Despite the conservation of the methylation sites, the abundance of site-specific modifications shows variability across species and tissues, suggesting that rRNA methylation may provide a means of controlling gene expression. As all Box C/D RNPs are thought to adopt a similar structure, it remains unclear how the methylation efficiency is regulated. Here, we provide the first structural evidence that, in the context of the Box C/D RNP, the affinity of the catalytic module fibrillarin for the substrate–guide helix is dependent on the RNA sequence outside the methylation site, thus providing a mechanism by which both the substrate and guide RNA sequences determine the degree of methylation. To reach this result, we develop an iterative structure-calculation protocol that exploits the power of integrative structural biology to characterize conformational ensembles. In a wide variety of cellular processes, ranging from biosynthesis to signalling and regulation of gene expression, RNA is chemically modified both co- and post-transcriptionally. All classes of RNA are modified, and RNA processing and editing mechanisms are highly conserved, with more than 140 chemical modifications supporting RNA function in all three domains of life . In rRNARecent evidence shows that, while methylation sites are largely conserved and cluster in functionally important regions of the ribosome , the abuBesides their role in guiding 2’-O-methylation, Box C/D RNPs are involved in a variety of other functions, ranging from rRNA processing .The lack of an in vitro reconstitution protocol yielding an active snoRNP currently precludes mechanistic and structural studies of the eukaryotic Box C/D complex. All structural and in vitro functional work to date has focused on the archaeal Box C/D sRNP . The valIn archaea, Box C/D sRNPs consist of three proteins assembled around the guide sRNA . Within In the absence of substrate RNA (apo form), the archaeal Box C/D sRNP has been found to assemble mainly as a dimeric RNP, comprising four copies of each protein and two copies of the guide sRNA  di-RNP,. Upon saPyrococcus furiosus (Pf) sR26 guide RNA, whose corresponding sRNP methylates substrate D’ more efficiently than substrate D, we demonstrated that methylation levels depend on — among other factors — the nature of the first base-paired nucleotide of the substrate , small angle X-ray (SAXS) and neutron (SANS) scattering data, we demonstrate that, in the context of the sRNP complex, the affinity of fibrillarin for the substrate depends on the RNA sequence beyond the methylation site. This difference in affinity is explained by the energetics of a global conformational transition of the sRNP from an inactive to an active state and provides a further route, besides the modulation of product dissociation described previously , to tunePf sR26 RNP we set out to determine the structure of the corresponding half-loaded sRNPs, bound to either substrate D or substrate D’. We used a stabilized version of the Pf sR26 guide RNA, where the apical K-loop has been substituted by a K-turn sequence , measured by SAXS or SANS. To estimate the compatibility of experimentally determined Rg values with the mono- or di-RNP assembly states, we evaluated the theoretical Rg distributions of 5000 di-RNP models with randomized positions of the fibrillarin copies not bound to the RNA in both apo and holo (fully-loaded) conformations from g value of 55.9 Å with a standard deviation (SD) of 2.0 Å for the apo di-RNP and a mean Rg of 58.1 ± 3.6 Å for the holo di-RNP. The SAXS curves of the apo sRNP assembled with st-sR26 (g) of 54.3 Å, which is consistent with a di-RNP architecture   . These ro state) . The samo state) . This iso state) . The RNPo state) .Before embarking upon the structural study of the sRNPs containing st-sR26, we wanted to understand which elements are responsible for the different oligomerization states of the holo ssR26- and holo st-sR26-RNPs. The ssR26 and the st-sR26 RNAs differ only in the sequence of the guide RNA at the box D position, which in the case of ssR26 is identical to that of guide D’. Thus, we generated two additional guide RNAs with distinct D and D’ sequences, st-sR26-1 and st-sR26-2: in st-sR26-1 (st-sR26-2), guide sequence D is a chimeric sequence, formed by the 5’ half of st-sR26 guide D (st-sR26 guide D’) and the 3’ half of st-sR26 guide D’ (st-sR26 guide D) . Interes2H-fibrillarin in 42%:58% D2O:H2O solvent: the number and relative intensities of the maxima are compatible with the presence of two fibrillarin copies but incompatible with the presence of four  distribution calculated from the SANS curve of the complexes assembled with  of four . As mono of four , the sRNTo investigate whether the difference in methylation efficiency of substrate D and D’ correlates with structural differences, we assembled the Box C/D RNP with the st-sR26 guide RNA and saturated either its D or D’ guide site  to obtai1H-13C correlation spectra of fibrillarin and L7Ae labelled specifically at the methyl groups of Ile, Val and Leu residues . In this technique, a paramagnetic tag (spin-label) carrying an unpaired electron is coupled to a unique cysteine engineered on one protein subunit within the complex. The PREs elicited on the methyl groups of a second protein subunit by the unpaired electron are translated into distance restraints , which d2H-labelled and contribute to the observed scattering signal, while the scattered intensity of the unlabelled proteins is masked by the solvent, which is prepared as a 42%:58% D2O:H2O mixture. A combination of such datasets provides sufficient information to restrain the relative position of several molecules within a multi-subunit complex. In our case we acquired SANS curves for 2H-L7Ae, 2H-Nop5, 2H-Fib, 2H-RNA, 2H-Fib/2H-RNA and 2H(70%)-Nop5/2H-RNA in 42%:58% D2O:H2O  for fibrillarin upon RNA binding should be few and relatively small in magnitude. As expected, the methyl-group NMR spectrum of the substrate-bound RNPs showed only moderate CSPs; nonetheless, these were mainly localized in the spectral region containing V110, V151 and V185, thus confirming that fibrillarin recognizes the substrate D’–guide duplex  or exchanging between the on- and off-states -Nop5/2H-RNA SANS and SAXS curves   . Conversr (>250) .When we fitted the PRE and SAS data with ensembles consisting of both - and -conformers, we could reproduce all experimental data satisfactorily with a population of -conformers of 34 ± 10%  and 5.The combination of the NMR and SAS data demonstrated the existence of a conformational equilibrium between - and -conformers for both substrate D- and substrate D’-loaded RNPs. The ensemble of conformations representing the substrate D’-loaded sRNP  containeS. solfataricus enzyme  simulation. In the -state, both half-loaded RNPs display a regular A-form helix of 11 base-pairs formed by the guide and substrate RNAs and positioned far from the Nop5 coiled-coil domains. The geometry of this helix was given as a restraint in the structure calculations, because of the perfect complementarity of the substrate–guide sequences over these 11 nucleotides. Binding of fibrillarin pushes the substrate–guide duplex towards the Nop5 coiled-coil domain, thereby perturbing the base-pairing at the substrate 3’ end . This obs enzyme . During s enzyme . In addis enzyme  and 7. Fs enzyme . Converss enzyme . The firWe conclude that the stability of the fibrillarin-bound form depends on a delicate balance between the loss of entropy due to fibrillarin localization, and the positive and negative enthalpy changes associated with base-pair melting and formation of new protein–RNA contacts, respectively. Given that:2’-O-rRNA methylation is one of the most extensive modification processes occurring during ribosome synthesis and maturation. The strong conservation of the methylation sites over different species, together with the lethal effect of methylation suppression, led to the conclusion that methylation is a constitutive modification of functional ribosomes. However, rRNA methylation has recently been proposed to exert a regulatory function by generating an heterogeneous ribosome population with differential methylation levels .2’-O-methylation is implemented by the Box C/D RNP enzyme through an RNA-guided catalysis. In addition to methylation, Box C/D complexes are involved in a plethora of other functions related to RNA processing. In the context of the multiple roles of Box C/D complexes, the question arises as to how Box C/D RNPs distinguish whether the RNA substrate bound to the guide sequence should be methylated and to what extent.To address this question, we studied the structure-function relationship of Box C/D RNPs in solution through a combination of NMR and SAS data. Using the archaeal Box C/D sRNP, we determined the solution-state structures of the half-loaded substrate D- and substrate D’-bound mono-RNPs. Instead of a single, well-defined conformational state, we found that the copy of fibrillarin at the substrate-loaded site exchanges between the substrate-bound and unbound states, with the substrate D’-loaded complex displaying a higher population of the methylation-competent -state than the substrate D-loaded RNP. Accordingly, the substrate D’-loaded RNP achieves higher levels of methylation . The exiOur results suggest that the proportion of the methylation-competent complex is subtly tuned by the free-energy difference between the active - and inactive - conformations . Recognition of the RNA ribose by fibrillarin is accompanied by a loss in entropy at the junction between the Nop5-NTD and the Nop5 coiled-coil and between the box C/D (or box C’/D’) RNA elements and the substrate–guide duplex. In addition, upon fibrillarin binding, the substrate–guide helical structure must deviate from the ideal A-form geometry, in order