Biochimica et Biophysica Acta (BBA) - Gene Regulatory Mechanisms
ReviewXRN 5′ → 3′ exoribonucleases: Structure, mechanisms and functions☆
Introduction
Messenger RNA turnover is a critical modulator of gene expression. Transcripts are constantly exposed to an array of proteins, small RNAs, and turnover mechanisms primarily devoted to regulating their stability. The implementation of these mechanisms also results in the elimination of defective mRNAs. Accordingly, mutations that cause defects in mRNA turnover can have significant consequences.
In the cytoplasm, most eukaryotic mRNAs are degraded by the 5′ → 3′ exoribonuclease, XRN1 (PACMAN), and/or by the exosome complex, which has both endoribonucleolytic and 3′ → 5′ exoribonucleolytic activities [1], [2], [3]. In the nucleus, mRNA precursors are degraded by the XRN1 paralog XRN2 (RAT1), or the nuclear exosome complex [2], [4], [5], [6]. Additionally, XRNs also participate in diverse aspects of RNA metabolism such as RNA silencing, rRNA maturation and transcription termination [4], [6]. Plants lack an XRN1 ortholog but have an ortholog of XRN2, called XRN4, which resides in the cytoplasm and functions like XRN1 [4], [7]. The XRNs are extremely important at the organismal level as well; loss of nuclear XRN function is lethal in diverse systems and the loss of XRN1 or XRN4 can cause a range of defects such as those affecting growth, development, and responses to hormonal and environmental stimuli [1], [4].
This review discusses the XRN family of exoribonucleases, focusing on their molecular functions and biological impacts. We first indicate the roles of XRNs within fundamental RNA decay pathways, and then describe the structures, locations and contributions of different XRNs to RNA decay in a variety of processes. In addition to reviewing these topics, we highlight three case studies in more detail: small RNA-associated functions of plant XRN4, the role of human XRN2 in 5′ → 3′ decay during pausing and termination by RNAP II, and the role of XRN1 (PACMAN) in Drosophila development. Note that although different systems differ in their nomenclature for protein and gene names, primarily with regard to capitalization, this review will use all uppercase for simplicity (with genes italicized and mutant alleles lowercase and italicized).
Much of our understanding of the XRN family and the mechanisms of mRNA decay comes from studies in the yeast Saccharomyces cerevisiae. However, studies using other eukaryotic organisms have added to our understanding of the molecular and biological roles of XRNs in multicellular organisms [1], [3], [4], [5], [6], [7], [8]. In this section, major mechanisms of both cytoplasmic and nuclear decay are discussed, all of which involve XRN activity.
In general, the decay of most eukaryotic mRNAs occurs by three major pathways 1) deadenylation-dependent 2) deadenylation-independent and 3) endonucleolytic cleavage-dependent decay (Fig. 1). As its name implies, the first rate-limiting step of deadenylation-dependent mRNA decay involves shortening of the poly(A) tail prior to 5′ cap removal (i.e. decapping) and subsequent degradation [2], [8]. One of more deadenylase enzymes, CCR4-CAF1-NOT1 or PARN, progressively trims and nearly removes the 3′ poly(A) tail [2], [9]. Following this deadenylation, the mRNA can undergo degradation in either the 5′ → 3′ or 3′ → 5′ direction (Fig. 1A). As deadenylation is completed, in the 5′ → 3′ decay pathway, the LSM1–7 proteins bind to the 3′ end of the mRNA and recruit the decapping complex [10], [11], [12]. Decapping enzymes such as DCP2, with additional cofactors, hydrolyze the 5′ cap, exposing the mRNA to decay that is carried out by XRN1, a processive exoribonuclease that completely hydrolyzes decapped (5′ monophosphorylated) RNA in the 5′ → 3′ direction (Fig. 1 A1) [4], [5], [8], [13], [14]. This pathway bears a similarity to the 5′ → 3′ RNA decay in prokaryotes which is also specific for 5′ monophosphorylated RNA [15], [16]. In eukaryotes, after deadenylation the mRNA can also be degraded in the 3′ → 5′ direction, primarily through the activity of the multi-subunit exosome complex (Fig. 1 A2) [17], [18]. This macromolecular complex has a central core arranged in a ring consisting of six catalytically inactive 3′ → 5′ exoribonucleases [18]. Depending on the subcellular localization, the exosome core associates with catalytically active subunits: a distributive RNase D 3′ → 5′ exoribonuclease, RRP6 (nucleus and nucleolus), and/or a processive RNase II 3′ → 5′ exoribonuclease RRP44/DIS3 (cytoplasm and nucleus) [19], [20], [21], [22]. RRP44 also has a highly conserved PilT N-terminus (PIN) domain with endoribonucleolytic activity [23], [24], [25], [26]. Exosome-mediated 3′ → 5′ degradation in the cytoplasm is followed by hydrolysis of the remaining cap-structure by DCPS (DCS1 in yeast), a “scavenger” type decapping enzyme [27], [28], [29]. These two directions of mRNA degradation occurring after poly(A) shortening are referred to as deadenylation-dependent RNA decay and represent the major decay mechanisms for RNA turnover in the cytoplasm, at least in yeast. SOV, another component of cytoplasmic 3′ → 5′ RNA decay, was first identified in Arabidopsis as a suppressor of VARICOSE/HEDLS, a decapping scaffold protein [30]. SOV is a member of the RRP44/DIS3 family that contains a conserved RNaseII domain but SOV lacks the PIN-domain required interacting with the core exosome and falls in a separate cluster within the family [30]. RNA stability data indicate that substrates of SOV overlap with those of the decapping complex [30]. Recently the function of SOV homolog, DIS3L2 has been described in yeast Schizosaccharomyces pombe and humans [31], [32]. DIS3L2 preferentially degrades uridylated substrates in S. pombe in vitro and it appears to function independent of the exosome in all three systems [30], [31], [32]. This suggests that SOV/DIS3L2 represents a cytoplasmic RNA decay pathway alternative to XRN1- and exosome-mediated degradation (Fig. 1 A2).
Another mRNA degradation mechanism involves the internal cleavage of mRNA to create unprotected 5′ and 3′ fragments that are substrates for exoribonucleolytic decay (Fig. 1B). Until recently, study of mRNA degradation had focused mainly on exoribonucleolytic decay from the ends. Yet, it is now apparent, that many pathways utilize endoribonucleases (e.g. AGO, SMG6, and RRP44/DIS3) [33]. One example of this in both plants and animals occurs via small RNAs (20–30 nt long) acting as guides in silencing complexes by directing AGO proteins to specific target mRNAs [34], [35], [36], [37]. Endonucleolytic cleavage is achieved by an AGO slicer activity if the small RNA is highly complementary to the target, if not, other decay mechanisms that may be linked to translational inhibition can take place [34], [36]. If cleavage by AGO does occur, XRN1 or XRN4 degrade the 3′ mRNA fragment while the 5′ fragment is degraded by the exosome [36], [38], [39]. In Drosophila, transcripts containing premature termination codons (PTCs) are degraded via a SMG6-mediated endonucleolytic mechanism, followed by exoribonucleolytic decay of the cleaved 5′ and 3′ fragments by the exosome and XRN1, respectively [40], [41], [42].
As shown in Fig. 1B2, some mRNAs undergo 5′ → 3′ decay without the removal of poly(A) tail (e.g. S. cerevisiae transcripts recognized for Nonsense-Mediated Decay (NMD), and RPS28B and EDC1 mRNAs) [43], [44], [45]. As part of the cytoplasmic mRNA surveillance system, aberrant mRNAs, mainly NMD substrates, also predominantly undergo 5′ → 3′ degradation without the need for deadenylation [44], [46]. NMD factors (e.g. UPF and SMG proteins) recognize and facilitate decay of transcripts containing PTCs, and thereby prevent the generation of 3′ truncated proteins that could be detrimental to the cell [44], [47]. XRN1 degrades these aberrant RNAs following 5′ cap removal by DCP2 initiated by the NMD factors [5], [44]. An alternative pathway contributes to the degradation of PTC-containing mRNAs and relies on rapid deadenylation followed by 3′ → 5′ decay via the exosome [46].
