The eukaryotic RNA exosome processes and degrades RNA by directing substrates to the distributive or processive 3′ to 5′ exoribonuclease activities of Rrp6 or Rrp44, respectively. The non-catalytic nine-subunit exosome core (Exo9) features a prominent central channel. Although RNA can pass through the channel to engage Rrp44, it is not clear how RNA is directed to Rrp6 or whether Rrp6 uses the central channel. Here we report a 3.3 Å crystal structure of a ten-subunit RNA exosome complex from Saccharomyces cerevisiae composed of the Exo9 core and Rrp6 bound to single-stranded poly(A) RNA. The Rrp6 catalytic domain rests on top of the Exo9 S1/KH ring above the central channel, the RNA 3′ end is anchored in the Rrp6 active site, and the remaining RNA traverses the S1/KH ring in an opposite orientation to that observed in a structure of a Rrp44-containing exosome complex. Solution studies with human and yeast RNA exosome complexes suggest that the RNA path to Rrp6 is conserved and dependent on the integrity of the S1/KH ring. Although path selection to Rrp6 or Rrp44 is stochastic in vitro, the fate of a particular RNA may be determined in vivo by the manner in which cofactors present RNA to the RNA exosome.
The eukaryotic RNA exosome core is an essential nine-subunit complex formed by two stacked rings of six RNase PH-like proteins on the bottom and three S1/KH domain ‘cap’ proteins on the top. Exo9 associates with the endo- and 3′ to 5′ processive exoribonuclease Rrp44, and the 3′ to 5′ distributive exoribonuclease Rrp6 (ref. 1). In Saccharomyces cerevisiae, the cytoplasmic exosome includes Exo9 and Rrp44 (Exo10Rrp44), the nuclear exosome contains Exo9, Rrp44 and Rrp6 (Exo11Rrp44/Rrp6)2, and a nucleolar exosome with Exo9 and Rrp6 (Exo10Rrp6) has been posited in human3. Each exosome complex may be uniquely equipped to target different RNA substrates, in conjunction with protein cofactors, to catalyse RNA turnover, quality control, or processing in the context of their respective cellular compartments4.
Rrp6 is a RNase D family member5, and is proposed to hydrolyse RNA via two metal ion catalysis6,7. It includes an amino-terminal PMC2NT domain that associates with a cofactor Rrp47 (ref. 8), an exoribonuclease domain (EXO), an HRDC domain, and a carboxy-terminal domain (CTD) that associates with the Exo9 core9. Structural studies revealed that the EXO and HRDC domains constitute the catalytic module10,11 and a Exo10Rrp44+Rrp6Cterm structure revealed how Rrp6 CTD residues interact with Exo9 (ref. 12). No structures yet exist for Rrp6 in complex with Exo9 or RNA.
We proposed that Rrp44 and Rrp6 use an overlapping channel within the S1/KH ring to engage RNA, that Rrp6 activities are modulated by Exo9, and that Rrp6 stimulates Rrp44 in binding and degradation of single stranded RNA in the context of Exo11Rrp44/Rrp6 (ref. 13). To provide a structural basis for these observations, we crystallized Rrp6 with Exo9 and a 24-nucleotide single-stranded poly(A) RNA (poly(A)24 RNA).
Global architecture of Exo10Rrp6–poly(A) RNA
The 3.3 Å structure of Exo10Rrp6 was obtained in the presence of a 24-nucleotide (nt) single-stranded poly(A) RNA using Rrp6 (128–685) that lacked exoribonuclease activity (D238N)13,14, the PMC2NT domain (1–127)13 and its last 48 C-terminal residues (Extended Data Fig. 1; Extended Data Table 1). Rrp6 is positioned atop the Exo9 S1/KH ring with Exo9 subunits resembling those of human Exo9 (ref. 15) and yeast Exo9 (ref. 12) in Exo10Rrp44+Rrp6Cterm (Fig. 1a, b). Poly(A)24 RNA is coordinated within the S1/KH ring with Rrp6 active site residues contacting the RNA 3′ end (Fig. 1c). The Rrp6 EXO domain contacts two of the three S1/KH ring proteins, Rrp4 and Rrp40, and the HRDC domain is proximal to Rrp4, although no direct contacts are apparent. Consistent with previous results12, the Rrp6 CTD wraps around the N-terminal domain (NTD) of Csl4 and PH-like subunit, Rrp43, before emerging at the top of the Exo9 core (Fig. 1a; Extended Data Fig. 2a, b). No electron density was observed for CTD residues 629–684 and for residues (517–524) that link the HRDC domain and CTD. The location of the Rrp6 N terminus (Met 128) places the Rrp6 PMC2NT domain over the Exo9 central channel in an ideal position to interact with Rrp47 to facilitate substrate recruitment (Fig. 1a). Alignment of exosome-associated Rrp6 to the yeast Rrp6 catalytic module (EXO-HRDC) reveals few differences (Extended Data Fig. 2c).