to adapt to the proteins. This is particularly evident at the 3’ end of the substrate, where any base-pair beyond the tenth is melted . These eIn conclusion, methylation efficiency appears to be regulated by a complex interaction network depending on the substrate rRNA sequence beyond the methylation site. We propose that, together with substrate turnover , the abiTo calculate the structural models of the substrate D- and substrate D’-loaded RNPs we developed a novel hybrid structure-calculation protocol that fits a combination of NMR and SAS data to an ensemble of conformations. The application of integrative structural biology approaches is particularly relevant to the detection of inter-domain dynamics of RNP complexes, as the different types of structural data are sensitive to conformational changes in different ways. In this case, the combination of NMR PRE data and SAS data was essential for revealing the equilibrium between RNA-bound and RNA-unbound fibrillarin states. The computational workflow developed here allows interpretation of hybrid structural data in terms of structural ensembles, rather than as individual conformations. The protocol proceeds in a step-wise fashion, where the structural ensemble becomes progressively well-defined while increasing the demand on the quality of the fit between predicted and experimental data. We anticipate the methodology developed here to be generally applicable to modular enzymes undergoing domain reorientation during catalysis.2O M9 minimal medium using 2H-glycerol as the sole carbon source. Deuterated proteins with 1H,13C-labelled ILV methyl groups were produced following protocols developed in the Kay laboratory . Leucine-specific labelling was achieved using the protocol described by 2H-Nop5, and, in the case of 1H,13C -ILV methyl-labelled L7Ae, with both 2H-Nop5 and 2H-fibrillarin. The 2H(70%)-Nop5 sample for SANS experiments was obtained by expression in 100% D2O M9 minimal medium with 1H-glucose as the sole carbon source; deuteration levels for this sample were verified by MALDI mass spectrometry.L7Ae (UniProtKB accession code Q8U160), Nop5 (Q8U4M1) and archaeal fibrillarin (Q8U4M2) were expressed, purified and reconstituted with sRNAs as described previously . Nop5 waboratory . Stereos2H-RNA samples, RNA synthesis was performed using 2H-labelled rNTPs (Silantes).Guide-RNAs were produced by in vitro transcription from double-stranded plasmid DNA templates using T7 RNA polymerase produced in-house and rNTPs (Roth). RNAs were purified by denaturing 12–20% polyacrylamide gel electrophoresis, and extracted by electro-elution. For GCGAGCAAUGAUGAGUGAUGGGCGAACUGAGCUCGAAAGAGCAAUGAUGACGGAGGUGAUCACUGAGCUCGC-3’ st-sR26-1: 5’-CGAGCAAUGAUGAGUGAUGGGCGAACUGAGCUCGAAAGAGCAAUGAUGACGGAGGGGCGAACUGAGCUGCG-3’st-sR26: 5’-CGAGCAAUGAUGAGUGAUGGGCGAACUGAGCUCGAAAGAGCAAUGAUGAGUGAUGUGAUCACUGAGCUGCG-3’ sR26: 5'-GCGAGCAAUGAUGAGUGAUGGGCGAACUGAAAUAGUGAUGACGGAGGUGA UCUCUGAGCUCGC-3’st-sR26-2: 5’-Substrate RNAs for st-sR26 were produced in-house using synthetic DNA oligonucleotides:GCUUCGCCCAUCAC-3’Substrate D′: 5′-GUAGAUCACCUCCG-3’Substrate D: 5′-GUAUCGCCCCUCCG-3’st-sR26-1 substrate D: 5’-GUAGAUCACAUCAC-3’st-sR26-2 substrate D: 5’-13C-HMQC spectra, acquired on ILV and ILVproS-labelled samples, in combination with 3D TOCSY–13C-HMQC spectra and by comparison to the NOEs expected from the fibrillarin structure. The assignment was transferred stepwise from the free fibrillarin to the Nop5-NTD–fibrillarin complex, the Nop5–fibrillarin complex and finally to the full Box C/D complex. For the ILV-labelled Nop5-NTD–fibrillarin complex, we also acquired a 3D NOESY–13C-HMQC spectrum; for all complexes we acquired 13C-HMQC spectra on ILV-labelled, ILVproS-labelled and L-labelled samples. For the ILV-labelled Nop5–fibrillarin complex, pairings of HMQC peaks from the diastereotopic methyl-groups of leucine and valine residues were verified with the assistance of a 3D experiment in which the 1H and 13C resonances of the methyl groups were correlated with the 13C resonances of the directly bonded methine carbon , thereby allowing methyl-pairs to be identified from their common methine resonance. The pulse-sequence for this experiment comprises an out-and-back magnetization-transfer-pathway starting and ending on the methyl protons, using COSY-type transfers between the methyl and methine carbons and constant-time chemical-shift evolution periods for both indirect 13C dimensions.In the free state, fibrillarin methyl resonances were stereospecifically assigned by means of 3D NOESY–2O buffer , prior to concentration with a 10 kDa-cutoff Amicon centrifugal concentrator (Merck Millipore).Mutants were generated following the QUIKCHANGE-XL protocol (Agilent Technologies) and purified in the presence of 5 mM β-mercaptoethanol in order to prevent disulfide bond formation. For L7Ae, the native C68 was mutated to serine prior to the introduction of cysteine residues at other sites. The purified protein was then buffer exchanged into 50 mM NaPi, 500 mM NaCl, pH 6.6 using a HiPrep 26/10 desalting column  and eluted directly into tubes containing a 10-fold molar excess of the 3-(2-iodoacetoamido)-PROXYL radical (Sigma-Aldrich) in the dark. The spin-labelling reaction was allowed to proceed overnight at room temperature. Spin-labelled proteins were used for complex reconstitution; the free spin-label was removed during the gel-filtration step. The final reconstitution step was carried out in 100% D1H spectra of the sRNP upon addition of increasing concentrations of substrate RNA. Sharp peaks began to appear after a 1:1 molar ratio of substrate:guide RNA was reached.All substrate-loaded sRNPs were obtained by addition of 1.25 molar equivalents of substrate RNA. This ratio yields full saturation of the substrate RNA-binding sites of the guide RNA. We verified this by monitoring the appearance of peaks indicative of free RNA (sharp peaks) in one-dimensional 13C-HMQC spectra were acquired on Bruker Avance 800 and 850 MHz spectrometers, equipped with TCI cryoprobes, at 55°C with sample concentrations between 10 and 40 μM (2–8 mg/ml). Diamagnetic spectra were recorded after reduction of the spin-label by addition of ascorbic acid to a final concentration of 5 mM.http://www.ucl.ac.uk/hansen-lab/fuda/) assuming Lorentzian line-shapes. When necessary, overlapped peaks were fitted as groups. The fitted volumes and line-widths were then converted into peak-heights. The heights in the paramagnetic and diamagnetic states were used to calculate the distance between the nitroxide group of the paramagnetic tag and the respective methyl-group (see below).All spectra were processed using apodization with an exponential function in order to preserve Lorentzian line-shapes. Peaks were fitted with the program FUDA (2 rates corresponding to the transverse relaxation rates of 1H single-quantum coherence (R2diaH) and 1H-13C multiple-quantum coherence (R2diaHC) of each individual peak were quantified using the pulse-schemes from the Kay laboratory  and diamagnetic  peak-heights corresponding to known distances within fibrillarin in complexes reconstituted with the Fib-R109C mutant. For L7Ae, we used known distances between the Nop5-CTD and L7Ae in complexes reconstituted with the Nop5-S343C mutant. The ratios of peak-heights were converted into PREs (Γ2), using 2diaHC and R2diaH rates measured for the respective peaks.tHMQC represents the magnetization transfer time in the HMQC sequence (7.6 ms). As this equation is non-invertible, Γ2 was derived by plotting the simulated bleaching ratio, Ipara/Idia, as a function of Γ2 for a given set of diamagnetic rates, with the experimental errors on Ipara/Idia, R2diaH and R2diaHC used to determine the upper and lower bounds of the derived PRE. These PREs were then used as restraints in the protocol developed in the Clore Lab  and ω is the proton Larmor frequency in rad/s. The errors on the distances were again estimated by using the errors in τC, experimental Ipara/Idia ratios and R2 rates to yield upper and lower bounds on a calibration curve. A lower-bound of 10% was used for the errors of Ipara/Idia, as recommended by Battiste & Wagner . A lower-bound of 2 Å was imposed for the errors on the distances in order to account for tag flexibility. Finally, a minimum error of −4 Å was used as lower bound for the distances extracted from the PRE ratios in the calculation of the -structures, to account for the possibility that the methyl group of only one fibrillarin copy is close the paramagnetic tag: in this case, the effective distance of the methyl group of the one fibrillarin copy to the paramagnetic tag would be smaller than the distance calculated from the sum of the two overlapping fibrillarin peaks (one with PRE intensity-ratios < 0.8 and one with PRE intensity-ratios close to 1).For a given value of τIn the structure calculations (CNS), distances were imposed from the nitrogen atom of the nitroxide group of the paramagnetic tag to the carbon atoms of fibrillarin methyl groups. For L7Ae, where stereospecific assignment of LV methyl groups was not available, the distance restraint was imposed to both methyl group carbons with an ‘OR’ statement. For complexes with both fibrillarin copies positioned away from the RNA, the same set of distance restraints was imposed on each fibrillarin copy; for complexes with one fibrillarin copy close to the RNA, distance restraints were imposed with an ‘OR’ statement.g and I(0) values were extracted according to the Guinier approximation using PRIMUS in ATSAS 2.7.5 , as recommended for globular particles.Box C/D sRNPs reconstituted in 50 mM NaPi pH 6.6, 500 mM NaCl were recorded at 40°C and concentrations varying from 0.4 to 5 mg/ml, unless otherwise specified. In most experiments a temperature of 40°C instead of 55°C was used for SAXS measurements due to the difficulty in