RNA lifetime control, from stereochemistry to gene expression

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ScienceDirect RNA lifetime control, from stereochemistry to gene expression Tom Dendooven, Ben F Luisi and Katarzyna J Bandyra Through the activities of various multi-component assemblies, protein-coding transcripts can be chaperoned toward protein synthesis or nudged into a funnel of rapid destruction. The capacity of these machine-like assemblies to tune RNA lifetime underpins the harmony of gene expression in all cells. Some of the molecular machines that mediate transcript turnover also contribute to on-the-fly surveillance of aberrant mRNAs and non-coding RNAs. How these dynamic assemblies distinguish functional RNAs from those that must be degraded is an intriguing puzzle for understanding the regulation of gene expression and dysfunction associated with disease. Recent data illuminate what the machines look like, and how they find, recognise and operate on transcripts to sculpt the dynamic regulatory landscape. This review captures current structural and mechanistic insights into the key enzymes and their effector assemblies that contribute to the fate-determining decision points for RNA in post-transcriptional control of genetic information. Address Department of Biochemistry, University of Cambridge, Tennis Court Road, Cambridge CB2 1GA, UK Corresponding authors: Luisi, Ben F ([email protected]), Bandyra, Katarzyna J ([email protected])

Current Opinion in Structural Biology 2020, 61:59–70 This review comes from a themed issue on Macromolecular assemblies Edited by Xiaodong Zhang and Tom Blundell

https://doi.org/10.1016/j.sbi.2019.10.002 0959-440X/ã 2019 Elsevier Ltd. All rights reserved.

Introduction It is helpful to consider a transcript as a transient, information-rich molecule. Beyond the codon pattern that it might carry for protein translation, RNA can also encode structural features that define its lifetime or the potential to be translated before turnover, and its maturation can unmask hidden information made available through interactions with other transcripts [1,2]. Regulation of RNA lifetime, in coordination with adjustments to rates of synthesis, maintains transcript homeostasis, enables rapid change of gene expression, and signals cellular status [3–6,7,8

,9]. The life www.sciencedirect.com

experience of prokaryotic and eukaryotic messenger RNA, starting from biogenesis and maturation, then passing through quality controls, transport and arrival at subcellular destination, and then on to its eventual destruction, are depicted schematically in Figures 1 and 2. Multi-subunit molecular machines orchestrate these events in the individual cellular organelles and compartments. This review presents recent findings and new concepts on the structure and function of some of these machines in eukaryotes and prokaryotes, and it is thematically organised by the compartments where the different types of assemblies operate (summarised in Table 1).

Events in the bacterial cytoplasm Prokaryotic mRNA degradation has been extensively characterised using representative model organisms such as the gram-negative Escherichia coli and the gram-positive Bacillus subtilis, which arose from highly divergent bacterial lineages (See Figure 1). RNA decay in these bacteria involves multi-enzyme complexes built around a membrane-bound multi-domain ribonuclease scaffold: RNase E in the case of most gram-negative bacteria [10] (Figure 3d) and RNase Y in model gram-positive microorganisms [11] (Figure 1b). RNase E and RNase Y are both endonucleases with preference for single stranded, 50 -monophosphorylated RNA species [12–14] (Figure 1a,b). Recent structural studies have provided insights into how an RNA duplex structure is recognised and used to align substrates for preferred cleavage by RNase E [15]. The ribonucleases assemble into functionally conserved multi-enzyme machineries, often referred to as RNA degradosomes, with both functional and structural parallels with the exosome and the RISC (RNA-induced silencing complex) machinery [16] (Figure 3c and d). Small regulatory RNAs (sRNAs) and RNA chaperones have been shown to co-regulate RNA turnover in conjunction with degradosome assemblies [2]. In many bacterial species, different RNAs are degraded by PNPase, also mentioned in Figures 2 and 3c and d for the roles of its homologue in the mitochondrium and structural similarities with the exosome core assembly [17]. Apart from RNA decay, PNPase can play a structural role in protecting and facilitating some regulatory RNAs in complexes with the RNA chaperone Hfq, with potential analogy to carrier functions for PNPase in human mitochondria [18]. In general, RNA chaperones, such as Hfq and ProQ, seem to be key assistants in almost all riboregulatory processes in bacterial cells. For example, in addition to its role to repurpose PNPase as a stable carrier of sRNA, Hfq has also been shown to engage regulatory sRNAs and mRNAs and form Current Opinion in Structural Biology 2020, 61:59–70

60 Macromolecular assemblies

Figure 1

(a)

70S

RNA

5’PPP

RNA

5’PPP

polyA polymerase

RppH RNA degradosome, RNase G

5’P

exonuclease

AAAAAA

oligoribonuclease

PHASE SEPARATION

(b)

