A structural biology view on the enzymes involved in eukaryotic mRNA turnover

Christina Krempl; Daniela Lazzaretti; Remco Sprangers

doi:10.1515/hsz-2023-0182

Open Access Published by De Gruyter September 15, 2023

A structural biology view on the enzymes involved in eukaryotic mRNA turnover

Christina Krempl , Daniela Lazzaretti and Remco Sprangers

From the journal Biological Chemistry

https://doi.org/10.1515/hsz-2023-0182

Abstract

The cellular environment contains numerous ribonucleases that are dedicated to process mRNA transcripts that have been targeted for degradation. Here, we review the three dimensional structures of the ribonuclease complexes (Pan2-Pan3, Ccr4-Not, Xrn1, exosome) and the mRNA decapping enzymes (Dcp2, DcpS) that are involved in mRNA turnover. Structures of major parts of these proteins have been experimentally determined. These enzymes and factors do not act in isolation, but are embedded in interaction networks which regulate enzyme activity and ensure that the appropriate substrates are recruited. The structural details of the higher order complexes that form can, in part, be accurately deduced from known structural data of sub-complexes. Interestingly, many of the ribonuclease and decapping enzymes have been observed in structurally different conformations. Together with experimental data, this highlights that structural changes are often important for enzyme function. We conclude that the known structural data of mRNA decay factors provide important functional insights, but that static structural data needs to be complemented with information regarding protein motions to complete the picture of how transcripts are turned over. In addition, we highlight multiple aspects that influence mRNA turnover rates, but that have not been structurally characterized so far.

Keywords: conformational changes; deadenylation; exoribonuclease; mRNA decapping; mRNA turnover

1 Introduction

The abundance of cellular bio-molecules is determined by the relative rates of synthesis and degradation. For eukaryotic mRNAs, synthesis is a complex hierarchical and multi-step process that sequentially involves RNA polymerase II mediated transcription, 5′ m⁷G capping, 3′ polyadenylation, splicing, export from the nucleus and, in many cases, base editing. Two main features protect mRNA transcripts against rapid and uncontrolled degradation: the 5′ m⁷G cap structure, that blocks access for the exoribonuclease Xrn1, as well as a 3′ poly(A) tail, that associates with one or multiple copies of the poly(A) binding protein (Pab1). Both elements serve multiple purposes and are e.g. also important for efficient translation.

The simplest form of the protecting m⁷G mRNA cap structure (termed cap0) consists of an N7-methylated guanosine that is connected to the 5′ end of the mRNA by a 5′–5′ triphosphate bridge (Shatkin 1976). The ribose 2′ OH group of the most 5′ RNA base is often methylated to form the cap1 structure. 2′ O-methylations of the following riboses result in the formation of cap2 to cap4 structures, where mRNA with a cap2 appears to have a mildly increased stability compared to cap1 mRNA (Despic and Jaffrey 2023) by a mechanism that remains elusive. The downstream bases in the mRNA body may also be modified in a wide range of manners, which can influence mRNA stability in multiple ways (Boo and Kim 2020), through molecular mechanisms that are not yet fully understood. Given the pivotal role of the cap in mRNA metabolism, it is desirable to make use of capped RNA in biochemical and structural studies, which can easily be produced in vitro using the capping enzyme complex from the vaccinia virus (Fuchs et al. 2016).

The length of the mRNA-protecting poly(A) tail ranges from 70 to 80 nucleotides in yeast to over 250 nucleotides in mammalian cells (Lima et al. 2017). Additionally, the length of the poly(A) tail differs between transcripts, and developmental, differentiation and cell cycle dependent stages, and has important implications for cellular function (Jalkanen et al. 2014). The 3′ poly(A) tail generally interacts specifically with Pab1, that thereby protects the 3′ end of the transcript against rapid exonucleolytic degradation (Eckmann et al. 2011; Mangus et al. 2003).

The half-lives of different mRNAs vary significantly, from 3 min to over 90 min in yeast (Wang et al. 2002), and can be even longer in human cells (Yang et al. 2003). These large variations in mRNA decay rates can in part be attributed to poly(A) tail length, but also depend on specific sequence elements (e.g. AU-rich elements in the 3′ UTR) or abnormalities in the transcript that can influence degradation rates significantly (Kurosaki et al. 2019; Xu et al. 1997). Generally, mRNA turnover is initiated by the stepwise shortening of the 3′ poly(A) tail (deadenylation), and proceeds through the action of decapping enzymes that remove the protecting mRNA cap structure and 5′→3′ or 3′→5′ exoribonucleases that degrade the transcript processively from either end.

Below, we describe the structural details of the Pan2-Pan3 and Ccr4-Not deadenylation complexes and of the 5′→3′ and 3′→5′ mRNA decay pathways. Known structures of the different sub-complexes depicted here originate from different species (e.g. human or yeast). Importantly, these complexes are highly conserved and their core structures appear to be species-independent. However, there is considerable variation in the details of the interaction between the sub-complexes and the mechanism through which additional factors are recruited.

2 mRNA turnover step 1: deadenylation by Pan2-Pan3

For canonical mRNA turnover, the degradation of the transcript is generally initiated by a two-step shortening of the 3′ poly(A) tail, which is often the rate limiting step in mRNA decay. As long as the poly(A) tail is long enough to interact with at least two copies of Pab1, the transcript is shortened by the 200–230 kDa Pan2-Pan3 complex (Brown and Sachs 1998; Schäfer et al. 2019). Multiple structures of Pan2, Pan3, and complexes thereof have been determined (Table 1), providing important insights into the mechanism of Pan2-Pan3-mediated deadenylation.

Table 1:

Structures of the Pan2-Pan3 complex.

PDB ID	Pan2	Pan3	Other
4CZW (Jonas et al. 2014)	n.c. UCH, RNase
4CZV (Jonas et al. 2014)	n.c. WD40
4Q8H (Schäfer et al. 2014)	s.c. UHC, RNAse
4Q8G (Schäfer et al. 2014)	s.c. UHC
6R9I (Tang et al. 2019)	s.c. UCH, RNase
6R9J, M, O, P, Q (Tang et al. 2019)	s.c. UHC, RNase		Different RNA substrates
4BWK, 4BWX (Christie et al. 2013)		n.c. PK, CC, CK
4BWP (Christie et al. 2013)		d.m. PK, CC, CK
4CYI (Wolf et al. 2014)		c.t. PK, CC, CK
4CYK (Wolf et al. 2014)		c.t. ZF
4CYJ (Wolf et al. 2014)	c.t. Linker	c.t. PK, CC, CK
4D0K (Jonas et al. 2014)	c.t. WD40	c.t. CK
4CZX, (Jonas et al. 2014)	n.c. WD40	n.c. CK
4CZY (Jonas et al. 2014)	n.c. WD40	n.c. PK, CC, CK
4XR7 (Schäfer et al. 2014)	s.c. UCH, RNase	s.c. PK, CC, CK
6R5K (Schäfer et al. 2019)	s.c.	s.c. PK, CC, CK	3 copies of Pab1

Abbreviations: s.c., Saccharomyces cerevisiae; c.t., Chaetomium thermophilum; n.c., Neurospora crassa; UCH, ubiquitin C-terminal hydrolase (Pan2); RNase, DEDD superfamily 3′→5′ exo-RNase domain (Pan2); WD40, WD40 domain (Pan2); CK, C-terminal knob domain (Pan3); CC, coiled coil (Pan3); PK, pseudokinase domain (Pan3); ZF, Zinc-Finger (Pan3).