collecting data with high salt concentrations at the higher temperature. For all measurements, 2 mM dithiothreitol (DTT) was added to mitigate radiation damage. Data collection was performed at the ESRF bioSAXS beamline BM29 with exposure of 10 frames each of 1 s duration. The curves were compared, merged, and the buffer contribution subtracted by the beamline software BsxCube . ForwardAS 2.7.5 . All Rg g values with the mono- or di-RNP assembly states, we evaluated the theoretical Rg distributions of 5000 di-RNP models in both apo and holo conformations from g value of 55.9 Å with a standard deviation (SD) of 2.0 Å; the holo di-RNP showed a mean Rg of 58.1 ± 3.6 Å; the -state of the mono-RNP showed a mean Rg of 44.7 ± 1.4 Å; and the -state of the mono-RNP showed a mean Rg of 48.5 ± 1.7 Å -Nop5/2H-RNA samples were measured in 50 mM NaPi pH 6.6, 500 mM NaCl, 42%:58% D2O:H2O solutions, in order to mask the contribution of the 1H-proteins. The curves corresponding to 2H-L7Ae, 2H-Nop5, 2H-RNA and 2H(70%)-Nop5/2H-RNA were acquired at D22 at the Institute Laue Langevin , with a neutron wavelength of 6 Å. The 2H-fibrillarin and 2H-fibrillarin/2H-RNA curves were acquired at KWS-1 at JCNS  (Germany)  with a nGermany) . All SAN2H-L7Ae in 42%:58% D2O:H2O. Restraints positioning the Nop5-α9’ helix between the two guide regions (from Nop5-K301 and K304 to the phosphate backbone of the nucleotide linking the K-turn and substrate–guide helix) were also used. With this set up, we started an iterative procedure, to generate two lists of PRE-derived distance-restraints compatible with either the - or -state. 500 structures were calculated per iteration. At the end of each iteration, restraint violations were evaluated: restraints violated by more than 10 Å in either set of calculations were classified, eliminated from that particular set, but kept in the other. After 5 iterations, this led to two restraint-lists per sRNP, corresponding to the - and -states of the sRNP.Structures were calculated using an adapted version of the protocol described in With these four sets of restraints (two for the substrate D-loaded and two for the substrate D’-loaded sRNP), four separate runs of torsion-angle simulated-annealing calculations were performed; we generated 2500 structures per run, using the settings described in 2 statistic , Iexp is the corresponding experimental value, N is the number of experimental points, σ represents the experimental error and c is the scaling factor:The fitness of the experimental SAS and PRE data with respect to the calculated structures was assessed by calculation of the χtatistic  and by v2 distribution of the 2H-Nop5, 2H-L7Ae and 2H-RNA SANS curves. The SAS curves including the contribution from fibrillarin were left out, because we expected the position of fibrillarin to be variable when it is not in contact with the RNA. SAS fitness was calculated with the programs CRYSOL and CRYSON, from the ATSAS suite, version 2.7.5 , the top 50% by 2H-L7Ae fitness , and the top 80% by 2H-Nop5 fitness ; for the substrate D-loaded complex, we selected structures within the top 90% ranking by 2H-RNA fitness , the top 80% by 2H-L7Ae fitness  and the top 90% by 2H-Nop5 fitness . The average pair-wise RMSD of the structures of each ensemble, calculated over the Cα and P atoms of the protein and RNA structured domains, including the fibrillarin units not bound to the RNA, was below 5 and 7 Å for the  and  conformers, respectively, with a maximum RMSD value of less than 10 Å in all cases.The structures ranking in the top 2% in both total energy and restraint energy were selected. To further narrow down the selection on the basis of the SAS data, we evaluated the χon 2.7.5 . Based o2 distributions for each run and curve, whereby we allowed more structures to be selected when the χ2 distribution was flat.Among the selected structures of each of the four runs , the one with the lowest restraint-violation energy that maintained the correct RNA topology was chosen as the starting point for refinement in Cartesian space. The four refinement runs comprised 1500 structures each spanning up to 10 Å RMSD of Cα and P atoms relative to the starting structure . At the end of the refinement, we applied stringent selection criteria with respect to the SAS curves and loose criteria with respect to the energy. The cut-offs for the SAS data were set upon visual inspection of the χ2H-RNA ; top 67% for 2H-L7Ae ; top 33% for 2H-Nop5 ; top 10% for 2H(70%)-Nop5-RNA . Applying these criteria we selected 1 structure for the substrate D’-loaded -state and 12 structures for the -state. The -state structures displayed an average RMSD of 2.4 Å, calculated on all Cα and P atoms ; 66% for 2H-L7Ae ; 33% for 2H-Nop5 ; 10% for 2H(70%)-Nop5-RNA . The final ensembles for the substrate D-loaded - and -states consist of 3 and 20 structures, respectively, with a Cα and P RMSDs of 4.6 and 2.4 Å, respectively.For the substrate D-loaded sRNP the cut-offs are as follows: top 33% of restraint-violation, van der Waals and total energy; top 83% for Representative structures in the final ensembles were minimized in explicit water using Amber 14 and the corresponding Amber99SB force field .2H-fibrillarin, 2H-fibrillarin/2H-RNA SANS and SAXS curves were therefore fitted to a mixture of structures in the - and -states.The PRE data and the SAS data indicated the presence of a conformational equilibrium between the - and -states, as discussed in the main text. The In order to address the flexibility of the Nop5-NTD–fibrillarin modules not in contact with the RNA, we sought to generate ensembles containing different orientations of these modules that would improve the fit to the SAS curves. This conformational diversity is in addition to the equilibrium between the - and -states, resulting in a pool of structures containing both - and - states and multiple conformations of Nop5-NTD–fibrillarin modules in each state.To generate these ensembles we proceeded as follows. Starting from the representative structure of each ensemble of In a separate run comprised of 300 structures, the template structures were kept entirely rigid while the spin-label side-chains were allowed to rotate in order to generate different orientations, as multiple conformations of the spin-label have been demonstrated to fit the PRE data more accurately than a single conformation .Ensemble scoring was carried out for substrate D’- and substrate D-loaded sRNPs via the pseudo-genetic algorithm shown in 2 value with respect to the experimental data was calculated by OLIGOMER -Nop5/2H-RNA and SAXS. The normalized χ2 values for each curve were then summed and renormalized into a single value, obtained with the same The theoretical scattering curve of the ensemble was computed as the linear combination of the scattering curves of each individual conformer . The χOLIGOMER . The norpara/Idia ratios from mixed - and -state ensembles requires an estimation of the timescale of the exchange rate kex (kex = k1 + k-1) between the - and -conformers. This can be easily done by inspecting the ILV-methyl 1H-13C spectra of fibrillarin: in the case of slow conformational exchange, the methyl groups in the fibrillarin copy sampling the on- and off-states should each yield two separate NMR peaks, while for fast conformational exchange these methyl groups should each show only a single peak, at a position corresponding to the population-weighted average of the positions corresponding to the on- and off-states. To investigate this, we used the spectrum of the RNP assembled with ssR26 and loaded with substrate RNA as a reference for the slow-exchange situation: in this complex, two of the four fibrillarin copies adopt a stable on-state, while the other two are in the off-state, and a subset of the fibrillarin methyl groups show separate and resolvable peaks corresponding to the two states. In the spectra of the half-loaded st-sR26 RNPs we did not detect any peak at the positions corresponding to RNA-bound fibrillarin in the holo ssR26 RNP spectra, indicating that in the half-loaded mono-RNP, either the kex is faster than the differences in the resonance frequencies of the fibrillarin methyl groups in the on- and off-states (~40–100 Hz), or the population of the on-state is too small to be detected. In the second case, one would expect no CSPs upon substrate RNA binding, which does not correspond with the observed spectra . The resulting ΓThree independent runs of the scoring algorithm were performed for substrate D’- and substrate D-loaded sRNPs, with the top scoring ensemble, judged by the consensus PRE-SAS score, displayed in After this selection, the conformations of each individual tag were refined by generating additional 3000 conformers per tag and by using the same pseudo-genetic algorithm to select the ensembles of tag conformations that best fitted each individual PRE dataset. During this refinement step the positions of all proteins and RNA, as well as the populations of fibrillarin conformers in the ensemble, were left invariant, in order not to alter the fit to the SAS data.Molecular dynamics simulations of the substrate D’- and substrate D-bound structures representing the -states were carried out in AMBER 2018 . The sim In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.Acceptance summary:A relatively large macromolecular RNA protein complex, Box C/D RNP, was known to methylate ribosomal RNA at ribose 2'-OH, but how this activity is regulated has been poorly understood. The authors show here that the substrate RNA sequence outside of the methylation site can impact C/D RNP mediated interactions and rRNA methylation. This work provides a strong example of the power of using multiple biophysical methods in addressing important structural biology problems, as well as a compelling example of the role of NMR in such