AAAAAA

PPP PP

‘Decapping’

RppH

Polyadenylation

polyA polymerase

3’->5’ Degradation

5’->3’ Degradation

PNPase RNase II RNase R

RNase J1/J2

Endonucleolytic cleavage

RNase E RNase G RNase Y RNase J1/J2 Current Opinion in Structural Biology

Bacterial RNA decay. (a) RNA decay pathway in most known bacteria. RNA substrates are monophosphorylated at the 50 end by RppH activity and a poly-A tail is added at the 30 end by Poly A polymerase. Subsequently endoribonucleases, exoribonucleases and ribonuclease assemblies engage the tagged RNA targets to generate short oligomeric RNA products. Oligoribonuclease completes the decay pathway by further degradation of the RNA oligos into nucleotides. The RNase E scaffold domain is natively unstructured and was shown to undergo liquid–liquid phase separation forming ribonucleotprotein droplets that resemble P-bodies to locally drive forward RNA decay reactions. (b) Overview of prokaryotic RNA degradation enzymes. Many of these ribonucleases and helper enzymes can transiently associate in pleiotropic degradosome derivates depending on cellular conditions.

higher order assemblies with other partner proteins into effector complexes that coordinate gene expression in response to changing environments [19,20]).

Events in the eukaryotic nucleus Surprisingly, RNA genesis in the nucleus is grossly inefficient, with an estimated 50% of coding transcripts turned over under normal growth conditions in yeast [21]. Current Opinion in Structural Biology 2020, 61:59–70

Nonetheless, this apparent wastefulness can help to optimise transcript fidelity and response time of regulation. For example, the nuclear RNA decay machinery of Saccharomyces cerevisiae has been shown to control the abundance of protein coding mRNA [22

]. This machinery also processes functional RNA molecules and eliminates those that are incorrectly processed. Often, the prime determinant of the activity of the decay machinery is accessibility of the www.sciencedirect.com

RNA lifetime Dendooven, Luisi and Bandyra 61

Figure 2

MITOCHONDRIUM

(a) AAAAAAAAAAAA

7-meGpppG

PNPase-SUV3 CYTOPLASM NUCLEUS

PNPase Rat1/Xrn2

AAAAAAAAAAAA

7-meGpppG

RNA TPase, GTase, Guanine-N7 MTase

CPF/CPSF

Pab1 AAAAAAAAAAAA

7-meGpppG Dcp2

DXO/Rai1

TUT4/TUT7 Ccr4 7-meGpppG

AAAAAAAAAAAA

AAAAAAAAAAUUU

7-meGpppG Xrn1

AAAAAAAAAAAA

7-meGpppG

AAAAAAAAAAAA

Pan2/Pan3 Ccr4-Not

7-meGpppG

Dis3L2 Exosome?

Exosome

P-BODY

(b) A A

m7-GDP

A

AAAAAAAAAAAAAAA

7-meGpppG GpppN

AA A

Decapping

Dcp2 Nudt16 Dxo1 Rai1

Deadenylation

Pan2-Pan3 complex

3’->5’ Degradation

Exosome

5’->3’ Degradation

Xrn1 Rat1(Xrn2)

Ccr4-Not complex Current Opinion in Structural Biology

Eukaryotic RNA decay. (a) RNA decay pathway in the eukaryotic nucleus and cytoplasm. In the nucleus, nascent mRNA species are polyadenylated by CPF at the 30 end, capped at the 50 end and transported to the cytoplasm for translation. RNA decay in the nucleus is initiated by deadenylation by Pan2/Pan3 and Ccr4-Not complexes and decapping by DXO/Rai1 for aberrant transcripts, followed by exoribonucleolytic decay by the nuclear exosome. In the cytoplasm, mRNA can be transported to the mitochondrium or translated. In the mitochondrium PNPase together with the helicase SUV3 is responsible for the bulk RNA turnover, but PNPase can alternatively take on RNA carrier roles. In the cytoplasm Pab1 coats the poly-A tail and either protects the transcript or recruits deadenylation machineries. Ccr4 deadenylates most transcripts so that the cytoplasmic exosome can access the 30 end and digest mRNAs. Alternatively, Dcp2 can decap aberrant transcripts, followed by 50 -30 Xrn1-mediated decay. Finally, TUT4/ TUT7 can extend the 30 poly A tail with an oligo-U stretch, which triggers degradation by Dis3L2. RNA degrading complexes can, together with their RNA substrates compartmentalise locally into P-bodies via liquid–liquid phase separation to drive forward RNA decay reactions. (b) Overview of eukaryotic canonical RNA degradation machineries acting on a transcript. The exosome core is depicted in yellow. The helicase complex (Skicomplex or TRAMP complex) is a beige hexagon and the Dis3/rrp4 and rrp6 ribonucleases are depicted in brown.