The C-terminus of the Pan2 protein contains a catalytically active distributive exoribonuclease (RNase) domain (Figure 1), which implies that the active site of the complex dissociates from the substrate between successive deadenylation steps (Lowell et al. 1992). Pan2 activity is enhanced by its cofactor Pan3 that contains a Pab1-interacting motif 2 (PAM-2 motif) (Uchida et al. 2004; Wolf et al. 2014). This motif binds specifically to the PABC domain of Pab1 and thereby ensures the recruitment of the Pan2-Pan3 complex to mRNA substrates with longer poly(A) tails (Siddiqui et al. 2007). In addition, Pan3 interacts with tryptophan-rich stretches in TNRC6/GW182 (Christie et al. 2013; Fabian et al. 2011), a protein that is central in the miRNA-mediated mRNA decay pathway. This exemplifies a strategy that is often exploited in mRNA turnover, where the mRNA degradation machinery is recruited to specific transcripts that have been designated for degradation through different pathways.

Figure 1:

Structure of the Pan2-Pan3 complex. One Pan2 (blue) protein interacts with an asymmetric Pan3 dimer (light and dark green). The 3′ end of the mRNA (orange) interacts with the Pan2 active site (magenta). The upstream bases interact with three Pab1 proteins (light pink, pink and violet). Pan3 contains interaction sites for ATP (orange) and for TNRC6/GW182 (yellow). The figure is based on a superposition of the structures in PBD IDs 6R5K and 4BWP (Table 1).

In isolation, Pan3 forms an almost symmetric homodimer, with a kink in one of the two central coiled-coil domains (Christie et al. 2013). This slight asymmetry favours the association of a single copy of Pan2, mediated by an over 100 amino acid long linker region (Schäfer et al. 2014; Wolf et al. 2014) and the WD40 domain (Jonas et al. 2014) of the Pan2 enzyme (Figure 1). The mRNA substrate is then recognized by the Pan2-Pan3 complex through at least three independent interactions. First, the active site of Pan2 specifically recognizes the unique stacked helical A-form structure that poly(A) RNA adopts (Tang et al. 2019), without forming specific interactions with the adenine bases. Second, the far N-terminal region of Pan3 contains a Zinc-Finger that enhances the interactions with the substrate (Wolf et al. 2014). Finally, the poly(A) substrate is recruited to the Pan2-Pan3 complex via Pab1, as is visualized in the impressive structure of a Pan2-Pan3-Poly(A)-Pab1 complex (Schäfer et al. 2019), where two to three copies of Pab1 exploit one surface to interact specifically with Pan2 and Pan3 and another surface to simultaneously interact with the poly(A) sequence in the substrate. These data also provide a mechanistic explanation for the inefficiency of Pan2-Pan3 in degrading RNA substrates with short poly(A) tails that interact with less than two Pab1 proteins, and for the stimulatory effect exerted by Pab1 on Pan2-Pan3 catalytic activity (Schäfer et al. 2019).

Importantly, the Pan2-Pan3 domain orientations and interactions in multiple independent sub-complexes from different organisms (Table 1) superpose well on the structure of the full complex (Figure 1). This highlights that the structural biology bottom-up approach, where smaller sub-assemblies are studied in isolation, is able to provide relevant structural and mechanistic insights into the full Pan2-Pan3 assembly. Nevertheless, highly dynamic regions often remain unresolved, especially in atomic models obtained from cryo-electron microscopy and X-ray crystallography. For the Pan2-Pan3 complex, over 200 residues of the N-terminal region of Pan3, including the Zinc-Finger and the PAM-2 motif, are invisible. It thus remains unclear how these elements function in the regulation of the deadenylation process. Furthermore, the static images of the active Pan2-Pan3 complex are unable to reveal how the poly(A) tail is translocated towards the active site during deadenylation. The slow nature of the Pan2-Pan3-mediated deadenylation might in that regard be correlated with the very large intermolecular interface between the substrate and the Pan2-Pan3-Pab1 complex, where many interactions need to be broken to shift the substrate towards the active site.

3 mRNA turnover step 2: deadenylation by the Ccr4-Not complex

After Pan2-Pan3 has shortened the poly(A) tail to less than 25–110 nucleotides (Decker and Parker 1993; Yamashita et al. 2005), the second step in deadenylation is performed by the multi-subunit Ccr4-Not complex (Tucker et al. 2001; Yamashita et al. 2005) that contains the catalytically active exonuclease CNOT7/Caf1/Pop2 and the deadenylase CNOT6/Ccr4.

The core of the Ccr4-Not complex is the CNOT1/Not1 protein (267 kDa in humans/240 kDa in yeast), which contains multiple consecutive helical domains connected by linker regions that are likely unstructured and flexible (Figure 2) (Raisch et al. 2019). The Not1 protein acts as a scaffold for interactions with other subunits of the Ccr4-Not complex (Figure 2; Table 2) to assemble a complex that has a molecular weight of around 670 kDa (in humans). From N- to C-terminus, the protein can be roughly divided into four different modules.

Figure 2:

Structural model of the Ccr4-Not complex. The domain organization of the scaffold protein CNOT1 (top), where residue numbers refer to the approximate domain boundaries in the human protein. The four modules form more or less independently structured units that are connected via linker regions. For the NOT1-N module, PDB IDs 8BFI and 8BFJ have been superposed (Table 2), while the TTP complex is based on PDB ID 4J8S. For the nuclease module, (parts of) the PDB IDs 4B8C, 3NGO, 2D5R, 5ANR and 2A1R have been superposed. For the Caf40 module, PDBI Ds 4CRV, 5ONA, 5LSW and 6HOM have been superposed. For the NOT module, PDB IDs 4C0D, 4CQO, 5FU7, 5AJD and 6H3Z have been superposed. The Not1 protein is shown in blue, other Ccr4-Not factors are coloured in different shades of green (with active sites highlighted in magenta), Ccr4-Not-associated proteins are in yellow, and RNA is shown in orange.

Table 2:

Structures of the Ccr4-Not complex.

PDB ID	Not1	Not10	Not11	Caf1	Ccr4	Caf40/Not9	Not2	Not3	Other
8BFI, 8BFJ (Mauxion et al. 2023)	h.s. N-MIF4G N-HEAT	h.s.	h.s.						h.s. GGNBP2
4B8B (Basquin et al. 2012)	c.s. N-HEAT M-HEAT
4J8S (Fabian et al. 2013)	h.s. M-HEAT								h.s. TTP
4B8C (Basquin et al. 2012)	s.c. M-MIF4G			s.c. ND	s.c.
4B8A (Basquin et al. 2012)	c.s. M-MIF4G			s.c. ND
4GMJ (Petit et al. 2012)	h.s. M-MIF4G			h.s. ND
2A1R (Wu et al. 2005)					h.s. PARN				(A)₃
7VOI (Zhang et al. 2022)	h.s. M-MIF4G			hs. ND	h.s. ND
7AX1 (Chen et al. 2021)				h.s. nd	h.s. nd
3NGO (Wang et al. 2010)					h.s. ND				(A)₄ DNA
2D5R (Horiuchi et al. 2009)				h.s. ND					h.s. TOB
4CT4 (Mathys et al. 2014)	h.s. M-MIF4G								h.s. DDX6
5ANR (Ozgur et al. 2015)	h.s. M-MIF4G								h.s. DDX6, Eif4e
4CRW (Chen et al. 2014)	h.s. M-MIF4G								h.s. DDX6
4CRV, 4CRU (Chen et al. 2014)	h.s. CN9BD					h.s.			h.s. TRP
4CT6, 4CT7 (Mathys et al. 2014)	h.s. CN9BD					h.s.			h.s. TRP
4CV5 (Mathys et al. 2014)	s.c. CN9BD					s.c.
5ONA (Sgromo et al. 2018)	h.s. CN9BD					h.s.			d.m. Bag-of-marbles
5LSW (Sgromo et al. 2017)						h.s.			d.m. Roquin
6HOM, 6HON (Keskeny et al. 2019)						h.s.			d.m. Not4
6H3Z (Raisch et al. 2018)	c.t. C-MIF4G
4C0D (Boland et al. 2013)	h.s. C- HEAT						h.s.	h.s.
4BY6 (Bhaskar et al. 2013)	s.c. C-HEAT						s.c.	s.c. Not5
4CQO (Bhandari et al. 2014)	h.s. C-HEAT								h.s. Nanos
5FU6, 5FU7 (Raisch et al. 2016)	h.s. C-HEAT						h.s.	h.s.	D.m. Nanos
5AJD (Bhaskar et al. 2015)	s.c. C-HEAT								s.c. Not4
6TB3 (Buschauer et al. 2020)								s.c. Not5	Ribosome