studies.Decision letter after peer review:eLife. Your manuscript has been reviewed by two experts and both they and I are impressed with the range of advanced methodologies that has been used to study such a large complex by NMR. We also feel that this can be an important contribution should you and your colleagues address the concerns listed below. Please note that further extensive experiments are not required, but that conclusions must be strongly supported by experiment.Thank you for submitting your manuscript, "The guide sRNA sequence determines the activity level of Box C/D RNPs" for consideration in The reviewers, which have opted to remain anonymous, have discussed the reviews with one another, and the Reviewing Editor has drafted this decision to help you prepare a revised submission.Major1) Previous work by the group described a di-RNP complex in the presence of substrate (holo di-RNP); this work describes a mono-RNP complex in the presence of substrate (holo mono-RNP). The authors mention this but don't provide details. The questions that arise are: a) why are the previous and the present complexes different, is there a specific RNA contact that drives the formation of the di-RNP? and b) in case small differences in the guide RNA have a pronounced effect on the fully assembled complex, what stoichiometry do the authors expect for the Box C/D complex with the native guide RNA?2) The substrate RNA that the authors use is significantly shorter than the natural substrate RNA (rRNA is long). Are the presented structures compatible with much longer substrate RNAs and is the proposed mechanism between the methylation efficiency of D' and D substrates also valid in the context of longer substrates? Can the terminal bases of the 3' and 5' ends of the substrate RNA that is used here adopt the same conformation when extra nucleotides are present?3) The authors should be more accurate regarding the description of the RNA that they used. In the text and Materials and methods, the authors write that they use a stabilized version of the sRNA . In addition, Figure 1B displays two sRNA sequences (the WT and the stabilized one), the legend refers to this as "Sequence of the sRNAs used…" We assume the authors mean that the lower one of the two is the used sRNA sequence. Likewise, Figure 2 refers to a ssR26 species. We assume that this is the ss-sR26 RNA, the RNA that was used in a previous study. Such inaccuracies often cause confusion regarding the type of the RNA that is used. One suggestion is to add this info to the cartoons in Figure 1A, where cartoon of the di-RNP-apo, mono-RNP-holo and di-RNP-holo complexes are shown.4) "In the di-RNP complex, the methyl-group NMR spectra indicated the presence of fibrillarin in two states, one close to the substrate-guide duplex (on-state) and one far from the substrate (off-state) ." We don't see this. In Figure 2A, the NMR spectra of the complex assembled with ss-sR26  and the complex assembled with st-sR26  are overlaid. According to SAXS data these are both di-RNPs. However, we don't see any sign of the presence of two states in those NMR spectra. In addition, later in the manuscript, the authors conclude that the RNP is dynamic in the presence of substrate, whereas the displayed spectra are from complexes in the absence of substrate.5) "In contrast, both half-loaded mono-RNPs (substrate D- bound and substrate D'-bound) show only one set of fibrillarin resonances; however, the peaks that are split in two in the di-RNP are noticeably elongated in the mono- RNP, suggesting a conformational equilibrium between on- and off-states interchanging at a rate that is fast compared to the corresponding difference in NMR frequencies." Again, we are not following this:a) In Figure 2B, we see one resonance for the st-sR26 apo complex  For the st-sR26 holo complex  (is this after addition of both D and D' or after either D or D'?), we see one maybe slightly broadened resonance. For the ssR26-holo  complex one can imagine two resonances . This could suggest that the holo complexes (mono- or di-RNA) are actually sampling multiple states. The text, however, mentions that the half-loaded mono-RNPs only show one set of resonances.6) We also don't fully understand the statement in the legend of Figure 2B "a single but elongated peak, characteristic of one copy of fibrillarin being in fast-exchange between the two states", refering to the green contours from the mono-RNP holo complex. In case there is fast exchange between 2 states, the green resonance should appear between the two purple resonances. In our view, this is not the case; the broad resonance appears exactly at one of the two signals that form the purple "doublet". Also, the broadening of the green resonance appears to be mainly in the carbon dimension, and not in the proton dimension. Are the spectra recorded with exactly the same experimental settings, or is it possible that differences in processing, viscosity, temperature or acquisition time is the cause of the differential linewidth? In summary, the conclusion regarding the dynamics of the complex seems not supported by the displayed NMR spectra.7) Does Figure 2C belong to the st-sR26 holo complex in Figure 2B? If yes, this should be clearly indicated.para/Idia be (significantly) larger than 1? For readability, please label in the figure that the blue curve is for D' bound and that the red one is for D bound and indicate that this is the mono-RNP holo complex.8) Figure 2D, top. How can Ipara/Idia values depend on the excess of the D or D' RNA? Are the binding sites for D and D' in both cases fully saturated? We assume that less saturation results in more mobility, so it is important to ensure that one compares fully saturated complexes.9) Do the I10) For Figure 2D, bottom, the deviations between the experimental and simulated P(r) values are similar for the short distances (0 – 40 A) and for the large distances (70 – 140 A). The deviations for the small r values are independent of conformational changes and these are thus intrinsic to the method. The question is, if the deviations at larger r values are thus significant enough that one can conclude from these SANS data that there are two states? If yes, does this agree with the population ratio of the  and  states that the authors determined later in the manuscript?11) Subsection “The conformational ensembles of the half-loaded mono-RNPs in solution” paragraph three: "show a reasonable similarity". The authors need to provide at least an RMSD and an overlay of their two structures with the structure of Lin et al. to support this statement.12) In context of sRNP complex, the affinity of fibrillarin for substrate depends on RNA sequence beyond recognition site– It is unclear what the recognition site is. Referring to substrate sequences? Without a clearly defined recognition site, this claim is not supported by the data– Do the authors mean methylation site rather than recognition site as specified in the Discussion? If this is the case, the Abstract/Introduction should be modified to reflect this level of detail.Life. The paper has been re-evaluated by the reviewers. The reviewers feel that the manuscript is much improved over the initial submission but bring up a number of critical points that must be addressed properly should the manuscript be accepted for publication.Thank you for submission of a revised version of your manuscript "The guide sRNA sequence determines the activity level of Box C/D RNPs" to eIn short, there are serious questions about the NMR data. For example, it is extremely difficult to establish that a dynamic equilibrium is occurring between on/off conformations for one of the copies of fibrillarin from the CSP data. Second, it would be useful to show that the PRE data are inconsistent with the off state and presumably also with the on state. The authors attempt to say this and refer to Figure 2C but it is not possible to see much from this figure, as it is small. It would be important to show the structures of the on and off state along with the PRE values and establish that the PRE values can simply not be explained by a static entity. We suggest separating the on and off structures pictorially to make the point. Second, given that some of the PREs are > 1 it is important that they be validated. One way of doing this is to establish that intra-protein PREs  are consistent with expectations based on distances.Major commentsPrevious points #4, 5 and 6; Legend Figure 2B: "Despite the absence of peak splitting, peak budding is visible in the spectrum of the substrate D'-loaded st-sR26 RNP in all instances for which peak splitting or budding is observed in the spectrum of the holo ssR26 di-RNP."1) This figure and its legend have been remade for this version of the manuscript. Nevertheless, we cannot follow what the authors see in their spectra. The only instance, where we can see a peak splitting (not sure if "budding" is a term that can be used), is in the purple (ssR26 holo) spectrum for residue L58 and, with some imagination, for I62. For all other panels and all other complexes, we fail to see any change in the resonance. In other words, for V118, L160, V141 and L200, there are effectively no chemical shift perturbations that are larger than 0.01 ppm in proton. In carbon there is no CSP whatsoever. For the open-closed conformations, one would expect many and large CSPs.This is a very important point, as the authors have little convincing NMR data that show that the complex exists in two states with fibrillarin either in the on- or the off-state.para/Idia still are very problematic. It is physically not possible that this ratio is larger than 1. In the plot, ratios that are 1.25 are shown. The authors write in their letter that there are only two values over 1, in the plot there appear to be at least 6. The remark that