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62 Macromolecular assemblies

Table 1 Assemblies and components of RNA processing and turnover described here Assembly

Organism

Compartment

Comment

References

Exosome-helicase complexes

yeast

nucleus

[49,50]

Exosome

human

nucleus

Pan2 exonuclease in complex with poly-A Pan2-Pan3 deadenylase complex with polyA/Pab1 Crc4-NOT complex

yeast

yeast

cytoplasm/ nucleus cytoplasm/ nucleus nucleus

DXO/Rai1

mouse & yeast

cytoplasm/ nucleus

Ski complex

yeast

cytoplasm

UPF1

many eukaryotes Caulobacter crescentus & Escherichia coli human

cytoplasm

14-subunit complex with 5.8S rRNA precursor. Reveals RNA substrate channeled from helicase yMtr4 into the exosome core Complex structures by cryo-EM. The major function of the human nuclear exosome is in quality control pathways that counteract pervasive transcription initiation and defective transcription termination from promoter upstream transcripts (PROMPTs) that arise due to antisense transcription from bidirectional promoters and prematurely terminated products of protein coding genes. Specificity is determined by recognition of the distinctive conformation of the phosphate backbone. Pab1 threads poly A into the Pan2-Pan3 complex, which recognises the poly-A bound Pab1 oligomer Polymerase module by cryo-EM. “CPFcore” complex together with accessory cleavage factors IA and IB (CF 1A and CF 1B). Ysh1 is positioned in proximity to the mRNA CAAA motif and allosterically activated by CF IA for cleavage The crystal structures of deNADding complex and evidence that DXO plays role in turnover of NAD capped RNAs in mammals Specialised SKI complex assists the cytoplasmic RNA exosome Helicase involved in nonsense mediated decay.

RNA degradosome

Polynucleotide phosphorylase

yeast

[62]

[29

] [28

] [23

,24

]

[37]

[55,56] [58,59,93]

cytoplasm

Liquid–liquid phase transition, from fluorescence microscopy. The assembly is associated with the cytoplasmic membrane in E. coli.

[88

,89]

mitochondria

RNA shuttling in and out of the mitochondria; in complex with RNA helicase involved in RNA turnover

[66,67]

transcript’s 50 and 30 ends to exonuclease attack, and accordingly, the termini of nascent transcripts are protected by modifications as they spool from the polymerase. Transcripts encoding proteins are capped at the 50 end, spliced if bearing introns, cleaved in the 30 untranslated region, polyadenylated on the freshly generated 30 end, and finally exported to the cytoplasm (Figure 2a).

the final module in the adenylation cascade, defines the configuration of the four subunits that cooperate to generate the poly-A tail [24

]. Remarkably, the polymerase domain has similar architecture to complexes involved in DNA repair and mRNA splicing [24

], illustrating how intricate assemblies can be repurposed for dedicated tasks in the course of evolution.

Polyadenylation of nascent transcripts is mediated by the cleavage and polyadenylation factor complex (known as CPF in yeast) (Figure 2a). Structural data indicate this to be a dynamic assembly, licensed to cleave only when all protein factors converge at the polyadenylation site [23

]. The subunits of CPF are organised into three functional modules with nuclease, polymerase and phosphatase activity [24

] (Figure 3a). The latter serves as the first step in the adenylation cascade and dephosphorylates the C-terminal tail of RNA polymerase II to signal the approach to transcription termination. The nuclease domain includes the ribonuclease subunit Ysh1, which is activated when incorporated into the CPF complex. Ysh1 is positioned in proximity to the mRNA cleavage site and allosterically activated by cleavage factor IA (CF IA) [23

] (Figure 3a). Cryo-EM analysis of the polymerase module,

The nascent poly-A tail protects the transcript’s 30 end, and removing it is the primary, rate-limiting step in mRNA turnover and a key regulatory control point. Tail removal occurs through the hydrolytic activities of the Pan2-Pan3 deadenylase complex and the multi-functional Ccr4–Not complex [25,26,27] (Figures 2a and b, 3b). Models for decay propose a biphasic process, in which the poly-A tail is quickly shortened by the Pan2/ 3 nuclease complex, followed by kinetically slower cleavage by the Ccr4-Not complex (Figure 3b). Ccr4-Not can target specific transcripts through facilitator proteins, such as Puf3, which forms a bridge between Ccr4-Not and mRNAs recognised by a destabilising motif [26]. The Pan2-Pan3 complex has a complicated relationship with the poly-A-binding protein Pab1 which both protects the tail but can also trigger tail trimming by the complex