Abbreviations: h.s., Homo sapiens; s.c., Saccharomyces cerevisiae; c.t., Chaetomium thermophilum; d.m., Drosophila melanogaster; PARN, Poly(A)-specific ribonuclease that is structurally related to Caf1.

First, the NOT1-N module contains an N-terminal Not1 MIF4G domain (middle portion of eIF4G, termed N-MIF4G here), a number of N-terminal helix-turn-helix HEAT (Huntingtin, elongation factor 3, protein phosphatase 2A, and the yeast kinase TOR1) repeats (termed N-HEAT here) and several middle HEAT repeats (termed M-HEAT here). This N-terminal region of Not1 interacts with the mammalian factors CNOT10 and CNOT11 to form an RNA-interacting module, termed the NOT1-N module, that enhances deadenylase activity (Raisch et al. 2019) through an unknown mechanism. The structure of the NOT1-N module (Mauxion et al. 2023) shows an intertwined core, composed of all three proteins, and a flexibly attached so-called antenna that consists of the CNOT11 C-terminal domain. This C-terminal domain of CNOT11 is a binding hub for factors that can be recruited to the Ccr4-Not complex, including the tumour repressor protein GGNBP2 (gametogenetin-binding protein 2) (Mauxion et al. 2023). The M-HEAT repeats that follow the CNOT1-CNOT10-CNOT11 unit have also been structurally characterized and interact with the protein TTP (tristetraprolin) (Fabian et al. 2013), mediating the direct recruitment of the Ccr4-Not complex to mRNAs with destabilizing AU-rich elements (Figure 2). The M-HEAT region of the yeast Not1 protein appears to interact stably with the rest of the NOT1-N module (Basquin et al. 2012). This part of Not1 is not present in the structure of the human NOT-N module and it is thus not clear, whether the stable association of the M-HEAT region with the NOT-N module is conserved.

The second module in the Ccr4-Not complex is the nuclease module, that is built around the CNOT1 middle MIF4G domain (termed M-MIF4G here). This M-MIF4G domain binds to the exoribonuclease CNOT7/Caf1/pop2, which in turn recruits the deadenylase CNOT6/Ccr4. The latter is the only component of the Ccr4-Not complex that does not directly interact with the Not1 scaffold protein (Basquin et al. 2012). Caf1 and Ccr4 are structurally distinct ribonucleases: like Pan2, Caf1 belongs to the DEDDh family of nucleases, whereas Ccr4 belongs to the heterogeneous EEP (exonuclease-endonuclease-phosphatase) family of phosphoesterases. Both nucleases have a strong preference for poly(A) RNA with a 3′ OH adenine residue (Chen et al. 2021), which has been structurally visualized for Ccr4 (Wang et al. 2010). Even though both ribonuclease activities are in principle redundant, both are required to achieve a fully functional deadenylation complex in cells (Piao et al. 2010). Biochemically, it has been observed that the Ccr4-Not activity is efficiently blocked when two or more non-adenine nucleotides are present (Raisch et al. 2019), which prevents non-specific RNA degradation. Single non-adenine bases are tolerated, and mainly removed by Caf1 (Chen et al. 2021). Interestingly, even non-adenine bases up to two nucleotides from the free 3′ end are detected and result in a slowdown of deadenylation (Chen et al. 2021), suggesting a complex mechanism that tunes the mRNA decay speed and thus mRNA stability. The relative orientation of the Not1 M-MIF4G domain and Caf1 appears invariant, as both domains adopt the same orientation in multiple crystal structures (Basquin et al. 2012; Petit et al. 2012; Zhang et al. 2022). The orientation of Caf1 and Ccr4, on the other hand, is not fixed, as these domains have been observed in different relative orientations (Chen et al. 2021; Zhang et al. 2022). This structural plasticity might relate to the way both nuclease active sites act together. It is tempting to speculate that the Ccr4 and Caf1 proteins rotate when e.g. non-adenine bases are encountered in the substrate, given the different degree of inhibition displayed in the presence of distinct non-adenine bases by the two nucleases (Chen et al. 2021) and the loss of the helical structure that a pure poly(A) RNA adopts (Tang et al. 2019). The second function of the M-MIF4G domain in the nuclease module is the recruitment of the DEAD box helicase DDX6/Dhh1 that links the Ccr4-Not complex to mRNA decapping (see below) and translational repression (Chen et al. 2014; Mathys et al. 2014; Ozgur et al. 2015). The M-MIF4G domain of Not1 induces an active conformation in the DDX6 helicase, which is important for efficient miRNA repression (Mathys et al. 2014). A superposition of the available structures reveals that the DDX6 helicase is in close spatial proximity to the nuclease domains (Figure 2). It is, however, still unclear whether DDX6 helicase activity stimulates deadenylation, and if such stimulation would be achieved actively, through the unwinding of secondary structure elements in the mRNA, or passively, by providing an additional RNA binding platform. Additionally, the nuclease module interacts, through Caf1, with the TOB family of antiproliferative proteins (Horiuchi et al. 2009; Hosoda et al. 2011), thereby linking the deadenylation complex to cell cycle regulation and providing a mechanism through which specific RNAs can be recruited to the deadenylase complex.

The Ccr4-Not nuclease module is followed by the Caf40 module that consists of the Not1 CNOT9-binding domain (CN9BD) and the CNOT9/Caf40 (Chen et al. 2014; Mathys et al. 2014). On the one hand, CNOT9 can interact with Trp rich sequences, and thereby bind the TNRC6/GW182 protein, enabling direct recruitment of the Ccr4-Not complex to miRNA targets (Chen et al. 2014; Mathys et al. 2014). On the other hand, Caf40 directly interacts with proteins that contain a Caf40-binding motif (CAB) that is present in e.g. the Drosophila protein Bag-of-marbles (Sgromo et al. 2018), in a number of Roquin proteins (Sgromo et al. 2017), and in metazoan Not4, a conserved E3 ubiquitin ligase (Keskeny et al. 2019). These interactions either target specific mRNAs to the Ccr4-Not complex, or facilitate ubiquitylation of multiple substrates, including ribosome-associated factors.