The NTD of Rrp4 and S1 domains of Rrp4 and Rrp40 interact with the Rrp6 EXO domain, burying 2,230 Å2 of surface area in the complex. Rrp6 interactions with the Rrp4 NTD (Region 2) and S1 domain (Region 3) are more extensive (1,750 Å2), whereas interactions with the Rrp40 S1 domain and Rrp4 S1 domain encompass a region proximal to the RNA binding site (Region 1). These surfaces are highly conserved (Fig. 2a, b) in comparison to the Rrp6 CTD (Extended Data Fig. 2a), which is important for Rrp6 interaction with the exosome core as illustrated by analytical gel-filtration studies showing that the CTDs of both yeast and human Rrp6 are required for association with the exosome (Extended Data Fig. 3).
The structure of Exo10Rrp6 bound to poly(A)24 RNA reveals how an RNase D family member interacts with RNA (Fig. 2c). The Rrp6 catalytic domain and active site bind A21A22A23A24 with the 2′-OH and 3′-OH of the terminal A24 coordinated via two main-chain hydrogen bonds to the backbone amide and carbonyl oxygen of His 241 and Glu 240 side-chain carboxylate, similar to that observed for AMP in the Rrp6 catalytic domain10. An additional contact is observed to A24 between the N7 adenine atom and Gln 345 side chain amide. The scissile phosphate of A24 and A23 is coordinated by a single magnesium ion that bridges the phosphate and the side chain carbonyl of Asn 238 (Asp in wild-type enzyme). The scissile phosphate is within hydrogen bond distance of the invariant side chain hydroxyl of Tyr 361 (Extended Data Fig. 4a,c). Consistent with the proposed reaction mechanism7, two metal ions coordinated by conserved side-chain carboxylates in the yeast Rrp6–AMP complex are ideally positioned to contact the scissile phosphate (Extended Data Fig. 4c, d). Several contacts between metal ions and RNA are missing in our structure owing to the D238N substitution.
The remaining contacts to poly(A)24 RNA reveal several non-specific interactions, consistent with the ability of Rrp6 to degrade RNA of any sequence. A23 2′-OH is within hydrogen-bonding distance to the side-chain carboxylate of Asp 296, whereas the A23 phosphate is within hydrogen-bonding distance of the Leu 328 backbone amide. The A23 base is situated over the A24 base with van der Waals contacts provided by side chains of Trp 299, Met 295 and Phe 294. Phe 294 is positioned between the A23 and A22 bases, presumably disrupting base-stacking interactions. The A22 base and ribose are cradled by van der Waals contacts to Phe 294, Gly 292, His 291 and Tyr 315 while its phosphate is within hydrogen bonding distance with the backbone amide of His 291. Lys 319, His 326 and Tyr 315 contribute the remaining contacts to the A21 base. Importantly, none of the aforementioned contacts seem to interrogate the identity of the base. Sixteen of eighteen residues observed in direct contact to the RNA or metal cofactors are conserved in human RRP6 (Extended Data Fig. 4a, b)10,11.
RNA passes through the S1/KH ring
Four out of six nucleotides of poly(A)24 RNA observed in electron density (5′-A19-A24-3′) are coordinated by Rrp6 placing the 5′ end (A19) proximal to conserved, basic side chains from Rrp4 (Lys 122 and Arg 123), while Rrp40 Lys 108 points towards A24 and Arg 110 is proximal to the phosphate of A21 and ribose of A20 (Fig. 3a). Additional densities with spacing consistent with the RNA phosphate backbone, modelled as PO4a and PO4b, are observed within the S1/KH ring nearest to Arg 150 and proximal to Arg 145 and Arg 202 of Csl4 (Extended Data Fig. 1a–c). The aforementioned Csl4, Rrp4 and Rrp40 residues are highly conserved15. No additional density consistent with RNA is observed within the PH-like ring.