Current Opinion in Structural Biology 2020, 61:59–70

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RNA lifetime Dendooven, Luisi and Bandyra 63

Figure 3

(a)

(b)

Pab1

7-meGpppG

RNA Pol II termination

Pan2/Pan3 complex recruitment

Phos Nuc

CPF

5’ UAUAUA CF1B

A

A AU

AA

Ysh

1

CA

7-meGpppG Pan2/Pan3 Deadenylation Ccr4-Not recruitment

Pol 7-meGpppG

Ccr4-Not Deadenylation Degradation

CF1A A

3’

U-rich

(c)

(d)

Ski2

RNase E

Ski3 Ski8

Endonuclease

Helicase Mtr4

Helicase

RhIB

Enolase Rrp47

Exosome core Rrp6 Exo-9

PNPase

Exonuclease

Exonuclease Rrp44

Nucleus

Cytoplasm Current Opinion in Structural Biology

Schematic structures of RNA processing and decay machineries. (a) Polyadenylation of nascent transcripts: the yeast CPF assembly. The Mpe1/Ysh1 subcomplex of the CPF nuclease core has been structurally characterised (adapted from Ref. [23

]), as well as the polymerase module [24

]. A model for the cooperation of the modules to recognise cleavage sites in a transcript. The scaffold (Cft1, Pfs2, Yth1) of the polymerase module (Cft1, Pfs2, Yth1, Fip1, Pap1) directly recognises the upstream stimulatory RNA sequence (i.e. the canonical AAUAAA element (see Refs. [60,61]), or AAGAA in a CYC1 model substrate). In contrast, the nuclease module proteins do not bind RNA very tightly (see Refs. [23

]). The polymerase module also has a more general role as an assembly hub for the complete machinery. (b) Schematic model for eukaryotic cytoplasmic deadenylation. During the lifetime of the transcript, the poly-A tail is shortened sequentially, finally resulting in abrupt mRNA degradation. The model proposes a biphasic process, in which the polyA tail is quickly shortened by the Pan2/3 nuclease complex, followed by kinetically slower cleavage by the Ccr4 nuclease in the Caf1/Ccr4 nuclease complex. Pab1 physically protects the end of the transcript from non-specific decay but it is needed for poly-A tail trimming. Pab1 stimulates Pan2-Pan3, the Ccr4-Not deadenylase complex then trims the tail to a short poly(A), rendering the transcript susceptible to decapping followed by degradation via Xrn1 and/or to 30 ->50 degradation by the exosome-Ski complex. How the Pan2-Pan3 specifically recognises early poly(A) RNPs and why it stops after shortening them to a discrete size has been illuminated by cryo-EM structure of the Pan2-Pan3/polyA/Pab1 RNP. This explains the poly(A) RNA-dependent oligomerisation of Pab1 and PABPC1 in the poly-A RNP. The structure also explains why Pan2-Pan3 degrades the poly-A tail in bursts, due to stepwise removal of Pab1 molecules. When the tail is very short in human cells, it can gain a poly-U tail (by the action of terminal uridyl transferase), which makes it a target for the 30 exonuclease Dis3L2 and the 50 exonuclease Xrn1. Adapted from [9]. (c) The nuclear and cytoplasmic exosome. The major function of the human nuclear exosome is quality control that deals with products of pervasive transcription initiation and defective transcription termination. As such, the nuclear exosome targets promoter upstream transcripts, from

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Current Opinion in Structural Biology 2020, 61:59–70

64 Macromolecular assemblies

(Figures 2a, 3b). How the Pan2-Pan3 complex switches between those states, and also how it focuses degradation on just the poly-A tail and not the rest of the transcript has been illuminated by the cryo-EM structure of the Pan2Pan3 complex in association with Pab1 and poly-A [28

]. These data reveal that the oligomeric structure of the RNA-binding domains of Pab1 acts as a molecular ruler that matches a defined surface of the Pan2-Pan3 complex. Moreover, the crystal structure of the Pan2 exonuclease in complex with poly-A RNA shows that specificity is also achieved through recognition of the distinctive conformation of the ribophosphate backbone of that homopolymer [29