Finally, the C-terminal Not1 MIF4G-HEAT repeat domains interact with Not2 and Not3 (or the paralogous protein Not5) to form the Not module (Bhaskar et al. 2013; Boland et al. 2013). This module recruits the Ccr4-Not complex to numerous specific mRNA transcripts, including miRNA targets and AU-rich element-containing mRNAs such as the transcription regulator Nanos (Bhandari et al. 2014; Raisch et al. 2016). Even though the structure of the core C-terminal Not module is conserved, the interaction surface of recruited proteins can differ between vertebrates and invertebrates as it is observed for e.g. Nanos. Likewise, yeast Not4 interacts differently with the metazoan and the yeast Ccr4-Not complex, as in the former case interaction is mediated by Caf40 (see above), and in the latter the Not module directly binds Not4 (Bhaskar et al. 2015). Yeast Not5 and its human orthologue CNOT3 can interact directly with stalling ribosomes and thereby target transcripts with non-optimal codon usage for degradation (Absmeier et al. 2022; Buschauer et al. 2020). The exoribonuclease Xrn1 (see below) has also been shown to interact with the Not module (Chang et al. 2019), though the structural details of the interaction are still unknown. This interaction inhibits the Caf1-mediated deadenylation activity of the Ccr4-Not complex (Chang et al. 2019) and thus reveals a mechanism that regulates the interplay of deadenylation and 5′→3′ transcript degradation. Finally, the Not module specifically interacts with poly(U) RNA in a structurally undetermined manner (Bhaskar et al. 2013).

Despite the abundant structural information on the isolated modular components of the Ccr4-Not complex (Figure 2), it remains unclear how these building blocks are assembled in three dimensional space. A low resolution electron microscopy map indicates that the modules are arranged into an L-shaped complex (Nasertorabi et al. 2011) where the relative positioning of the arms varies between reconstructions, suggesting flexibility. It is thus plausible that the full complex is highly dynamic in solution, which would explain the observed cross-talk between different modules and which could provide additional means of regulating activity and/or substrate recruitment. This flexibility, however, results in challenges in the determination of higher resolution structures of the complete assembly (Raisch et al. 2019). In that light, it is worth mentioning that it is not possible to simply superpose the structures of the individual Not modules onto AlphaFold-derived models (Jumper et al. 2021) of the full length Not1 protein from either yeast or humans, as this results in severe structural clashes between the modules. This computational exercise underscores the importance of experimental methods to obtain information on highly complex assemblies as relative domain orientations are not well-predictable.

4 mRNA turnover step 3

After 3′ deadenylation, the mRNP (messenger ribonucleoprotein) complex undergoes a transition from a translationally competent state to one that is primed for rapid and irreversible transcript degradation (Tharun and Parker 2001). This mRNA decay process can occur via either of two pathways (Garneau et al. 2007). One involves 5′ mRNA decapping by the Dcp2 enzyme, followed by the processive 5′→3′ exoribonucleolytic hydrolysis of the mRNA body by Xrn1. Alternatively, the transcript can be degraded in the 3′→5′ direction by the cytoplasmic RNA exosome complex, followed by decapping of the residual small mRNA fragment by the scavenger decapping enzyme DcpS. Mechanistically, these pathways can have a different functional consequence, as the decapping of a transcript results in an immediate inhibition of translation initiation and thus prevents the production of truncated proteins that could still occur in the 3′→5′ degradation pathway. It is still unknown how an mRNA is committed to either of these decay pathways. Nevertheless, it is plausible to postulate that this step, like all other steps in mRNA turnover, is tightly regulated.

4.1 mRNA turnover step 3a: Xrn1-mediated degradation of the mRNA body

The 5′→3′ mRNA degradation pathway appears to be the most prominent one in yeast (Muhlrad et al. 1995). Over the years, a large number of structures of the components involved in this pathway have been elucidated (Table 3). Nevertheless, the exact molecular and structural details that result in the recruitment of the 5′ decapping machinery to the 3′ deadenylated mRNA remain unclear. Based on the knowledge gathered from sub-complexes, it is possible to obtain insights into the structural links between deadenylation, decapping and exonucleolytic degradation of the mRNA body.

Table 3:

Structures of proteins involved in 5′→3′ mRNA decay.

PDB ID	Xrn1	Dcp1	Dcp2	Edc1	Edc3	Pat1	Dhh1	Lsm1-7	Other	RNA
2Y35 (Jinek et al. 2011)	d.m.									DNA
3PIE, 3PIF (Chang et al. 2011)	k.l. E178Q
6Q8Y (Tesina et al. 2019)	s.c.								Ribosome	mRNA
2LYD (Braun et al. 2012)	d.m. IDR	d.m.
1Q67 (She et al. 2004)		s.c.
2WX3 (Tritschler et al. 2009a)		h.s. CTD
4B6H (Lai et al. 2012)		h.s.							PNRC2
2WX4 (Tritschler 2009a)		d.m.
5MP0, 5QOH-5QOZ, 5QP0-5QP9, 5QPA-5QPC			h.s. CD						Inhibitors
2A6T (She et al. 2006)			s.p. RD, CD
2JVB (Deshmukh et al. 2008)			s.c. CD
4K6E, 4KG3, 4KG4 (Aglietti et al. 2013)			s.c. ND
2QKL, 2QKM (She et al. 2008)		s.p.	s.p. RD							+/− ATP
5LON (Charenton et al. 2016)		k.l.	k.l. RD, CD
5J3Y (Valkov et al. 2016)		s.p.	s.p. RD, CD
5KQ1 (Mugridge et al. 2016)		s.p.	s.p. RD, CD						h.s. PNRC2
5KQ4 (Mugridge et al. 2016)		s.p.	s.p. RD, CD						h.s. PNRC2	Cap analogue
5J3Q (Valkov et al. 2016)		s.p.		s.p.
5JP4 (Wurm et al. 2016)		s.p.		s.p.
5N2V (Wurm et al. 2017)		s.p.	s.p. RD, CD	s.p.						m⁷Gpp
5J3T (Valkov et al. 2016)		s.p.	s.p. RD, CD	s.p.
6AM0 (Mugridge et al. 2018)		k.l.	k.l. RD, CD	k.l.	k.l. LSm					Cap analogue
5LOP (Charenton et al. 2016)		k.l.	k.l. RD, CD		k.l. LSm					m⁷GDP
4A54 (Fromm et al. 2012)			s.p. HLM		s.p. LSm
5LM5, 5LMF, 5LMG (Charenton et al. 2017)			s.c. HLMs			s.c. CTD
6Y3Z (Charenton et al. 2020)		s.c.	s.c.		s.c. LSm				s.c. Pby1
6Y3P (Charenton et al. 2020)									k.l. Pby1
2RM4 (Tritschler et al. 2007)					d.m. LSm
2VC8 (Tritschler et al. 2007)					h.s. LSm
3D3J, 3D3K (Ling et al. 2008)					h.s. YjeF-N
4A53 (Fromm et al. 2012)					s.p. Lsm
4OGP, 4OJJ (Fourati et al. 2014)						s.c. CTD
2XEQ, 2XER, 2XES (Braun et al. 2010)						h.s. CTD
1S2M (Cheng et al. 2005)							s.c.
2WAX, 2WAY (Tritschler et al. 2009a)					h.s. FDF		h.s. RecA2
4BRU (Sharif et al. 2013)					s.c. FDF		s.c. RecA2
4BRW (Sharif et al. 2013)						s.c. N-IDR	s.c. RecA2
6S8S (Peter et al. 2019)					h.s. FDF		h.s. RecA2
6F9S (Brandmann et al. 2018)							h.s. RecA2		c.e. Lsm14
5ANR (Ozgur et al. 2015)							h.s. RecA2		h.s. CNOT1
2VXG (Jinek et al. 2008)									d.m. Ge-1
4Q2S (Fromm et al. 2014)									s.p. Pdc1
4N0A (Wu et al. 2014)						s.c. CTD		s.c. Lsm2,3
4C8Q (Sharif and Conti 2013)						s.c. CTD		s.c. Lsm1-7
4C92 (Sharif and Conti 2013)								s.c. Lsm1-7
4EMG (Wu et al. 2012)								s.p. Lsm3
3BW1 (Naidoo et al. 2008)								s.c. Lsm3
4EMK (Wu et al. 2012)								s.p. Lsm5,6,7
3SWN (Mund et al. 2011)								s.p. Lsm5,6,7
4M75 (Zhou et al. 2014)								s.c. Lsm1-7
6PPQ, 6PPV (Montemayor et al. 2020)								s.p. Lsm1-7		RNA
4EMH (Wu et al. 2012)								s.p. Lsm4