As in the cytoplasm, the nucleus also has 5′ → 3′ and 3′ → 5′ decay mechanisms that are critical for nuclear RNA turnover. Unlike mRNAs stabilized by polyadenylation, several nuclear RNAs, including rRNAs, are destabilized via polyadenylation by the TRAMP complex, leading to accelerated exosome-mediated 3′ → 5′ decay [48]. Unspliced and incorrectly polyadenylated RNAP II transcripts undergo degradation by the nuclear exosome [17], [49]. Work in yeast indicates that nuclear-restricted mRNAs are degraded by XRN2 (RAT1) after they are decapped by machinery recruited by the LSM2–8 proteins [6], [50].
Section snippets
Structure and mechanistic functions of XRNs
The XRN family members were first identified in S. cerevisiae (XRN1 175 kDa and XRN2, 115 kDa) and have been studied extensively over the past three decades [51], [52], [53], [54]. Orthologs of XRN1 and XRN2 have been identified in most key model organisms, and in humans [55], [56], [57], [58], [59], [60], [61], [62], [63], [64], [65].
XRN1 and XRN2 (Fig. 2A and B) show extensive conservation within their N-terminal regions and share an active site and mechanism of action. The sequence similarity
Molecular functions of XRN1
XRNs have molecular functions in a number of key processes (listed in Table 2). In addition to bulk 5′ → 3′ mRNA turnover (Fig. 1 A1), XRN1 degrades a wide range of cytoplasmic RNAs, including noncoding RNAs and NMD-substrates. Yeast XRN1 degrades a novel class of long noncoding RNAs (lncRNAs) called XRN1-sensitive Unstable-Transcripts (XUTs) after they are decapped, particularly those antisense to open reading frames [72]. XRN1 substrates also include GAL lncRNAs, which overlap with GAL
Co-factor interactions of XRN1 and XRN2
The diversity of molecular functions of XRN1 and XRN2 predict that multiple interacting factors are involved. This is indeed the case, as highlighted in Table 3. This table and the sections below are not intended to provide an exhaustive description of interacting partners, but instead focus on key interactions of XRN1 and XRN2 with cofactors from specific decay pathways.
Location of XRN family proteins within eukaryotic cells
The location of XRN enzymes in the cell has provided information on their function and probable targets. XRN2 and related enzymes (e.g. Arabidopsis XRN2/3 and Trypanosome brucei XRND) are known to be located in the nucleus and nucleolus, where they are involved in the maturation (5′ trimming) of rRNAs and snoRNAs, as well as contributing to transcription termination by nuclear RNAP I and II [64], [65]. The larger XRN1s and functionally-related proteins (e.g. Arabidopsis XRN4, Drosophila PACMAN,
Biological functions of XRNs
Although the molecular functions of XRNs have been elucidated (see Section 3), the biological functions of XRNs responsible for observed phenotypes in mutants, are less well known (Table 4). Experiments to determine the biological consequences upon mutation or knockdown of XRN1 or XRN2 homologs are essential in order to determine the true biological targets of XRNs.
Small RNA-associated functions of plant XRN4
The Arabidopsis system has been the source of most information about plant XRNs and their roles in interesting small RNA-associated RNA decay mechanisms. The earliest report of an XRN enzyme playing a role in an RNA decay pathway initiated by small RNA-mediated cleavage came from Arabidopsis with the demonstration that XRN4's substrates include miRNA targets. Specifically, XRN4 degrades the downstream cleavage product produced when a miRNA guides an AGO-protein to cleave the target in the
Conclusions and future prospects
The 5′ → 3′ exoribonucleases XRN1, XRN2 and related XRNs have proved to be fascinating enzymes, with key functions in RNA decay and roles in a number of other important cellular processes including NMD, PTGS and small RNA-mediated decay. During the past several years, especially exciting discoveries have been made. Solving the XRN1 crystal structure made it easy to understand the enzymatic mechanism of the XRN family in general. New substrates and interacting partners have been identified, and
Acknowledgements
We thank Renate Wuersig and Karl Franke for valuable comments on the manuscript. XRN work in the authors' laboratories was primarily funded by grants from the National Science Foundation (MCB1021636) and the National Institute of Health (GM096471) to P.J.G. and the Biotechnology and Biological Sciences Research Council (BB/I021345/1 and BB/I007989/1) to S.F.N., with additional support from the Department of Energy (DE-FG02-07ER64450) to P.J.G.
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This article is part of a Special Issue entitled: RNA Decay mechanisms.