], and not through base-specific contacts as seen in many other poly-A-binding proteins [30]. Enzymes that modify or remove 50 ends also provide a regulated route into decay pathways [31,32]. The eukaryotic 50 RNA cap is usually a 7-methyl guanosine (m7G). In the nucleus, the DXO/Rai1 family of decapping enzymes provide a mechanism of 50 -end capping quality surveillance that removes RNAs with incomplete caps [33–36] (Figure 2b). Strikingly, capping of mRNAs with nicotinamide adenine dinucleotide (NAD) has the opposite effect of the canonical m7G protective cap in that it can promote decay through DXO mediated removal of the NAD, a process called ‘deNADding’ [37]. The crystal structures of DXO and Rai1 in complex with the NADcapped RNA mimic, 30 -phospho NAD+ (30 -NADP+), demonstrate that deNADding is accomplished by the same active site that removes m7G [37]. Without a normal 30 poly-A tail or 50 cap, nuclear transcripts become accessible for RNA degradation machineries. The key machinery of the nucleus for RNA turnover in widely diverse eukaryotes is the highly conserved, multi-subunit exosome (Figures 2b and 3c). As an indication of the functional importance of the exosome, its dysfunction is associated with severe neurological disorders [21]. Importantly, the exosome turns over aberrant transcripts, including defectively processed stable RNAs, and coding transcripts that are not spliced properly or

have aberrant 30 ends [38–43]. The exosome effectuates not only degradation, but also the processing of precursors of rRNA, small nucleolar RNA (snoRNA) and small nuclear RNA (snRNA) [44,45]. Although the assembly has little apparent preferences per se, the exosome can target defined substrates by recruiting cooperating specificity factors to recognise RNA sequence elements, secondary structures and modifications. In Saccharomyces pombe, for example, the RNA-binding protein Mmi1 recognises a sequence pattern and tags meiotic mRNAs cotranscriptionally for rapid exosome-dependent decay [21]. ATP-dependent RNA-unwinding activities are central to the regulation of the exosome, and they facilitate threading of the RNA substrate through the narrow channel entrance which can accommodate only single stranded RNA (Figure 3c). In S. cerevisiae, the conserved DExHbox helicase Mtr4 is required for RNA degradation in the nucleus and is part of the Trf4/5-Air1/2-Mtr4 polyadenylation (TRAMP)-complex that aids in the degradation of numerous surveillance targets such as hypomodified precursor of initiator tRNA, defective pre-rRNAs and cryptic RNA polymerase II (RNAPII) transcripts [9,46]. Furthermore, Mtr4 can act together with PABPN1 and the zinc finger protein ZFC3H1 to stimulate exosome degradation of defective nuclear transcripts [9] and to help mature the 60S large ribosomal subunit, together with other ribonucleoproteins (RNPs), which can be considered a form of pre-translational surveillance. In S. cerevisiae Mtr4 interacts with the early-acting and late-acting ribosomal biogenesis factors Utp18 and Nop53 that direct the exosome to specific pre-ribosomal RNP substrates [47,48]. A recent cryo-EM structure shows how the nuclear exosome interacts with the Mtr4 helicase to thread RNA substrates through the central channel to the Dis3 nuclease subunit at the opposite end [49,50] (Figure 3c). Mtr4-containing exosomes can displace a DNA strand while translocating on RNA, potentially explaining the role of the exosome in controlling targeted strand-specific DNA mutagenesis in antibody genesis [49,51]. Finally, cryo-EM reveals how

(Figure 3 Legend Continued) antisense transcription from bidirectional promoters, and prematurely terminated mRNAs [62]. The exosome core is a hexameric ring comprising six proteins related to the phosphoryolytic exoribonuclease RNase PH and is structurally conserved in Archaea (brown). At one end of the central channel there are three subunits that are RNA-binding modules belonging to the S1 and KH structural groups. The exosome has endo-ribonucleolytic and 30 ->50 exo-ribonucleolytic activities, and in eukaryotes, exosome catalytic activity is dependent on the associated processive 3-0 ->50 exonuclease Dis3/Rrp44 (blue), which is positioned at the opposite end of the barrel from the RNA-binding modules. Structural and functional data indicate that approximately 30 nt of single-stranded RNA must be threaded through the long central channel to reach the Dis3/Rrp44 active site (red). In the yeast complex, Rrp44 can adopt either a closed conformation to support the RNA channel path or an open conformation to support a short direct path to the exoribonuclease site [63,64

]. In the human complex, the RNA channel path is achieved by an open conformation of hDIS3. Alternatively, RNA can be degraded by Rrp6, a distributive exonuclease, assisted by its cofactor Rrp47. Interestingly, Rrp6 can allosterically stimulate the nucleolytic activity of Dis3 in budding yeast, probably by direct RNA-binding as well as through widening of the channel [64

,65]. The exosome has different compositions of accessory factors in different compartments (nucleus and cytoplasm). For example, in most organisms, the nucleolar exosome complex is associated with a poly-A polymerase. Furthermore nuclear exosomes carry Mtr4 as a helicase (left) whereas cytoplasmic exosomes are associated with the Ski-complex (Ski2 as a helicase, right). The structure used for the figure is 6fsz [50]. (d) Correspondence of analogous and homologous components of the bacterial RNA degradosome with the exosome. The exoribonuclease PNPase has a similar architecture to the exosome core yet has nucleolytic activity. The DEAD box helicase RhlB unwinds RNA as part of the degradosome. Initial cleavage is performed by RNase E, an endonuclease. Current Opinion in Structural Biology 2020, 61:59–70

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RNA lifetime Dendooven, Luisi and Bandyra 65

the exosome transiently associates with the pre-60S ribosomal subunit during ribosome maturation [50]. As part of the exosome effector complex, Mtr4 docks onto the 25S rRNA and funnels the 30 end of the 5.8S rRNA towards the exosome channel and Dis3.