Rrp6 and Rrp44 use a shared portion of the central channel to engage RNA substrates13, and alignment of Exo10Rrp6 and Exo10Rrp44+Rrp6Cterm reveals overlapping paths for RNA within the S1/KH central channel (Fig. 3a, b), albeit in opposing directions. A subset of side chains from the S1/KH cap proteins contact RNA in both structures, but none make contacts that would enforce directionality. To determine the importance of the S1/KH ring for RNA degradation, yeast Exo10Rrp6 and Exo11Rrp44/Rrp6 were reconstituted with Csl4 and Rrp40 containing amino acid substitutions for Arg 145, Arg 150 and Arg 202 in Csl4, and for Lys 107, Lys 108 and Arg 110 in Rrp40. These mutations severely diminish Rrp6 activity in Exo10Rrp6 and Exo11Rrp44/Rrp6 as well as Rrp44 activity in Exo11Rrp44/Rrp6 (Fig. 3c; Extended Data Fig. 5a). In contrast, Rrp6 activity is not inhibited by an insertion in Rrp45 that occludes the PH-like ring channel below the S1/KH ring13, although this mutation inhibits Rrp44 activity. Reconstituted human Exo10RRP6 (Extended Data Fig. 3b) reveals similar dependencies as a channel-occluding insertion in RRP41 near the S1/KH ring impaired RRP6 activity, whereas a channel-occluding insertion in RRP45 well below the S1/KH ring failed to inhibit RRP6 (Fig. 3d; Extended Data Fig. 5b). Combining insertions inhibits RRP6 to a greater degree, probably because the RRP45 insertion restricts movement of the RRP41 insertion. These data indicate that the RNA path to Rrp6 depends on the integrity of the S1/KH ring, but does not require the full extent of the PH-like ring central channel. In contrast, at least for yeast Exo11Rrp44/Rrp6, the RNA path to Rrp44 relies on the integrity of central channel throughout both S1/KH and PH-like rings. These data are consistent with contacts observed to RNA in structures of Exo10Rrp6 and Exo10Rrp44+Rrp6Cterm.
To map RNA interactions through Exo9 to Rrp6 and Rrp44, catalytically dead yeast exosome complexes were reconstituted and subjected to long-wavelength ultraviolet crosslinking using 36-nt poly(A) or AU-rich RNA substrates bearing single 4-thioU substitutions 6, 21 or 29 nt from the 3′ end (Fig. 4a, Extended Data Fig. 6a–c). As the 4-thioU probe is moved 3′ to 5′, AU-rich RNA crosslinks to Rrp44, followed by the PH-like ring, and finally to the S1/KH cap in Exo10Rrp44 and Exo11Rrp44/Rrp6, in agreement with previous studies12,13,16,17. In contrast, crosslinks are only observed to Rrp44 in Exo10Rrp44 with poly(A)36 RNA; however, addition of Rrp6 in Exo11Rrp44/Rrp6 results in crosslinks that progress from Rrp44 through the PH-like ring to the S1/KH ring and Rrp6 as the 4-thioU probe is moved 3′ to 5′. The results with poly(A)36 RNA are in line with previous biochemistry showing that Rrp6 stimulates Rrp44 binding to poly(A) RNA in Exo11Rrp44/Rrp6 (ref. 13).
Crosslinking to the Exo10Rrp6 complex reveals RNA contacts to Rrp6 and Rrp4 when 4-thioU is 6 nt from the 3′ end. Although no structural impediment exists to prevent RNA from entering the PH-like ring, poly(A)36 and AU-rich RNA crosslinks are only observed to Rrp4, Csl4 and Rrp40 even when the 4-thioU is positioned 21 or 29 nt from the 3′ end (Fig. 4c; Extended Data Fig. 6). This pattern contrasts with that observed for Exo10Rrp44 and Exo11Rrp44/Rrp6, where crosslinks are observed to PH-like ring subunits when 4-thioU is placed 21 nt from the 3′ end. Crosslinking patterns to human Exo10RRP6 used a 36-nt AU-rich substrate as reported previously13. Similar to results obtained for yeast, ultraviolet-induced crosslinks are observed to the three S1/KH ring proteins and RRP45, but to none of the other five PH-like proteins (Fig. 4b). Ultraviolet crosslinking to complexes with loop insertions mirror results observed in decay assays (Fig. 3d) with diminished crosslinking when a loop insertion is placed proximal to the S1/KH ring (Extended Data Fig. 7). These data indicate that the integrity of the S1/KH ring central channel is important for Rrp6 activity in both human and yeast systems.