it is hard to extract reliable data for crowded spectral regions is correct, however, in case the data are unreliable or highly uncertain (which is not reflected in the presented error bars), it is not possible to draw solid conclusions. So how do the authors come to their conclusions based on the presented data? As an example, a ratio of 0.75 is mentioned in the text as highly relevant for the closed conformation, whereas a ratio of 1.25 is rebutted  as resulting from difficult spectral interpretations.2) Previous point #8; Figure 2C. The PRE data with an IAs the NMR data do not agree with SANS data, the authors involve conformational rearrangements between open and closed conformations. Clearly, in case things are not in agreement, it can always be explained by dynamics. Because of the very weak NMR evidence (2 points above), it seems a far stretch to conclude that the complex undergoing open-closed motions.Previous point #10; Figure 2D. The figure is still hard to understand. The substrate D loaded experimental (red) and theoretical (brown) are very different. The substrate D' loaded experimental (cyan) and theoretical (blue) are also very different.eLife. Your revised article has been evaluated by Philip Cole (Senior Editor) and a Reviewing Editor.Thank you for resubmitting your work entitled "The guide sRNA sequence determines the activity level of Box C/D RNPs" for further consideration by Thank you for your letter addressing the comments that were raised based on the second revision and you are invited to resubmit a revised manuscript. Let me first address whether it is worth submitting a further revised paper. I agree that it is a lot of work on all sides – but there was very significant enthusiasm from the reviewers and me, and I think that we would all like to see some final issues clarified.eLife is that space is not nearly as constrained as in some other prestigious journals. I would suggest taking advantage of this. For example, include validation of the PREs (intra) for both fibrillarin and L7Ae, as a main figure, to convince the reader that your data are good (I believe that this will placate the reviewers). Then if you fit your inter-data to 1 structural model you should be able to show that the fit is worse. We request that you do this, as then you establish that there could be more than 1 structure. We request that you also include structural models of the sort in Figure 2C but larger so that these can be better viewed. Following this up with the SAXS data will then make your thought process transparent. As the reviewers had difficulty with the SAXS please make it very clear that the disagreement is a really powerful argument for why there are dynamics.First, when there are issues raised by the reviewers that seem unreasonable to the authors, this sometimes reflects the fact that the material was not explained as clearly as might be possible. One of the advantages of We suggest that the NMR CSPs are less convincing because they are small. We recommend that you might emphasize this before the PREs and SAXS  but tone it down a bit as the differences in shifts are not large. One can sort of imagine that the green peaks are between blue and purple but we are talking about very small changes. If you say this as a follow-in to the other data, we think that it may sit easier with the reviewers.Finally, regarding the errors in PREs. we can appreciate that some values could well be > 1 given statistics. One would hope that the errors would reflect this. For a couple of cases that we count, this is not the case. Perhaps if you show more PRE data this will help – and as you will hopefully be showing correlation plots assuming 1 state and that these are not as good as the intra-validation – showing more raw data would be justified. Major1) Previous work by the group described a di-RNP complex in the presence of substrate (holo di-RNP); this work describes a mono-RNP complex in the presence of substrate (holo mono-RNP). The authors mention this but don't provide details. The questions that arise are: a) why are the previous and the present complexes different, is there a specific RNA contact that drives the formation of the di-RNP? and b) in case small differences in the guide RNA have a pronounced effect on the fully assembled complex, what stoichiometry do the authors expect for the Box C/D complex with the native guide RNA?We thank the reviewers for this question. We have thoroughly investigated this point during the course of this work, as the discovery that the complexes assembled with the sR26 and st-sR26 RNAs form monomeric species upon substrate binding was unexpected.With regard to point b): RNPs assembled with both sR26 and st-sR26 RNAs transition from the di-RNP to the mono-RNP form upon substrate binding. To demonstrate this, we have added Figure 1—figure supplement 4, which shows the SAXS curves of apo and holo-complexes assembled with sR26.With regard to point a): our experiments revealed that, while the apo complex is always a di-RNP, the oligomerization state of the holo complex depends on the sequence of the guide RNA in the substrate recognition regions. In particular, the sequences directly upstream of either box D or box D’ play a fundamental role . Unfortunately, because of the lack of high-resolution data for the disordered parts of the RNA components of the complex, we do not yet know why this is the case and cannot propose a predictive set of rules. We are planning solid-state NMR experiments to answer this question. However, these experiments have never been tried before and are likely to require substantial experimental effort.Following the question of the reviewers we have now added new experimental data  and an entire new paragraph to address this point.“Before embarking upon the structural study of the sRNPs containing st-sR26, we first wanted to understand which elements are responsible for the different oligomerization states of the holo ssR26- and holo st-sR26-RNPs. The ssR26 and the st-sR26 RNAs differ only in the sequence of the guide RNA at the box D position, which in the case of ssR26 is identical to that of guide D’. Thus, we generated two additional guide RNAs with distinct D and D’ sequences, st-sR26-1 and st-sR26-2: in st-sR26-1 (st-sR26-2), guide sequence D is a chimeric sequence, formed by the 5’ half of st-sR26 guide D (st-sR26 guide D’) and the 3’ half of st-sR26 guide D’ (st-sR26 guide D) . Interestingly, the Box C/D enzyme containing st-sR26-1 maintained the di-RNP architecture upon binding of either substrate RNAs, while the sRNP comprising st-sR26-2 transitioned to the mono-RNP state . Mutation of the last nucleotide of st-sR26-1 guide D to either C or U (A61C and A61U with complementary substrate D) did not perturb the di-RNP architecture . We conclude that the guide sequence strongly influences the oligomerization state of the holo complex. “2) The substrate RNA that the authors use is significantly shorter than the natural substrate RNA (rRNA is long). Are the presented structures compatible with much longer substrate RNAs and is the proposed mechanism between the methylation efficiency of D' and D substrates also valid in the context of longer substrates? Can the terminal bases of the 3' and 5' ends of the substrate RNA that is used here adopt the same conformation when extra nucleotides are present?Both the 3’ and 5’ ends of the substrates used in this work are not involved in base-pairing and can be easily elongated without perturbing the structure of the Box C/D RNP. We use substrates that are longerthan the 11 base-pairing nucleotides, in order to account for the longer natural substrates.There are several reports in the literature stating that longer RNAs are methylated more efficiently in vitro, probably due to additional interactions within the Box C/D RNP, which however remain uncharacterized. rRNA methylation partially occurs during transcription; thus, another question one can ask is whether the sequences flanking the methylation regions at more than 5–6 nucleotides distance are involved in interactions with other proteins or other RNA stretches and thus unavailable for additional interactions within the Box C/D RNP. We do not know the answer to these questions. There is currently no good model to test the effect of the longer rRNA sequences in vivo.3) The authors should be more accurate regarding the description of the RNA that they used. In the text and Materials and methods, the authors write that they use a stabilized version of the sRNA . In addition, Figure 1B displays two sRNA sequences (the WT and the stabilized one), the legend refers to this as "Sequence of the sRNAs used…" We assume the authors mean that the lower one of the two is the used sRNA sequence. Likewise, Figure 2 refers to a ssR26 species. We assume that this is the ss-sR26 RNA, the RNA that was used in a previous study. Such inaccuracies often cause confusion regarding the type of the RNA that is used. One suggestion is to add this info to the cartoons in Figure 1A, where cartoon of the di-RNP-apo, mono-RNP-holo and di-RNP-holo complexes are shown.We apologize for this. We have now added the ssR26 RNA sequence to panel B of Figure 1. In addition, we have changed the figure caption to:“Two RNA sequences (st-sR26 and ssR26) were derived from the Pf sR26 RNA and used to assemble the Box C/D sRNPs either in this (st-sR26) or previous studies ). The sequence of st-sR26 is derived from the native sR26 RNA upon substitution of the apical K-loop element with the more stable K-turn element. This substitution does not affect the oligomerization state of the complex, as shown in Figure 1—figure supplement 4, but ensures the stability of the complex over several days.”This text explains why we used the st-sR26 RNA instead of sR26 and also addresses the question raised by the reviewers with