Events in the eukaryotic cytoplasm In the eukaryotic cytoplasm, the mRNA decay pathway involves shortening of the poly-A tail, followed by 50 end decapping and then exonucleolytic degradation (Figure 2a. The poly-A tail can be coated in the cytoplasm by protein Pab1 (PABPC1 in humans), which not only protects the 30 end from the deadenylases but also facilitates translation initiation [25,52] (Figure 2a). Ccr4-mediated deadenylation appears to be a key pathway for turnover of most cytoplasmic mRNAs. The addition of 30 uracils to short poly-A tails by TUT4 and TUT7, however, may stimulate decay through recruitment of the Lsm1–7–Pat1 complex to enhance decapping or through preferential degradation by the nuclease Dis3L2 [53] (Figure 2a). Transcriptomics data indicate that decapping is an important step for mRNA decay [46]. Deprotection of the 50 end is initiated by the Nudix hydrolase family of decapping enzymes (which is a different family from the nuclear decapping enzymes mentioned above) that initiate 50 -30 decay by hydrolyzing 50 N7-methylG caps and releasing m7GDP [54]. The decapped transcripts are then degraded by the processive 50 ! 30 exoribonuclease Xrn1 (Figure 2a). In vivo photocrosslinking experiments in yeast confirm that decapping factors preferentially engage mRNAs with suboptimal codons, supporting models of rapid turnover of transcripts that are inefficiently translated [46]. In the cytoplasm, exosome-dependent degradation occurs during translation (Figure 2, 4a). The first high resolution insights into the physical links between translation and degradation complexes was elucidated by cryoEM, revealing how the Ski helicase complex, which assists the cytoplasmic exosome in RNA 30 ! 50 decay and quality control pathways, can associate with a stalled 80S ribosome [55]. This interaction helps expose the 30 end of mRNA directly off the ribosome, via the Ski2 helicase channel, for degradation by the exosome (Figure 4a). The structure of the Ski-80S complex suggests that Ski2 helicase activity is triggered by association with the ribosome, which raises the question as to how mRNA 30 UTRs, devoid of ribosomes, are degraded by the Ski-exosome machinery (Figure 3c). Zhang et al. [56] discovered a new component, Ska1, that can associate with Ski-exosome complexes and assist in 30 -50 degradation of longer, ribosome-free, 30 UTRs of mRNA. By assisting the Ski-exosome complex in degrading mRNA stretches in non-translated sequences, Ska1 complements the canonical Ski-exosome-ribosome cooperative assembly for co-translational surveillance. It is plausible that www.sciencedirect.com

Ska1 is necessary for initial degradation of the 30 untranslated region of a defective mRNA but is jettisoned when the processive Ski-exosome complex reaches a stalled ribosome. Transcripts can also be degraded cotranslationally from the last translating ribosome [57]. This occurs in a 50 to 30 direction and is likely mediated by the exonuclease Xrn1. Another decay pathway that can impact on gene expression is nonsense-mediated mRNA decay (NMD), which is a translation-dependent surveillance mechanism that controls expression of 10% of mRNAs in mammals [58]. It was found that NMD can remodel the transcriptome mRNAs involved in differentiation by ensuring that key transcripts are tagged with NMD-inducing signatures. NMD is regulated by recruitment of the endoribonuclease SMG6 or the Ccr-Not complex through SMG5-7 factors and a helicase, UPF1 [58]. Recent data indicate that UPF1 helicase can associate with nascent RNAs at active transcription sites in Drosophila and as such assist in on-the-fly surveillance during translation [59].