Abbreviations: h.s., Homo sapiens; s.p., Schizosaccharomyces pombe; s.c., Saccharomyces cerevisiae; d.m., Drosophila melanogaster; k.l., Kluyveromyces lactis; c.e., Caenorhabditis elegans; CTD, C-terminal domain (Dcp1, Pat1); RD, regulatory domain (Dcp2); CD, catalytic domain (Dcp2); HLM, helical leucine rich motif (Dcp2); Lsm, Lsm domain (Edc3); RecA2, second RecA domain (DDX6/Dhh1); N-IDR, N-terminal IDR (Pat1).

Table 4:

Structures of proteins involved in 3′→5′ mRNA decay.

PDB ID	DcpS	Exo9	Rrp44/Dis3	Other	RNA/cap analogue
1VLR (Han et al. 2005)	m.m.
6GBS (Fuchs et al. 2020)	c.t
3BL7, 3BL9, 3BLA (Singh et al. 2008)	h.s.				Inhibitor
1ST0, 1ST4 (Gu et al. 2004)	h.s.				m⁷GpppG, m⁷GpppA
1XMM (Chen et al. 2005)	h.s.				m⁷GDP
1XML (Chen et al. 2005)	h.s.
4QDE, 4QEB, 4QDV (Hett et al. 2015)	h.s.				Inhibitor
6TRQ (Fuchs et al. 2020)	s.c.				m⁷GpppGU
5OSY (Wojtczak et al. 2018)	h.s.				m⁷G(5′S)ppSp(5′S)G
5BV3 (Neu et al. 2015)	s.c.				m⁷GDP
7TUV (Cesaro et al. 2023)			t.b. CSDs-RNB-S1, open		RNA: GGUU
6MD3 (Cesaro et al. 2019)			t.b. PIN domain
2VNU (Lorentzen et al. 2008)			s.c. CSDs-RNB-S1, open		RNA: (A)₁₀
4RO1 (Lv et al. 2015)			s.p. DIS3-like 2
4PMW (Faehnle et al. 2014)			m.m. Dis3-like 2, open		RNA: (U)₁₄
2WP8 (Bonneau et al. 2009)		s.c Rrp41-Rrp45	s.c.
2NN6 (Liu et al. 2006)		h.s.
4OO1 (Wasmuth et al. 2014)		s.c.		s.c. Rrp6	RNA: (A)₂₄
5OKZ (Falk et al. 2017)		s.c.		s.c. Mpp6
6H25 (Gerlach et al. 2018)		h.s.	h.s. Open	h.s. Mpp6	RNA: (U)₄₄
5JEA (Kowalinski et al. 2016)		s.c.	s.c. Closed	s.c. Ski7	RNA: 46 nt
5G06 (Liu et al. 2016)		s.c.	s.c. Open	s.c. Ski7
5K36 (Zinder et al. 2016)		s.c.	s.c. Open	s.c. Rrp6	RNA: 17 nt
4IFD (Makino et al. 2013)		s.c.	s.c. Closed	s.c. Rrp6	RNA: 45 nt
5C0X (Makino et al. 2015)		s.c.	s.c. Closed	s.c. Rrp6	RNA: 45 nt (including a hairpin)
5VZJ (Wasmuth et al. 2017)		s.c.	s.c. Open	s.c. Rrp6, Mpp6	RNA: 11 nt RNA: 19 nt
5C0W (Makino et al. 2015)		s.c.	s.c. Open	s.c. Rrp6, Rrp47	RNA: 18 nt
6D6Q, 6D6R (Weick et al. 2018)		h.s.	h.s. Open	h.s. Rrp6, Mpp6, Mtr4	RNA: 62 nt
6FSZ (Schuller et al. 2018)		s.c.	s.c. Closed	s.c. Rrp6, Rrp47, Mtr4, Mpp6	RNA: 23 nt
6LQS (Du et al. 2020)		s.c.	s.c. Closed	s.c. Rrp6, Rrp47, Mtr4, Mpp6	RRNA and pre-90S ribosome
6FT6 (Schuller et al. 2018)				s.c. Rrp6, Rrp47, Mtr4, Mpp6	RRNA and pre-60S ribosome
4BUJ (Halbach et al. 2013)				s.c. Ski2, Ski3, Ski8
7QDR, 7QDS (Kögel et al. 2022)				h.s. Ski2, Ski3, Ski8
7QDY, 7QDZ (Kögel et al. 2022)				h.s. Ski2, Ski3, Ski8	RNA: (U)₂₅ or (U)₆
5MC6 (Schmidt et al. 2016)				s.c. Ski2, Ski3, Ski8	mRNA and ribosome
7QE0 (Kögel et al. 2022)				h.s. Ski2	RNA: (U)₉

Abbreviations: m.m., Mus musculus; c.t., Chaetomium thermophilum; h.s., Homo sapiens; t.b., Trypanosoma brucei; nt, nucleotides; CSD, Cold Shock Domain (Rrp44); RNB, catalytic domain of ribonuclease (Rrp44); S1, S1 domain (Rrp44). Note that Rrp6, Rrp47 (Exosome complex protein LRP1), Mtr4 and Mpp6 are nuclear proteins and exosome complexes that contain those subunits are thus not directly involved in canonical mRNA turnover. These complexes are included in this Table to illustrate the structural plasticity of Rrp44.

The 3′ end of the transcript interacts with the pore of the ∼90 kDa heteroheptameric doughnut-shaped Lsm1-7 complex (Montemayor et al. 2020) (Figure 3A, red/black). The Lsm2 and Lsm3 subunits of this complex stably associate with the C-terminal helical region (Pat1-C) of the ∼90 kDa multi-domain scaffolding protein PATL1/Pat1 (Sharif and Conti 2013; Wu et al. 2014) (Figure 3A). Pat1 binding enhances the affinity and specificity of the Lsm1-7 ring for oligoA RNA, as present in a deadenylated 3′ poly(A) mRNA. This interaction is mediated by the structurally uncharacterized middle domain of Pat1 (Pat1-M) (Lobel et al. 2019), the C-terminal domain of Pat1 (Pat1-C), and the C-terminal extension of Lsm1 (Chowdhury et al. 2007; Lobel and Gross 2020). It would be exciting to determine the structural details of this Lsm-Pat1-mRNA complex, and thus to understand the molecular mechanisms that result in the specific recognition of deadenylated transcripts. At the same time, the Pat-C region interacts with one of the multiple helical-rich motifs in the C-terminal IDR (Intrinsically Disordered Region) of the Dcp2 mRNA decapping enzyme (Charenton et al. 2017) (Figure 3A, green). Note that this C-terminal IDR is not present in human Dcp2 protein, where a long C-terminal IDR is instead present in the Dcp2-interacting protein Dcp1. Functionally, Pat1-C thus links recognition of the deadenylated 3′ end of a transcript (by Lsm1-7) to the hydrolysis of the 5′ mRNA cap structure (by Dcp2).