respect to the oligomerization state expected for the complex assembled with the sR26 RNA.4) "In the di-RNP complex, the methyl-group NMR spectra indicated the presence of fibrillarin in two states, one close to the substrate-guide duplex (on-state) and one far from the substrate (off-state) ." We don't see this. In Figure 2A, the NMR spectra of the complex assembled with ss-sR26  and the complex assembled with st-sR26  are overlaid. According to SAXS data these are both di-RNPs. However, we don't see any sign of the presence of two states in those NMR spectra. In addition, later in the manuscript, the authors conclude that the RNP is dynamic in the presence of substrate, whereas the displayed spectra are from complexes in the absence of substrate.We apologize for having generated confusion with our typo. The sentence should have made referenceto Figure 2B and not 2A. As the reviewers and the editor correctly noticed, there is no evidence of multiple fibrillarin states in the apo form of the complexes. The apo complexes assembled with either ssR26 or st-sR26 are both di-RNPs, with all four copies of fibrillarin far from the RNA due to the absence of the substrate RNAs. The two spectra of Figure 2A  overlap well with each other and indeed are not expected to differ. We have corrected the typo, re-designed Figure 2, substantially changed the main text and the figure caption to provide answers to this as well as to the next points. With respect to this point, the figure caption now reads:1H-13C spectra of fibrillarin in the apo ssR26 (turquoise) and apo st-sR26 (blue) RNPs. In both di-RNPs, all four fibrillarin copies are distant from the RNA and the two spectra are identical.”“Overlay of ILV-methyl 5) "In contrast, both half-loaded mono-RNPs (substrate D- bound and substrate D'-bound) show only one set of fibrillarin resonances; however, the peaks that are split in two in the di-RNP are noticeably elongated in the mono- RNP, suggesting a conformational equilibrium between on- and off-states interchanging at a rate that is fast compared to the corresponding difference in NMR frequencies." Again, we are not following this:a) In Figure 2B, we see one resonance for the st-sR26 apo complex  For the st-sR26 holo complex  (is this after addition of both D and D' or after either D or D'?), we see one maybe slightly broadened resonance. For the ssR26-holo  complex one can imagine two resonances . This could suggest that the holo complexes (mono- or di-RNA) are actually sampling multiple states. The text, however, mentions that the half-loaded mono-RNPs only show one set of resonances.1H and the 13C dimensions and always appear in the same direction of the peak splitting or budding observed in the holo ssR26 di-RNP.The green spectrum of Figure 2B referred to the fully loaded st-sR26 RNP, which has a complicated behavior, as both fibrillarin copies are in equilibrium between the RNA-bound and the RNA-free states. The relative populations of the RNA-bound forms of the fibrillarin at the substrate-loaded site are different for the two different half-loaded mono-RNPs; in addition, the chemical shifts of the respective RNA-bound states will also show differences, as the substrates have different sequences (D and D’). By showing the holo form of the st-sR26 complex we have generated much confusion, especially as we did not explain the spectrum. In this revised version we have re-designed the representation of the spectra in Figure 2: we now show the spectrum of the substrate D’-loaded mono-RNP, which is directly comparable to that of the holo ssR26 di-RNP, as the substrate sequences are the same. We also show the expanded view of more methyl group peaks to show that peak “budding” effects are present in both the We have also added two paragraphs in the main text to explain our interpretation of the peak shape. However, we would like to stress that the presence of the conformational equilibrium is not proven by the minuscule peak budding in the HMQC spectra, but rather by the incompatibility of the SANS and PRE data. Thus, by comparing the HMQC spectra, we do not want to prove the existence of the conformational equilibrium, but rather verify that the peak shapes are compatible with the presence of this equilibrium. Here, we summarize the explanation that we give in the text:1) In the apo complexes all fibrillarin copies are distant from the RNA in both the st-sR26 and ssR26 RNPs. The spectra of the two species overlap perfectly, as shown in Figure 2A, left.2) In the holo di-RNP complex assembled with the ssR26 RNA (our previous work), all structural data are compatible with two fibrillarin copies being in stable contact with the RNA and two being far from the RNA. Two distinct NMR peaks are observed for some of the fibrillarin methyl groups, while many other peaks showed substantial asymmetric broadening (“budding”). Thus, the position of the second peak in the di-RNP HMQC spectrum is representative of fibrillarin bound to the substrate D’–guide duplex, as all substrate-binding sites in this di-RNP recognize substrate D’.3) In the mono-RNP half-loaded with substrate D’, the spectra appear to show only one set of peaks. However, the peaks are elongated (“budding”) in the direction of the second peak seen in the di-RNP spectra. The half-loaded mono-RNP bound to substrate D’ contains two fibrillarin copies. One fibrillarin copy never binds the RNA, as its corresponding substrate (in this case substrate D) is absent. Thus, its methyl group peaks are expected to appear at the same positions as in the apo complex. The other fibrillarin copy may bind the RNA. If this fibrillarin copy were stably bound to the RNA, we would see peaks at a similar position as the second peaks in the holo ssR26 di-RNP spectrum, but this is not the case. If this fibrillarin copy were rapidly exchanging between RNA-bound and -unbound forms, we should see a peak halfway between the two peaks seen in the holo ssR26 di-RNP HMQC spectrum, assuming that the population distribution is 50%:50% and that the line-widths of the two states are similar. This peak, however, would overlap strongly with the peak of the first fibrillarin copy, which stays in the unbound form, because of the modest chemical shift differences between the RNA-bound and RNA-unbound states (see spectra of the di-RNP) as well as the large line-widths. Thus, the observed peak-shape originates from the overlap of the peak of one exchanging fibrillarin molecule with the peak of one non-exchanging fibrillarin molecule in the RNA-unbound state. We have now provided more examples of peaks in Figure 2B and simplified the spectral overlay to allow the reader to appreciate this.6) We also don't fully understand the statement in the legend of Figure 2B "a single but elongated peak, characteristic of one copy of fibrillarin being in fast-exchange between the two states", refering to the green contours from the mono-RNP holo complex. In case there is fast exchange between 2 states, the green resonance should appear between the two purple resonances. In our view, this is not the case; the broad resonance appears exactly at one of the two signals that form the purple "doublet". Also, the broadening of the green resonance appears to be mainly in the carbon dimension, and not in the proton dimension. Are the spectra recorded with exactly the same experimental settings, or is it possible that differences in processing, viscosity, temperature or acquisition time is the cause of the differential linewidth? In summary, the conclusion regarding the dynamics of the complex seems not supported by the displayed NMR spectra.13C dimension. In addition, the clearer display of the peaks should allow better appreciation of the differences in the line-shapes of the peaks of the apo versus the substrate D’-loaded st-sR26 RNPs, which are consistent with (but do not prove) the conformational equilibrium.Please see the answer to the question above, as well as the new Figure 2. The spectra of the apo and substrate D’-loaded sRNPs were acquired and processed in exactly the same way . The display of additional peaks in Figure 2B now demonstrates that the “budding” does not occur exclusively in the The difference between “budding” peaks (those belonging to methyl groups which are split or budding in the holo di-RNP) and non-budding peaks (those belonging to methyl groups which are NOT split or budding in the holo di-RNP) can be appreciated from the lower leftmost panel of Figure 2B, where L160  shows budding upon addition of substrate D’ to the st-sR26 RNP, while L66 does not. This comparison also demonstrates that the budding is not an artifact generated by different experimental or processing parameters. We have now commented on this in the figure caption.7) Does Figure 2C belong to the st-sR26 holo complex in Figure 2B? If yes, this should be clearly indicated.We have amended the figure and figure caption accordingly.para/Idia be (significantly) larger than 1? For readability, please label in the figure that the blue curve is for D' bound and that the red one is for D bound and indicate that this is the mono-RNP holo complex.8) Figure 2D, top. How can Ipara/Idia are larger than 1 (between 1.1 and 1.2). We often encounter these cases when quantifying PREs, due to errors in line-shape fitting as a consequence of spectral overlap and due to noise. The quantification of the Ipara/Idia values in regions of spectral overlap may be difficult. To address this problem, we exclude the more overlapped peaks from the analysis and we apply a generous error-range on the extracted distances .Only two values of IWe have amended the figure according to the reviewers’ suggestion.para/Idia values depend on the excess of the D or D' RNA? Are the binding sites for D and D' in both cases fully saturated? We assume that less saturation results in more