RNA decay and processing in the mitochondria Multi-enzyme degradation machines are also found in the mitochondria of metazoans and single cell eukaryotes such as yeast (Figure 2a). In human mitochondria, polynucleotide phosphorylase (PNPase), which is an important exoribonuclease evolutionarily related to the exosome, interacts with the SUV3 helicase to provide efficient degradation of redundant RNAs [17]. Mutations of PNPase result in mitochondrial double-stranded RNA accumulation coupled with activation of innate immune defence mechanisms that have evolved to protect vertebrates against microbial and viral attack. The localisation of PNPase at the mitochondrial inter-membrane matrix suggests that it has a dual role in preventing the formation and release of mitochondrial double-stranded RNA into the cytoplasm [66] which in turn prevents the activation of these immune responses. PNPase also is involved in the import of long non-coding RNAs, and recent data suggest that it serves as a carrier protein to help import the RNA component of telomerase for processing and then exports the processed form back to the cytoplasm [67]. Hints about how the exoribonuclease might toggle between exoribonuclease and carrier modes has come from studies of the E. coli PNPase complexes with regulatory RNAs [18], which was described in the earlier section on bacterial RNA metabolism.

On-the-fly surveillance opportunities of transcription and translation in bacteria Transcription-translation-degradation coupling provides mRNA quality control, as synthesis of non-translated transcripts is terminated [68], and their decay is promoted [69]. In eukaryotes, cytoplasmic RNA decay happens Current Opinion in Structural Biology 2020, 61:59–70

66 Macromolecular assemblies

Figure 4

(a)

EUKARYOTA

(b)

BACTERIA Degradasome RhIB

RNAP

Nucleus

?

Tagging for protection

7-meGpppG

AAAAAAAAAAAA

RNAP

Ribosome stalling/pausing

Tagging for decay

P

AAAA

Exosome Ski complex

7-meGpppG Ribosome stalling/pausing

AAAAAAAAAAAA

Un-tagging

7-meGpppG Current Opinion in Structural Biology

On-the-fly surveillance modes of substrate access. (a) RNA Decay by the cytoplasmic exosome can happen during translation in Eukaryotes. A physical link between these two complexes is formed by the Ski helicase complex. Tollervey and colleagues have postulated that eukaryotic RNA transcripts are degraded unless protected by RNPs or 50 /30 tags. (b) Data suggest forms of co-translational and perhaps even co-transcriptional surveillance in bacteria. In E. coli, the RNA helicase, together with the RNase E scaffold domain was shown to bind stalled ribosomes so that the whole of the degradosome can rescue these by degrading the single stranded mRNA substrate. In bacteria RNA decay uses a different approach from eukaryotes, in that appended tags (poly A, 50 monophosphate) are triggers for decay rather than mRNA protection.

co-translationally, as discussed previously, and the Ski helicase complex is pivotal in this cooperation (Figure 4a). In prokaryotes, initial opportunities for tight regulation might occur in the ‘expressome’, a RNAPribosome complex [70] (Figure 4b), or the RNAP-30S complex, which differs in the small subunit orientation and may potentially represent a different state [71]. There is precedence for an interplay between translation and degradation in bacteria as well, in which mRNA lifetime is influenced mostly by the time during which it can support protein synthesis. In E. coli, the RNA degradosome can bind translating ribosomes and degrade Current Opinion in Structural Biology 2020, 61:59–70

the engaged, single stranded, mRNA substrate [72] (Figure 4b). In such cases, mRNA decay occurs as a consequence of translation inhibition and is involved only as a scavenging process [73,74]. Data suggest the bacterial equivalent of the Ski complex, the degradosome associated DEAD-box helicase RhlB, directly interacts with the stalled ribosome in such salvage assemblies (Figure 4b). Recently, an endoribonuclease with a PIN domain, also found in the eukaryotic exosome accessory, was discovered to bind the ribosome A site to cleave transcripts in a ribosome-dependent mRNA decay pathway in the model gram positive bacterium B. subtilis [75]. www.sciencedirect.com

RNA lifetime Dendooven, Luisi and Bandyra 67

Spontaneous compartmentalisation and the un-structural biology of riboregulation Organelles compartmentalise and concentrate the activities of the machines that act on RNA. This can drive forward binding as well as out-of-equilibrium processes supported by catalysis. Spontaneous compartmentalisation of RNA-binding proteins, arising from liquid–liquid phase separation, is proposed to serve a similar function, and could in principle drive ribonucleoprotein assembly and processing [76–82]. RNA–protein interactions can facilitate liquid–liquid phase separation [78], as seen in the occurrence of eukaryotic P-bodies and stress granules that contain mRNA decay machinery and translation initiation factors, respectively [83] (Figures 1a and 2a). In one mode of action, low-complexity regions that are associated with folded RNA-binding domains can become more ordered upon RNA-binding to enhance their affinity for target RNA [84], most likely by lowering off-rates. Another effect of the droplets is to provide a physico-chemical environment that behaves more like an organic solvent rather than water, resulting in destabilisation of nucleic acid helices but promoting other structures [85]. In this way, the droplet can be conducive to specific biochemical reactions.