Figure 3:

Structural link between 3′ deadenylation and 5′ decapping and degradation. (A) Superposition of known three-dimensional structures involved in 5′→3′ mRNA decay (Table 3). The Lsm1-7 complex (red) interacts with the 3′ end of the mRNA (black, PDB ID 6PPV) as well as with Pat1 (orange, PDB ID 4C8Q). Pat1 connects to Dcp2 (green, PDB ID 5LM5) via a number of linear motifs in its C-terminal IDR (only one of the interactions is shown). These motifs also interact with the Lsm domain in Edc3 (blue, PDB ID 6AM0). Edc3 dimerizes via a C-terminal Yjef_N domain (PDB ID 3D3J) and interacts through its central IDR with the DEAD box helicase DDX6/Dhh1 (purple, PDB ID 6S8S). This helicase also links the mRNA to the deadenylation machinery via Not1 (light blue, PDB ID 5ANR). The mRNA decapping enzyme Dcp2 interacts with the decapping activators Dcp1 (yellow) and Edc1 (pink; PDB ID 6AM0). Dcp1 interacts with the C-terminal IDR of Xrn1 (light brown, PDB ID 2LYD and 2Y35). (B) All known structures of Dcp2, superposed on the Dcp2 regulatory domain and Dcp1 (when present). The Dcp2 catalytic domain (light green) and the catalytic Nudix helix (red) can adopt a large number of orientations with respect to the regulatory domain. In solution, an open, closed and active orientation have been observed (PDB ID 2QKL, 5N2V). (C) Static structures of Xrn1 (PDB ID 2Y35, 6Q8Y), as well as NMR data on Xrn2, reveal motions in the N-terminal helix (red) that correlate with substrate turnover. The stacking interactions between the three most 5′ bases of the substrate RNA (orange) and the conserved His and Trp residues in Xrn1 (blue) are shown.

The 5′→3′ decay interaction network is significantly more extended as described above and involves multiple additional factors. The C-terminal IDR of Dcp2 (or Dcp1 in humans) can simultaneously recruit multiple Edc3 proteins via an N-terminal Lsm domain (Fromm et al. 2012) (Figure 3A, blue). These Edc3 proteins in turn dimerize through a C-terminal Yjef_N domain and exploit an internal IDR to interact with the second RecA domain of the helicase DDX6/Dhh1 (Sharif et al. 2013; Tritschler et al. 2009a) (Figure 3A, purple). Thereby, this latter interaction directly couples decapping (Figure 3) and deadenylation (Figure 2), as DDX6 can simultaneously interact with Edc3 and the Ccr4-Not scaffold protein Not1 (Mathys et al. 2014) (Figure 3A, light blue). However, it remains unclear how the Ccr4-Not complex dissociates from the mRNA to allow for the recruitment of the Lsm1-7 ring, and what the trigger for this exchange is.

The Dcp2 decapping enzyme (Figure 3A, green) contains a catalytic domain with a Nudix helix as well as an N-terminal regulatory domain. This latter domain forms part of the composite substrate binding site (Floor et al. 2010) and interacts tightly with the prime activator of decapping Dcp1 (She et al. 2008) (Figure 3A, yellow). The role of Dcp1 is threefold: first, it stabilizes the N-terminal domain of Dcp2 and thereby facilitates mRNA decapping (Wurm et al. 2017); second, it recruits the Edc1 protein (Figure 3A, pink) that stabilizes the Dcp2 enzyme in an active conformation (Charenton et al. 2016; Mugridge et al. 2018; Wurm et al. 2017); and finally, it provides a dynamic binding groove for proline rich sequences in mRNA decapping factors (Wurm et al. 2016) and Xrn1 (Braun et al. 2012) (Figure 3A, light brown). This latter interaction places the 5′→3′ exoribonuclease close to the decapped transcript, favouring a rapid and processive degradation of the mRNA body after decapping (Chang et al. 2011; Jinek et al. 2011). Xrn1-mediated degradation can also occur on transcripts that are still being actively transcribed, as is captured in a structure of Xrn1 in complex with the ribosome (Hu et al. 2009; Tesina et al. 2019). Structurally, Xrn1 contains a catalytic core in which the active site pocket is selective for 5′ mono-phosphorylated RNA and thereby strongly discriminates against capped RNA. This Xrn1 core is succeeded by four folded domains whose functions are not well understood. These additional domains are resolved and visible only in some of the determined Xrn1 structures (Chang et al. 2011; Jinek et al. 2011; Tesina et al. 2019), indicating that they can adopt different orientations with respect to the core domain.

The “model” of the 5′→3′ mRNA decay pathway depicted in Figure 3 should not be considered the only possible link between deadenylation and 5′→3′ mRNA degradation. It is important to realize that, similar to the Ccr4-Not deadenylation complex, the domain organization and the interaction details between the different 5′→3′ mRNA decay factors can vary between species. Differences include for example the direct and stable interaction of the Dcp1 and Dcp2 proteins in yeast, whereas in humans the additional protein Edc4 is required to form a functional decapping complex (Jinek et al. 2008). In addition, the yeast Dcp1 protein comprises only one modular domain, while this decapping factor trimerizes in humans via a long C-terminal extension (Tritschler et al. 2009b). Furthermore, the C-terminal IDR of mammalian Xrn1 has been shown to interact not only with Dcp1, but also directly with the Edc4 protein and with Pat1 (Chang et al. 2019). Future structural studies will be needed to reveal how these differences result in species, tissue or cellular conditional differences in the architecture and regulation of the mRNA degradation pathways.

The catalytic domain of Dcp2 possesses only a low intrinsic decapping efficiency that is enhanced by the Dcp2 regulatory domain and regulated by the Dcp2 IDR (He and Jacobson 2015; Paquette et al. 2018). In addition, Dcp2 decapping activity is increased by the interaction with various decapping factors, including Dcp1, Edc1, Edc3, Pat1 and Dhh1 (Nissan et al. 2010; Wurm et al. 2017). It has been experimentally shown that the Dcp1-Dcp2 complex is highly dynamic and that the catalytic domain can adopt different orientations with respect to the rigid conformation that is adopted in the complex of Dcp1 with the Dcp2 regulatory domain (Charenton et al. 2016; Mugridge et al. 2018; She et al. 2006, 2008; Wurm et al. 2017) (Figure 3B). So far, structural studies have described seven different Dcp2 conformations, of which only three have been directly observed in solution (Wurm et al. 2017): an open state that has a high affinity for the body of the RNA substrate, a closed state that is catalytically impaired, and an active state in which mRNA decapping can take place (Figure 3B). The motions in the Dcp2 enzyme have been quantified in detail using high resolution nuclear magnetic resonance (NMR) methods that thereby complement static structural information (Krempl et al. 2022; Wurm et al. 2017). The binding of the decapping activator Edc1 stabilizes the active conformation of Dcp2 and thereby enhances decapping activity directly through the modulation of protein motions (Charenton et al. 2016; Wurm et al. 2017). The regulation of enzyme activity through the modulation of the conformational space that a protein samples is likely a more general mechanism and could also be important for other complexes involved in mRNA turnover. If and how the Dcp2 enzyme is differentially active on cap0, cap1 or other cap structures has not been addressed in detail as far as we are aware, but modifications on or close to the cap structure could influence decapping and thereby provide a means to influence mRNA stability.