mobility, so it is important to ensure that one compares fully saturated complexes.9) Do the Ipara/Idia ratios do depend on the saturation of the D and D’ sites as they are different for the apo and holo states and for the half-loaded substrate D- and substrate D’-bound mono-RNPs. To verify that both sites are saturated we measured the 1D 1H spectra of the sRNP with increasing concentrations of substrate and followed the appearance of free substrate in the spectral region of the RNA H1’ peaks, which is devoid of protein peaks. We see no free-RNA peaks for a sub-stoichiometric ratio of substrate RNA:enzyme  For Figure 2D, bottom, the deviations between the experimental and simulated P(r) values are similar for the short distances (0 – 40 A) and for the large distances (70 – 140 A). The deviations for the small r values are independent of conformational changes and these are thus intrinsic to the method. The question is, if the deviations at larger r values are thus significant enough that one can conclude from these SANS data that there are two states? If yes, does this agree with the population ratio of the  and  states that the authors determined later in the manuscript?When we fit SAS data, we directly fit the scattering curves rather than the P(r) distributions. If we calculate the structures of the half-loaded sRNPs assuming only one state of each half-loaded complex, namely the -state as required by the PRE data, the resulting structures do not fit the directly measured SANS and SAXS curves. This is the main argument to support the existence of a conformational equilibrium. We show this now in Figure 2D and in Figure 2—figure supplement 3.2H-protein and 1H-RNA in 42%:58% D2O:H2O solvent mixture. This calculation does not take into consideration the fact that the scattering of the 1H-proteins is only perfectly matched at the beginning of the SANS curve and that atoms other than the Cα or P atoms contribute to the scattering. In the new version of Figure 2D we calculate the theoretical P(r) functions from the theoretical 2H-Fib SANS curves averaged over the ensemble of -structures that satisfy the PRE data of either the substrate D- or substrate D’-loaded sRNPs. Now, the theoretical P(r) functions at short distances perfectly match the experimental ones (as at short distances the P(r) function represents the pairwise distance distribution of atoms in one fibrillarin copy), while they differ at large distances demonstrating the incompatibility of the SANS data with a pool of -structures only.The choice of displaying the P(r) distributions in Figure 2D rather than the direct SAS scattering curves is done for visual reasons, as the P(r) distribution function is easier to interpret intuitively. The discrepancy between the experimental and theoretical functions noted by the reviewers in the first version of this manuscript was due to the fact that in the old Figure 2D, instead of calculating the P(r) curves from the simulated SANS curves of the structural ensembles, we displayed the histogram of the pair-wise Cα–Cα and P–P distances scaled by the respective scattering contrast of 11) Subsection “The conformational ensembles of the half-loaded mono-RNPs in solution” paragraph three: "show a reasonable similarity". The authors need to provide at least an RMSD and an overlay of their two structures with the structure of Lin et al. to support this statement.The overlays are shown in Figure 3—figure supplement 2. We have now added the RMSDs to the figure caption.12) In context of sRNP complex, the affinity of fibrillarin for substrate depends on RNA sequence beyond recognition site– It is unclear what the recognition site is. Referring to substrate sequences? Without a clearly defined recognition site, this claim is not supported by the data– Do the authors mean methylation site rather than recognition site as specified in the Discussion? If this is the case, the Abstract/Introduction should be modified to reflect this level of detail.We have changed “recognition site” to “methylation site” as this is what we mean.Major commentsPrevious points #4, 5 and 6; Legend Figure 2B: "Despite the absence of peak splitting, peak budding is visible in the spectrum of the substrate D'-loaded st-sR26 RNP in all instances for which peak splitting or budding is observed in the spectrum of the holo ssR26 di-RNP."1) This figure and its legend have been remade for this version of the manuscript. Nevertheless, we cannot follow what the authors see in their spectra. The only instance, where we can see a peak splitting (not sure if "budding" is a term that can be used), is in the purple (ssR26 holo) spectrum for residue L58 and, with some imagination, for I62. For all other panels and all other complexes, we fail to see any change in the resonance. In other words, for V118, L160, V141 and L200, there are effectively no chemical shift perturbations that are larger than 0.01 ppm in proton. In carbon there is no CSP whatsoever. For the open-closed conformations, one would expect many and large CSPs.This is a very important point, as the authors have little convincing NMR data that show that the complex exists in two states with fibrillarin either in the on- or the off-state.As noted also in the previous answers, it is not the NMR data by itself that indicates the existence of the conformational exchange but the combination of PRE and SAS data. To make this absolutely clear we have rewritten the entire Results section and remade Figure 2. We no longer discuss chemical shifts in terms of conformational exchange but focus only on the discrepancy between PRE and SAS data when attempting to fit them to a single conformational state. To provide further evidence that the conformational exchange is needed to fit the data we now compare the PRE and SAS fits for: the representative structures of  and -states ; ensembles of mixed - and -conformers ; ensembles of - OR -conformers only . While the PRE data can be fit in multiple scenarios, structural ensembles containing both - and -conformers  can properly fit the PRE and SAS data simultaneously. We are confident that this line of reasoning together with the new analysis presented in the supplements to Figure 3, supplement to Figure 4, new Figure 5 and its supplement, should be convincing.Please also note that we do not agree with the sentence “For the open-closed conformations, one would expect many and large CSPs.” Fibrillarin interacts with the RNA backbone and this interaction is driven by arginine and lysine amino acids, as well as other polar side-chains, but definitely not by methyl-group-containing amino acids. Indeed, of all the fibrillarin ILV methyl groups, only those of V110 are expected to be within 5 Å of the RNA. Thus, we expect only a few detectable methyl-group CSPs upon RNA binding. Nonetheless, to demonstrate more clearly that there are indeed CSPs upon RNA binding, we now show a different region of the spectrum containing peaks from the residues expected to be closest to the RNA. This region was not shown in the original version of the paper as the peaks are quite overlapped. Nevertheless, the CSPs are clear, although difficult to quantify due to the overlap.para/Idia still are very problematic. It is physically not possible that this ratio is larger than 1. In the plot, ratios that are 1.25 are shown. The authors write in their letter that there are only two values over 1, in the plot there appear to be at least 6. The remark that it is hard to extract reliable data for crowded spectral regions is correct, however, in case the data are unreliable or highly uncertain (which is not reflected in the presented error bars), it is not possible to draw solid conclusions. So how do the authors come to their conclusions based on the presented data? As an example, a ratio of 0.75 is mentioned in the text as highly relevant for the closed conformation, whereas a ratio of 1.25 is rebutted  as resulting from difficult spectral interpretations.2) Previous point #8; Figure 2C. The PRE data with an Ipara/Idia data-sets reported in the literature contain values higher than 1 and even 1.1:From the original PRE publication of Battiste and Wagner  onwards, all IExcerpt from the text of Battiste and Wagner: “The apparent noise or variation of intensity ratios for cross-peaks that should be far away from the spin-label nitroxide (no broadening effect) is approximately 10-15%. “Other examples of PRE values higher than 1.1 can be found in Huang et al. Scientific Reports 6, Article number: 33690 : e2207and many other publications.para/Idia values to at least 10%, as recommended by Battiste and Wagner, to take into account possible changes in intensity due to sample manipulation and/or sample ageing.Nevertheless, in order to address the concern regarding the values higher than 1, we have reprocessed all spectra to correct for a slight baseline offset present in three of them. We have then extracted all PREs from the newly processed spectra and confirmed that they are very similar to the old values. In addition we have increased the errors on the Ipara/Idia- error) larger than 1.1 among a total of 449 and 414 PREs, respectively. These numbers correspond to ~6% of the 258 (D’-loaded) and 261 (D-loaded) Ipara/Idia ratios higher than 0.9, namely those Ipara/Idia ratios corresponding to long distances. If one considers that the estimated error corresponds to the standard deviation of a Gaussian distribution, 68% of these “null PRE” values should be found between 0.9 and 1.10 (1 standard deviation), while another 15% should be found between 1.1 and 1.2. Thus the numbers of Ipara/Idia ratios higher than 1.1 are well within the statistics.For each of the D’-loaded and D-loaded complexes, we have ~15 values of , while the values to which we refer when invoking the closed conformation comprise 6 residues in a row, with values