The results are consistent with a model in which distinct but overlapping paths guide RNA to Rrp6 or to Rrp44 (Fig. 4c). So how is a path selected? Path selection seems stochastic in vitro because degradation products of Rrp6 and Rrp44 are observed under conditions of limiting enzyme (Fig. 3c) or limiting substrate (Fig. 4d). The distributive mechanism underlying Rrp6 activity suggests repeated substrate binding and release, whereas the processive mechanism used by Rrp44 suggests that it binds and holds onto substrates until completely degraded. Thus at steady state, binding and ultraviolet crosslinking probably reflect the stable interaction with Rrp44 even when Rrp6 is present13 (Extended Data Fig. 6a). Additional evidence for stochastic sampling of the two paths is evident by ultraviolet crosslinking under conditions of slight enzyme excess (Fig. 4d). As predicted based on the distributive and processive mechanisms of Rrp6 and Rrp44, respectively, crosslinked products are observed to Rrp44, Rrp6 and the S1/KH ring proteins at the earliest times, and this pattern is lost once most of the RNA finds its way to the Rrp44 active site.
Structural analysis of the Exo10Rrp6 poly(A) complex suggests at least four potential paths past the S1/KH ring to Rrp6, although paths 1 and 2 seem most likely with respect to electrostatics and conservation (Fig. 4e; Extended Data Fig. 8a). Importantly, these paths are available in Exo11Rrp44/Rrp6 as they do not involve surfaces from the PH-like ring or Rrp44. Modelling Rrp6 onto the Exo10Rrp44 RNA complex shows that the central channel is still accessible and that RNA paths to Rrp6 and Rrp44 are available in Exo11Rrp44/Rrp6 (Fig. 4e; Extended Data Fig. 8b).
A wider channel in Exo10Rrp6
Rrp6 can stimulate Rrp44 binding and decay activities13, a phenomenon readily apparent in crosslinking to poly(A) RNA (Fig. 4a; Extended Data Fig. 6a). These results indicate that Rrp6 enhances RNA access to the PH-like ring and central channel. Structures of Exo10Rrp6 and Exo10Rrp44+Rrp6Cterm were compared to query differences that might account for this activity. Whereas the global architecture of the PH-like ring does not differ (Fig. 5d), the Exo9 channel widens in Exo10Rrp6 through movement of Rrp4, Rrp40 and Csl4 away from the central channel, widening the gap between Rrp4 and Rrp40 S1 domains by ∼4 Å (Fig. 5a–c). Although Exo10Rrp6 and Exo10Rrp44+Rrp6Cterm structures are bound to RNA, the increase in channel width in Exo10Rrp6 might account for Rrp6-mediated stimulation of Rrp44, especially because Rrp44 and Rrp6 do not physically interact. It remains unclear how Rrp6 exerts this change or if an Exo10Rrp44 apo structure differs from its RNA-bound configuration, but it is clear that Rrp6 EXO and CTD are both required for this activity as addition of either element alone is not sufficient to stimulate Exo10Rrp44 when added in trans (Fig. 5e). These data suggest that the Rrp6 CTD is required to bring the catalytic module in proximity to the S1/KH ring, perhaps eliciting channel widening through EXO domain interaction with Rrp4 and Rrp40. Further data will be required to determine if channel widening is a regulated feature of Exo11Rrp44/Rrp6 or if additional cytoplasmic factors elicit channel widening of Exo10Rrp44.
The structure of Exo10Rrp6 shows Rrp6 positioned above the Exo9 S1/KH ring while the Exo10Rrp44+Rrp6Cterm structure shows Rrp44 below the PH-like ring. It is notable that Rrp6 activity is altered and becomes dependent on the S1/KH ring when associated with Exo9 (ref. 13). Although shorter RNAs may be directed to Rrp44 via a channel-independent “direct access” route18, Rrp44 remains highly dependent on the integrity of the central channel throughout both S1/KH and PH-like rings. The dependency on the Exo9 core is remarkable given that both Rrp44 and Rrp6 active sites are exposed to solvent in Exo10Rrp6 and Exo10Rrp44+Rrp6Cterm complexes.