The proteins involved in 5′→3′ mRNA decay can condensate into cellular processing bodies, or P-bodies (Sheth and Parker 2003), through a large dynamic network of redundant and weak intermolecular interactions (Li et al. 2012) that involve RNA, folded protein domains as well as disordered protein regions (Fromm et al. 2014; Luo et al. 2018; Schütz et al. 2017). The exact functional role of this self-assembly process remains a matter of debate and it is not clear if processing bodies are sites where mRNA is stably stored, or where active mRNA turnover takes place. It is tempting to speculate that the Dcp2 domain orientation (Figure 3B) is modulated upon condensation such that decapping can be inhibited or enhanced (Damman et al. 2019; Schütz et al. 2017; Tibble et al. 2021).

The importance of structural changes in enzyme function is further illustrated by the Xrn1 enzyme (Figure 3C). The three most 5′ bases of the decapped mRNA stack tightly between a pair of conserved histidine and tryptophan residues to prevent the premature release of the substrate from the enzyme. At the same time, the substrate needs to translocate one base towards the active site after each cleavage event. This substrate movement appears to go in concert with motions in the N-terminal Xrn1 helix (Figure 3C), as structurally different Xrn1 conformations have been observed in static structures of Xrn1 that differ in the translocation step of the substrate (Jinek et al. 2011; Tesina et al. 2019). Experimentally, the link between these motions and enzyme turnover rates has been characterized in solution for the closely related Xrn2 enzyme (Overbeck et al. 2022).

4.2 mRNA turnover step 3b: exosome-mediated degradation of the mRNA body

In mammals, mRNAs appear to be primarily degraded in the 3′→5′ direction (Wang and Kiledjian 2001). In this decay pathway the deadenylated transcripts recruit the cytoplasmic exosome complex that processively degrades the mRNA body.

The central unit of the exosome complex is the catalytically inactive 270 kDa Exo9 core. Exo9 contains a hexameric ring structure that is composed of the heterodimeric RNase PH domain proteins Rrp41-Rrp45, Rrp46-Rrp43 and Mtr3-Rrp42 and that is capped with the RNA binding proteins Csl4, Rrp4 and Rrp40 (Figure 4A) (Cvetkovic et al. 2017). The 3′ end of a single stranded RNA substrate can enter the Exo9 barrel via a narrow pore that is formed by the cap proteins (Liu et al. 2006). The Dis3/Rrp44 protein adds catalytic activity to the exosome (Dziembowski et al. 2007) and interacts with the Exo9 barrel on the site opposite the cap to form the Exo10 complex (Figure 4A). This Exo10 complex is found in the nucleus, where it plays a role in RNA processing and decay, as well as in the cytoplasm, where it is involved in mRNA turnover (Januszyk and Lima 2014).

Figure 4:

Structures of complexes involved in 3′→5′ mRNA degradation. (A) Structure of the exosome complex (PDB ID 4IFD; Table 4). The central hexameric barrel (dark and light blue) interacts with three cap proteins (teal) that form a narrow entrance channel for the substrate RNA (orange). This Exo9 complex interacts with the catalytic subunit Rrp44 (green) to form Exo10. (B) The Rrp44 enzyme can adopt different conformations that result in different RNA binding grooves. The individual protein domains have been coloured in different shades of green. The exo- and endonuclease active sites are indicated in pink. The image is based on PDB ID 4IFD (top, closed conformation) and 5C0W (bottom, open conformation). (C) The Ski2-3-8 complex can form a closed state (top, PDB ID 7QDY), in which the substrate RNA-binding channel is obstructed by Ski3. The dissociation of the Ski3-8 complex from the Ski2 enzyme opens the RNA channel and allows for the interaction of the Ski complex with Exo10 (PDB ID 7QE0 and 7QDS). (D) Model of how the exosome complex can be recruited to the ribosome in the cytoplasm. The model was constructed by superposing structures of the Ski2-ribosome complex (PDB ID 5MC6), the Ski2 protein in complex with RNA (PDB ID 7QE0) and the Exo10-Ski7 complex (PDB ID 5JEA) onto the structure of the nuclear exosome (PDB ID 6FSZ), structurally aligning Ski2 with Mtr4. (E) Structure of the human DcpS enzyme in the absence (top, PDB ID 1XML) and presence (bottom, PDB ID 1XMM) of substrate. The two DcpS protomers are coloured in green and blue. The catalytic H-x-H-x-H triad is highlighted in pink. The mRNA fragment is shown in orange.

The ∼110 kDa Rrp44 enzyme contains two active sites (Figure 4B), one endonuclease site that is located in the N-terminal PIN (Pilus-forming N-terminus) domain (Lebreton et al. 2008), and one exonuclease site that is found in the C-terminal RNB domain. Both active sites have been suggested to cooperate, but how this would mechanistically work remains unclear. The exonuclease site hydrolyses the substrate base-by-base in a highly processive manner (Lorentzen et al. 2008) and details of the enzyme:RNA interaction have been observed in multiple structures of the complex in the presence of substrate. Interestingly, the Rrp44 enzyme has been observed in two structurally different conformations that differ in domain orientations and RNA binding path (Figure 4B) (Makino et al. 2015). It remains to be determined if and how these large structural changes are linked with the processing activity of the Exo10 complex or with special features in the substrate.

The Exo10 complex interacts with additional factors in a manner that depends on the cellular localization (Januszyk and Lima 2014). In the nucleus, the Exo10 complex associates with the ribonuclease Rrp6, the RNA helicase Mtr4 and the co-factors Rrp47 and Mpp6 (Butler and Mitchell 2011). In the cytoplasm, Exo10 interacts with the adaptor protein Ski7 (Kowalinski et al. 2016; Liu et al. 2016), that can subsequently recruit the SKI (Superkiller) complex, which consists of the RNA helicase Ski2 (structurally highly similar to Mtr4), the tetratricopeptide repeat protein Ski3 and two copies of the WD40 repeat protein Ski8. The recruitment of the SKI complex is essential for exosome-mediated 3′→5′ mRNA decay (Anderson and Parker 1998).

The Ski2-7-8 module has been found in a conformation in which the Ski3-8 proteins block the exit channel of the Ski2 helicase domain, and thereby prevent translocation of the mRNA (Halbach et al. 2013; Kögel et al. 2022) (Figure 4C). In an ATP-dependent manner, the Ski3-8 module can “swing out”, which has two functional consequences. First, the RNA channel in Ski2 opens such that mRNA translocation is no longer obstructed, and second, the helicase domain of Ski2 is then able to dock directly onto the Exo10 complex. The interaction between the Exo10 complex and the Ski-2 helicase domain has not been structurally characterized. However, based on biochemical data and the structure of the nuclear exosome in complex with Mtr4, it is possible to construct a model of the Exo10-Ski7-Ski2 complex (Figure 4D), in which Rrp47 bridges the interaction between the Exo10 complex and Ski2.