RNase E core [89,90]. Strikingly, the unstructured character of degradosome machineries is conserved in most bacterial lineages, but not via horizontal gene transfer [91], and therefore must provide evolutionary benefits to the cell. Speculatively, the unstructured RNase E scaffold domain increases degradosome evolvability by relaxing foldability constraints on viable mutations [92]. That these unstructured C-terminal tails seem to play key roles in local, transient compartmentalisation via liquid–liquid phase separation, may pose a strong additional force for these scaffold domains to persist in evolution.

Summary and perspective

One key question is what physico-chemical properties of the peptides and nucleic acids drive the liquid–liquid phase separation. The peptides must be densely packed in the liquid phase and are associated with certain types of natively unstructured domains. Efforts to understand the phase separation have come from analyses of the low complexity domain of the RNA-binding protein FUS (Fused in Sarcoma), which forms liquid–liquid phase separated bodies in vivo and in vitro. Portions of the FUS form amyloid-like fibrils mediated by beta-strand interactions that have been structurally characterised by solid state NMR [86]. This finding supports a model, in which the flexible extensions from the structured core mediate the interactions for liquid–liquid phase separation. However, another recent analysis suggests that the portion that forms the liquid–liquid phase separated property is conformationally heterogeneous [87].

The ability to regulate gene expression quickly and robustly requires that transcripts have a limited lifetime. The association of degrading machineries with ribosomes and potentially with expressomes allows for on-the-fly surveillance of nascent RNA species. Signatures in the transcript can control intrinsic rates of decay and trigger destruction. One puzzle in molecular recognition is how aberrant transcripts are detected and destroyed faster than the non-aberrant forms. Bresson and Tollervey [9] have proposed that, in eukaryotic cells, the identification and degradation of defective RNAs and enormous numbers of spurious transcripts, does not require recognition of specific signatures. Instead, surveillance ensures that transcripts are subject to ‘Decay by Default’, but those that undergo correct and timely maturation acquire features that protect them from a fate of degradation by the decay machinery, such as 50 caps and poly A tails. Thus, RNA polymerase II is in a default surveillance-ready state, and the arrival of protective factors prevents RNA decay. A transcript will be automatically destroyed unless protected, so the problem becomes one of recognition on multiple levels. It is interesting to note in this regard that the degradative Ccr4Not complex interacts with proteins that participate in maturation of mRNPs coupled to their export, such as the nuclear poly(A)-binding protein Nab2 or Hrp1 of the THO complex, or Mlp1 of the inner basket of the nuclear pore complex [27]. These interactions of the Ccr4-Not complex might provide an opportunity for deadenylation of transcripts if the maturation is too slow or defective.

The property of phase separation may also occur in the much smaller cells of bacteria (Figure 1a). The best know example is that of the RNA degradosome from the Gram-negative bacterium Caulobacter aquatic crescentus. This degradosome assembly has natively unstructured scaffold domains that bear RNA-binding regions. The C. crescentus degradosome forms liquid– liquid phase-separated ribonucleoprotein bodies [88

]. Formation of these bodies is dependent on RNA-binding and can be reversed by RNA turnover. A similar phenomenon has been observed for the E. coli RNA degradosome, which is associated with the cytoplasmic membrane and has, just like the Caulobacter degradosome, a long continuous natively unstructured scaffold region in the

Another important factor in RNA lifetime are helicases, which are involved in all stages of the life of an RNA — from biogenesis to turnover. Discriminating defective transcripts must require an input of energy that could be supplied by ATP-dependent RNA helicases, potentially explaining why helicases such as Mtr4 are key components and physical linkers for surveillance pathways [9]. Further understanding the interactions of human MTR4 helicase with partner proteins and RNA exosome substrates will likely illuminate the mechanism of substrate identification and activity [48,49]. Complex roles of the helicase are also becoming apparent, as indicated by the association of UPF1 involved in NMD with mRNAs at transcription [59,93].

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68 Macromolecular assemblies

Compartmentalisation of riboregulatory processes in the form of liquid–liquid phase separation in both eukaryotic and prokaryotic cells allows for local shifts in substrate and enzyme concentrations. These organelle-like bodies can serve as transient microfactories with increased processivity, and they appear to be used in widely diverse species as an effective solution. It is striking in this regard to note that the natively unstructured portion of bacterial degradosomes has been sustained in evolution for at least a billion years, which is a strong indication of their importance in biology.

Conflict of interest statement Nothing declared.

Acknowledgements We are supported by the Wellcome Trust. TD is also supported by an AstraZeneca Studentship. We thank Chris Hill, Alex Borodavka, Marta Kubaska, Mandy Neumann, Stefan Bresson and David Tollervey for helpful comments and discussion.

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