That a similar segment of the S1/KH ring is used to engage RNA in opposing directions suggests that overlapping paths to Rrp6 and Rrp44 may serve to commit the exosome to distributive or processive degradation depending on how a particular RNA substrate is delivered to the exosome. Because the paths overlap, the exosome would be unable to interact with another substrate until completing the task at hand. Whereas path choice seems stochastic in vitro (Fig. 4d), the nuclear cofactors Mpp6 (ref. 19), Rrp47 (ref. 8) and TRAMP20 may bias selection of a particular path to facilitate transitions between editing, processing or degradation activities of the exosome.
Rrp6 can stimulate Rrp44 activities when associated with the Exo9 core, perhaps through widening of the S1/KH ring. Channel gating mechanisms as a point of regulation have been described in many systems including, but not limited to, the proteasome21,22,23,24,25. Whereas many degradation complexes place their catalytic activities in a protected compartment that is accessed by gating a substrate-responsive channel, the RNA exosome appears inside out, with its catalytic subunits located on the periphery of the exosome core. While the functional relevance of this architecture is not yet clear, it is evident that the exosome core can modulate the activities of its catalytic subunits by requiring RNA to pass through distinct elements of the central channel before being processed or degraded.
RNA exosome subunits were expressed, purified and reconstituted as described13,26; see Methods for details. S. cerevisiae Exo10Rrp6exo− (12–14 mg ml−1) was incubated with poly(A)24 RNA (Invitrogen) at 1:1.1 molar ratio before crystallization. The structure was solved by molecular replacement. Mutant exosome subunits were generated by PCR-based site-directed mutagenesis. Wild-type and mutant exosome complexes were assayed for RNA degradation activities as described13. Ultraviolet crosslinking used catalytically-dead Rrp44 or Rrp6 using conditions described13.
Yeast exosome reconstitution and purification
Expression, purification, and reconstitution of recombinant exosome subunits and complexes have been previously described in detail13,26. In brief, the two catalytic subunits (Rrp6 and Rrp44) and nine non-catalytic subunits of the Exo9 core (Rrp41, Rrp45, Rrp42, Mtr3, Rrp46, Rrp43, Rrp4, Rrp40 and Csl4) were cloned as N-terminal 6× histidine Smt3 fusions in pRSFDuet-1 expression vectors (Novagen), and expressed in the Escherichia coli expression strain BL21 (DE3) RIL (Stratagene) either as heterodimers (Smt3–Rrp41/Rrp45, Smt3–Rrp42/Mtr3, Smt3–Rrp46/Rrp43) or as distinct subunits (Smt3–Csl4, Smt3–Rrp4, Smt3–Rrp40, Smt3–Rrp6, Smt3–Rrp44). Generation of the Rrp40 and Csl4 point mutants was performed by PCR-mediated site-directed mutagenesis. For the mutant subunits, cells were grown in shaker flasks at 37 °C in Superbroth (Teknova), induced by cold shock on ice, addition of ethanol to a final concentration of 2% and 0.05 mM isopropyl-β-d-thiogalactoside (IPTG), and grown at 18 °C overnight. For all other proteins, cells were grown in shaker flasks at 37 °C in Superbroth and induced by cold shock on ice and addition of 0.4 mM IPTG, and then subjected to shaking at 18 °C overnight. Cells were collected and lysed as described previously. After high-speed centrifugation to pellet the insoluble material, the supernatant was loaded on a nickel-NTA column (Qiagen) and allowed to flow by gravity. After washing with 350 mM NaCl, 20 mM Tris pH 8.0, 1 mM BME, and 20 mM imidazole, the column was further washed with 350 mM NaCl, 50 mM KCl, 20 mM Tris pH 8.0, 1 mM BME, 2 mM ATP, 10 mM MgSO4 to displace chaperone impurities. Protein was eluted with 350 mM NaCl, 20 mM Tris pH 8.0, 1 mM BME, and 250 mM imidazole and then purified by size exclusion chromatography. All subunits were purified on the Superdex 200 (GE) with the exception of Smt3–Csl4 and Smt3–Rrp40, which were purified on a Superdex 75 (GE). Wild-type and mutant subunits eluted at the same volumes on gel filtration. Only Smt3–Rrp6, Smt3–Rrp44, and Smt3–Rrp42/Mtr3 were subjected to overnight cleavage by the SUMO protease Ulp1 (ref. 29) with another step of purification on the Superdex 200 to remove the 6× histidine Smt3 tags; Smt3–Rrp42/Mtr3 was cleaved by Ulp1 before formation of Exo9 as inclusion of this tag interferes with reconstitution. At this stage, subunits were concentrated to 6 to 12 mg ml−1 and stored at −80 °C. For reconstitution, Exo9 subunits were mixed together as Smt3 fusions, incubated on ice for 30 min, followed by addition of Ulp1 for another 30 min before overnight dialysis. After purification, exosomes were concentrated to 12–14 mg ml−1 and stored at −80 °C in a final buffer of 100 mM NaCl, 20 mM Tris pH 8.0, 0.1 mM MgCl2, 1 mM TCEP.