The Exo10-SKI complex also plays a role in mRNA surveillance pathways, where faulty mRNA transcripts are endonucleolytically cleaved upon ribosome stalling (Shoemaker and Green 2012). The resulting free 3′ end of the cleaved RNA can then be directly channelled into the exosome via the Ski2 protein (Kögel et al. 2022; Schmidt et al. 2016). Multiple parts of this process have been visualized and can be combined into a structural model (Figure 4D) that shows how the Ski2 protein first interacts with an arch region close to the mRNA exit pore in the ribosome, how afterwards the 3′ end of the mRNA is threaded through the helicase domain of Ski2, and how it then reaches the active site of Rrp44 via the Exo9 complex. At the same time, it remains unclear how the Exo10-SKI complex is targeted to a deadenylated mRNA during canonical mRNA turnover, as no physical link between the deadenylase machinery and the exosome has been identified so far.

The products of the exosomal degradation process are single nucleotides and a small capped RNA fragment of 2–5 nucleotides (Fuchs et al. 2020; Mitchell et al. 1997). These short 5′ cap mRNA fragments are subsequently decapped by the ∼80 kDa scavenger decapping enzyme DcpS/Dcs1p, that releases m⁷GMP and the 5′ diphosphorylated mRNA fragment (Liu et al. 2002; Nuss and Furuichi 1977; Wang and Kiledjian 2001).

In the apo state, DcpS forms a symmetric homodimer with two active sites, each between the N- and C-terminal lobes (Figure 4E) (Chen et al. 2005). The enzyme undergoes a large structural change upon substrate recruitment, which results in the formation of one catalytically competent active site (Gu et al. 2004). At the same time, the other substrate-binding site opens into an inactive conformation that allows for the concomitant recruitment of a second substrate. Upon hydrolysis of the first substrate, the N-terminal lid domain flips over to form a catalytically competent active site around the second substrate, and consequently releases the products from the first active site. These flipping motions become fast in the presence of a large excess of substrate, thereby inhibiting substrate turnover (Krempl and Sprangers 2023; Neu et al. 2015). Importantly, DcpS is only active on very short mRNA fragments as longer mRNAs prevent the formation of the closed active site due to steric clashes between the enzyme and the third base in the substrate (Fuchs et al. 2020). Functionally, this elegant mechanism prevents the decapping of long mRNAs that might still be actively involved in translation.

An interaction between DcpS and the exosome has been reported (Wang and Kiledjian 2001), although we have not (yet) been able to observe a direct interaction between these complexes in vitro using purified components (unpublished data). It thus remains unclear how the exosome products are transferred to DcpS.

5 Variations in the mRNA decay pathways

The steps described above form a basic pathway to facilitate an ordered and regulated turnover of mRNA. Numerous (cell- and organism-dependent) variations to these pathways exist that have not been structurally studied in detail so far. As an example, after deadenylation, or after cleavage of the poly(A) tail, terminal uridylyltransferase enzymes can add 3′ uracil stretches to short poly(A) tails that no longer interact with Pab1 (Lim et al. 2014). This uridylation results in enhanced mRNA decay rates either through the recruitment of the Lsm1-7 complex and subsequent Dcp2 mediated mRNA decapping (Rissland and Norbury 2009; Song and Kiledjian 2007), or through the action of the Dis3L2 enzyme (Malecki et al. 2013), a paralogue of the Rrp44 enzyme that is unable to associate with the exosome. It remains to be determined under which conditions, and for which substrates, uridylation takes place, and which benefits this may provide.

Likewise, numerous other triggers can lead to the rapid turnover of an mRNA transcript. As described above, the miRNA silencing machinery can directly recruit the Ccr4-Not complex through the TNRC6/GW182 protein or through AU-rich element-binding proteins like TTP. Additional pathways include cellular quality control or surveillance mechanisms that recognize faulty transcripts (Graille and Séraphin 2012), such as those containing a premature translation termination codon (PTC), which activates the nonsense-mediated decay (NMD) pathway (Kervestin and Jacobson 2012) via the translating ribosome. Central to this pathway is the Upf1 (up-frameshift 1) protein that serves at least three functions. First, it interacts with the Dcp2 C-terminal IDR to initiate mRNA decapping; second, it recruits the endoribonuclease SMG6 that cleaves the transcript in the vicinity of the PTC; and, finally, it interacts with the SMG5-SMG7 module, which in turn recruits the Ccr4-Not complex to initiate deadenylation. Similarly, the no-go decay (NGD) pathway (Harigaya and Parker 2010) is activated when ribosomes stall or collide on a transcript, e.g. due to stable secondary structures in the RNA. This can also result in the endoribonucleolytic cleavage of the transcript, followed by the recruitment of the 5′→3′ decay machinery. Analogous to the NGD pathway, the non-stop decay (NSD) pathway (Klauer and van Hoof 2012) is activated when a stop codon is lacking, and translating ribosomes slow down while translating the poly(A) tail (Chandrasekaran et al. 2019; Tesina et al. 2020). In all these cases the same enzymes that play a role in canonical mRNA turnover are recruited via interaction networks that are structurally only partially determined.

6 Concluding remarks

In the past decades our knowledge regarding the structures of the proteins involved in mRNA turnover has increased significantly. Based on that, a picture arises that reveals how these enzymes are embedded in large interaction networks. In the future, it will be important to understand how the activity of these enzymes is regulated on a structural level, which likely involves structural changes and protein dynamics, as was shown for e.g. Dcp2 and the Xrn1 homologue Xrn2. Based on recent technological advances in structural biology methods, including computational methods (Jumper et al. 2021), single-particle cryo-electron microscopy (Guaita et al. 2022) and NMR spectroscopy (Schütz and Sprangers 2019), we are looking forward to exciting future findings that should ultimately allow us to fully describe how mRNA decay is regulated on an atomic level.

Corresponding author: Remco Sprangers, Institute of Biophysics and Physical Biochemistry, Regensburg Center for Biochemistry, University of Regensburg, D-93053 Regensburg, Germany, E-mail: remco.sprangers@ur.de

Funding source: Deutsche Forschungsgemeinschaft

Award Identifier / Grant number: SFB 960 TP/B12

Acknowledgments

We would like to thank all lab members for discussions. The authors thank Jan Overbeck, Julian Hübner, David Stelzig and Martin Brehm for constructive comments on the manuscript. The authors would also like to thank Herbert Tschochner for his work as the SFB 960 speaker during the past 12 years.

Research ethics: Not applicable.
Author contributions: All the authors have accepted responsibility for the entire content of this submitted manuscript and approved submission.
Competing interests: The authors declares no conflicts of interest regarding this article.
Research funding: This work was supported by the Deutsche Forschungsgemeinschaft under SFB 960 TP/B12.
Data availability: Not applicable.

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Received: 2023-04-13

Accepted: 2023-08-24

Published Online: 2023-09-15

Published in Print: 2023-10-26

This work is licensed under the Creative Commons Attribution 4.0 International License.

A structural biology view on the enzymes involved in eukaryotic mRNA turnover

Abstract

1 Introduction

2 mRNA turnover step 1: deadenylation by Pan2-Pan3

3 mRNA turnover step 2: deadenylation by the Ccr4-Not complex

4 mRNA turnover step 3

4.1 mRNA turnover step 3a: Xrn1-mediated degradation of the mRNA body

4.2 mRNA turnover step 3b: exosome-mediated degradation of the mRNA body

5 Variations in the mRNA decay pathways

6 Concluding remarks

Acknowledgments

References

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