmann (University of Zurich Protein Crystallization Center) for performing crystallization screens, and Levi Kopp for assistance with protein crystallization. We thank Vincent Olieric, Takashi Tomizaki, and Meitian Wang (Swiss Light Source, Paul Scherrer Institute) for assistance with crystallographic data collection. We are grateful to Stefanie Jonas and members of the Jinek laboratory for critical reading of the manuscript. This work was supported by Boehringer Ingelheim Fonds PhD Fellowship and by the National Center for Competence in Research (NCCR) RNA & Disease, funded by the Swiss National Science Foundation.

Senior Editor

  1. James L Manley, Columbia University, United States

Reviewing Editor

  1. Eric J Wagner, University of Rochester Medical Center, United States

Reviewer

  1. William Marzluff, University of North Carolina, United States

Publication history

  1. Received: May 17, 2022
  2. Preprint posted: May 31, 2022 (view preprint)
  3. Accepted: September 7, 2022
  4. Accepted Manuscript published: September 8, 2022 (version 1)
  5. Version of Record published: September 26, 2022 (version 2)

Copyright

© 2022, Muckenfuss et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Lena Maria Muckenfuss
  2. Anabel Carmen Migenda Herranz
  3. Franziska Maria Boneberg
  4. Marcello Clerici
  5. Martin Jinek
(2022)
Fip1 is a multivalent interaction scaffold for processing factors in human mRNA 3′ end biogenesis
eLife 11:e80332.
https://doi.org/10.7554/eLife.80332

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