Human exosome reconstitution and purification
For human, expression, purification and reconstitution strategies of the Exo9 core were identical except Smt3–RRP43/MTR3/RRP42 was expressed as a trimer, Smt3–RRP46 was expressed alone (as described previously15,26), and RRP451–302/RRP41 was purified with a non-cleavable N-terminal hexa-histidine tag on RRP45. RRP6 (EXOSC10, PM/SCL-100) residues 180 to 804 were PCR-amplified from a previously generated full-length expression construct11, cloned into pET28a-Smt3 and transformed into BL21 (DE3) RIL. For expression, cells were grown in shaker flasks to ∼0.6 OD600 nm, and induced overnight at 18 °C with 0.25 mM IPTG in the presence of 2% ethanol. Subsequent purification of human Smt3–RRP6180–804 and reconstitution of HsExo10RRP6 is identical to that described for the yeast system. The purification method for the catalytic region, RRP6180–606, used for core interaction studies has been described previously11. The channel occluding RRP41 and RRP45 mutants were engineered based on those described for S. cerevisiae13, and were generated by PCR to encode electrostatic and steric loop insertions with the primary sequence GTGESEGESES between amino residues Gly 93 and Arg 94 of RRP45 and between Arg 62 and Ala63 of RRP41.
Crystallization and structure determination
Exosome samples were mixed with poly(A)24 RNA (Invitrogen) in a 1:1.1 molar ratio and incubated on ice for 1 h before crystallization. Crystals grew at 18 °C by vapour diffusion in either sitting (Greiner Bio One, Crystalquick) or hanging (Hampton Research, VDX) drop formats, in 7–11% PEG3350, 100 mM MES pH 6.7, 4–15% MPD, typically taking 3 to 5 days to appear. Crystals were harvested within 2 weeks. Partial degradation of RNA (20–24 nt) was observed in washed crystals (Extended Data Fig. 1d). For cryoprotection, well solution was replaced with crystallization buffer augmented with 25–30% MPD and crystals were incubated for three days before harvesting; one day before harvest, trays were transferred to 4 °C. A light polarizer was used to exclude multiple crystals and to identify crystals with single regions, which were broken off and flash frozen in liquid nitrogen for data collection. X-ray diffraction data were collected at the Advanced Photon Source 24-ID-E and 24-ID-C beam lines, and the National Synchrotron Light Source X29 beam line. Data was obtained from a single crystal diffracted at NSLS X29 at a wavelength of 1.075 Å at 100 K. Data were processed using HKL200030 and statistics reported in Extended Data Table 1 were obtained using Phenix31 including CC1/2 (%) and CC* (%) values of 99.9 (33.8) and 100.0 (71.1) for data between 50–3.3 (3.42–3.3) Å. The structure was solved by molecular replacement using Phaser32 and coordinates of human apo Exo9 (PDB: 2NN6) and yeast Rrp6CAT (PDB: 2HBL) as search models followed by docking yeast Exo10Rrp44+6Cterm (PDB: 4IFD) (Extended Data Table 1). The structure is refined to R/Rfree values of 0.227/0.265. The final model includes six nucleotides of the poly(A) 24mer, and two phosphate ions that likely represent the RNA backbone (Extended Data Fig. 1a–c), and 2,751 of the 3,156 amino acids present in the crystal (Extended Data Table 2). The asymmetric unit contains one complex. Iterative rounds of refinement were accomplished using Phenix31. RNA and side chains were manually built using O33 and Coot34. The model was initially refined using secondary structure restraints in conjunction with positional refinement followed by individual B-factor refinement. Secondary structure restraints were released and a final round of positional and B-factor refinement was performed followed by refinement of TLS parameters that resulted in a further decrease in R and Rfree values. Figures depicting the structure were prepared with Pymol27. Surface conservation was calculated using ConSurf35. Structure quality was assessed using MolProbity36 indicating the model has excellent geometry with 95.7% in favoured and 100% in allowed regions of Ramachandran space. The structure also scored in the 100th percentile for the Clash and MolProbity scores.
For yeast exosomes, unless otherwise noted, exoribonuclease assays were performed under multiple turnover conditions as described previously13. In brief, 10 nM synthetic 49-nt RNAs (Invitrogen) bearing 5′ fluorescein labels were incubated at 30 °C with 1 nM RNA exosome for various time points, and RNA degradation was monitored by resolving reaction intermediates by denaturing TBE-PAGE (Invitrogen), and detected using a Fuji FLA-5000 scanner (FITC filter). A similar protocol was followed for human exosome complexes, the exception being that these assays were performed at 37 °C, with 5 nM of the 49-nt RNA and 50 nM HsExo10RRP6. For mix-in experiments (Fig. 5e), twofold molar excess of Rrp6 protein was incubated with Exo10Rrp44 on ice for 1 h before initiating RNA decay by addition of RNA. For steady state ultraviolet-RNA crosslinking to 4-thioU-bearing RNAs, 500 nM of exosome complexes with mutations in the exoribonuclease active sites (D238N for Rrp6, D551N for Rrp44) were incubated for 20 min on ice with 50 nM 36-nt 5′ fluorescein RNAs with a single internal 4-thioU (Thermo Scientific) in a 70 μl reaction volume. Binding buffer includes 50 mM KCl, 20 mM Tris pH 8.0, 10 mM DTT, 0.5 mM MgCl2. Crosslinking was performed by subjecting the RNA-exosome mixture to long-range ultraviolet (365 nm) for 15 min in the dark using a 4 W handheld lamp. For the time course described in Fig. 4d, 350 nM of exosome was incubated with 150 nM of RNA, and the binding reactions allowed to proceed for indicated times before initiating crosslinking for 10 min. 15 μl was quenched with LDS loading buffer, and the crosslinked products were separated by SDS–PAGE (Invitrogen) and visualized with a Fuji FLA-5000 scanner (FITC filter). 4-thioU RNA included the following sequences, each of which contained a 5′-fluorescein: 5′-Fl-AAUUAU4thioUUAUUAUUUAUUUAUUAUUUAUUUAUUUAA; 5′-Fl-AAUUAUUUAUUAUU4thioUAUUUAUUAUUUAUUUAUUUAA; 5′-Fl-AAUUAUUUAUUAUUUAUUUAUUAUUUAUU4thioUAUUUAA; 5′-Fl-AAAAAA4thioUAAAAAAAAAAAAAAAAAAAAAAAAAAAAA; 5′-Fl-AAAAAAAAAAAAAA4thioUAAAAAAAAAAAAAAAAAAAAA; 5′-Fl-AAAAAAAAAAAAAAAAAAAAAAAAAAAAA4thioUAAAAAA. For ultraviolet crosslinking to human exosome complexes, samples were incubated with 250 nM of the AU-rich RNA and protein in binding buffer for 60 min on ice, and then placed in a ultraviolet-Stratalinker (Stragene) and subjected to 300,000 μJ of short-wave (254 nm) ultraviolet radiation. Samples were quenched with LDS loading buffer, and crosslinked products were separated and visualized as described above.
Protein Data Bank
Atomic coordinates and structure factors are deposited in the Protein Data Bank with accession code 4OO1.
We thank NE-CAT beamlines (Advanced Photon Source) supported by P41GM103403 (NIH NIGMS). APS is supported by the US Department of Energy, Office of Basic Energy Sciences, under Contract No. DE-AC02-06CH11357. Beamline X29 (National Synchrotron Light Source) supported by the US Department of Energy, the Office of Basic Energy Sciences and P41RR012408 (NIH NCRR) and P41GM103473 (NIH NIGMS). Research reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under award numbers F31GM097910 (E.V.W.) and R01GM079196 (C.D.L.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. C.D.L. is an investigator of the Howard Hughes Medical Institute.
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Cell Research (2016)