of Ipara/Idia = 0.5 in the D’-loaded complex and 0.75 in the D-loaded complex. We think that this piece of evidence, among others (see above), demonstrate that these intensity ratios are significant.The reviewers criticize that we invoke the presence of a closed conformation on the basis of IAs the NMR data do not agree with SANS data, the authors involve conformational rearrangements between open and closed conformations. Clearly, in case things are not in agreement, it can always be explained by dynamics. Because of the very weak NMR evidence (2 points above), it seems a far stretch to conclude that the complex undergoing open-closed motions.Please see the evidence for the conformational exchange now provided in Figures 3, 4 and 5 and their supplements. The ability of integrative structural biology to detect conformational exchange when data from only one technique could or would fail is exactly the point that we want to make here.Previous point #10; Figure 2D. The figure is still hard to understand. The substrate D loaded experimental (red) and theoretical (brown) are very different. The substrate D' loaded experimental (cyan) and theoretical (blue) are also very different.2H-Fib SANS curves in several figures and supplements.The figure has been removed. Instead of P(r) we now show the direct fit to the First, when there are issues raised by the reviewers that seem unreasonable to the authors, this sometimes reflects the fact that the material was not explained as clearly as might be possible. One of the advantages of eLife is that space is not nearly as constrained as in some other prestigious journals. […] Perhaps if you show more PRE data this will help – and as you will hopefully be showing correlation plots assuming 1 state and that these are not as good as the intra-validation – showing more raw data would be justified.In brief, for this revised version, we have re-analysed PRE data after reprocessing all spectra, redid all structure calculations and ensemble scoring with the new sets of PRE data (very similar to the previous ones), added new analysis and validation figures and rewritten the Results section. In addition, we no longer discuss chemical shift perturbations to support the existence of conformational exchange. The conclusions are identical to the first version of the manuscript: the fact that we now refer to a new round of structural calculations justifies the small differences in the exact numbers reported from Figure 3 onwards.


Oestrogen receptor-α (ERα) shuttles continuously between the nucleus and the cytoplasm, and functions as an oestrogen-dependent transcription factor in the nucleus and as an active mediator of signalling pathways, such as phosphatidylinositol 3-kinase (PI3K)/AKT, in the cytoplasm. However, little is known regarding the mechanism of ERα nucleocytoplasmic shuttling. In this study, we found that ERα is transported into the nucleus by importin-α/β1. Furthermore, we found that Transportin-2 (TNPO2) is involved in 17β-oestradiol (E2)-dependent cytoplasmic localisation of ERα. Interestingly, it was found that TNPO2 does not mediate nuclear export, but rather is involved in the cytoplasmic retention of ERα via the proline/tyrosine (PY) motifs. Moreover, we found that TNPO2 competitively binds to the basic nuclear localisation signal (NLS) of ERα with importin-α to inhibit importin-α/β-dependent ERα nuclear import. Finally, we confirmed that TNPO2 knockdown enhances the nuclear localisation of wild-type ERα and reduces PI3K/AKT phosphorylation in the presence of E2. These results reveal that TNPO2 regulates nucleocytoplasmic shuttling and cytoplasmic retention of ERα, so that ERα has precise functions depending on the stimulation. ERα proteins are primarily located in the nucleus, regardless of ligand binding, but shuttle constantly between the nucleus and the cytoplasm. Upon oestrogen binding, ERα changes conformation that enables dimer formation and active transcription of target genes. Oestrogen-bound ERα has been reported to decrease shuttling and further accumulate in the nucleus4. Outside the nucleus, a small population of ERα is localised to the plasma membrane5, which requires palmitoylation, in interaction with the membrane protein caveolin-16. Oestrogen-bound ERα on the plasma membrane can rapidly induce the mobilisation of intracellular calcium7 and activation of PI3K/AKT and MAPK/ERK pathways11. Abnormal phenotypes of nuclear-only ERα and membrane-only ERα transgenic mice suggested that normal organ development and function require convergence and crosstalk of the classical nuclear and extranuclear receptor pathways17. Moreover, the cytoplasmic ERα mutant, which forcibly changed the localisation of ERα to the cytoplasm, lost oestrogen-induced transcriptional responses, but still maintained MAPK/ERK activation18. Recently, cytoplasmic ERα mutant mice have been generated, and preliminary observations suggest that their phenotypes are similar to those of ERα deficient mice19. Therefore, nucleocytoplasmic shuttling plays an important role in the regulation of ERα signalling pathways.Oestrogen receptor-α (ERα) plays important roles in various physiological processes, such as male and female reproductive development, lipid and bone metabolism, and the maintenance of the cardiovascular and nervous systems20. Additionally, seven members of the importin-α family, which function as an adaptor between the NLS-containing cargo molecule and importin-β1, have been identified in humans21. Importin-α and importin-β families have been reported to be involved in the nucleocytoplasmic shuttling of steroid hormone receptors. For instance, importin-α/β1 heterodimers, importin-7, importin-8, and importin-13 mediate nuclear import of glucocorticoid receptor (GR)24. Importin-α/β1 heterodimers also mediate the nuclear import of mineralocorticoid receptor25 and androgen receptor (AR)26. Moreover, CRM1 (exportin-1)27 and exportin-528 can stimulate AR export. The distribution of thyroid hormone receptors is regulated by CRM129, exportin-4, exportin-5, and exportin-730. However, only a few studies have shown the nuclear export of ERα32; thus, the regulatory mechanism in ERα nucleocytoplasmic shuttling remains poorly understood.Nucleocytoplasmic transport of macromolecules occurs through the nuclear pore complex (NPC), and is generally dependent on the presence of a specific signal sequence, called nuclear localisation signals (NLSs) and nuclear export signals (NESs). Many nucleocytoplasmic transport pathways are mediated by the importin-β superfamily, which is composed of more than 20 family members in humans, and is functionally classified as importins and exportins. The importin-β superfamily exhibits cargo specificity, but has the common features that recognise the NLS or NES sequences of cargo molecules and the GTP-bound form of Ran (RanGTP), a member of the Ras superfamily, and mediate transport between the nucleus and the cytoplasmHere, we sought to gain possible insights into the mechanism of ERα nucleocytoplasmic shuttling. We found that ERα is transported into the nucleus via two different pathways: active transport by importin-α/β1 and passive diffusion. Our results also show that Transportin-2  enhances the cytoplasmic localisation of ERα, not as a nuclear export receptor, but rather as a cytoplasmic retention factor, while it also competitively inhibits the binding of importin-α. Furthermore, we confirmed that TNPO2 knockdown increased the nuclear localisation of endogenous ERα and suppressed PI3K/AKT activation in the presence of its ligand, 17β-oestradiol (E2). We propose that TNPO2 plays a critical role in determining the distribution of ERα between the nucleus and the cytoplasm in response to stimulation.34. To determine the essential amino acid residues for ERα active nuclear transport, we introduced amino acid substitution mutations in the NLS region by converting the three basic amino acid clusters into alanine   and the hinge regionine Fig. A. Since ed GFP ~ 03 kDa (FDa) Fig. A. These Da) Fig. B, or imped GFP ~ 03 kDa (FDa) Fig. A to an iDa) Fig. C. When iDa) Fig. . In contDa) Fig. ” below. Da) Fig. B, althoued GFP ~ 03 kDa (FDa) Fig. ” below.FNext, we analysed the subcellular distribution of 3 × HA-ERα (~ 70 kDa), which is smaller than GFP-ERα, in HeLa cells. 3 × HA-ERα was predominantly localised in the nucleus, corresponding to the distribution of GFP-ERα Figs. B. On theWe investigated the effect of E2 treatment on the subcellular distribution of ERα. 3 × HA-ERα showed only a slight change, but 3 × HA-ERα-mNLS was dramatically translocated to the cytoplasm by E2 treatment , an inhibitor of CRM1-dependent nuclear export38. As shown in Supplementary Fig. 36. These contrasting results on the LMB effect in the subcellular localisation of ERα are probably caused by the NLS mutant used in this study instead of the wild type ERα used in the previous reports. Therefore, it is possible that the imbalance in ERα-mNLS transport allowed us to observe the nuclear export more clearly. Meanwhile, we confirmed the binding of recombinant GST-CRM1 and 3 × HA-ERα-mNLS exogenously expressed in HeLa cells using a GST pulldown assay, as described previously32 41 genes using 3 × HA-ERα-mNLS, as a useful tool to easily observe ERα export. The knockdown efficiency of the corresponding siRNAs was determined using quantitative real-time PCR analysis   or basic motif, followed by a C-terminus R/H/K-X(2–5)-P-Y (X is any residue) consensus sequence46. The 61–180 residues of the AF-1 domain of ERα contain two similar PY-motifs: 64-AAAAANAQVYGQTGLPY-80 and 117-LSPFLQPHGQQVPY-130. First, in order to clarify the relationship between ERα and TNPO2, we generated two deletion mutants lacking the AF-1 domain of ERα-mNLS, residues—61–595 (ERαΔ1-mNLS) and 181–595 (ERαΔ2-mNLS), respectively  motifNext, we investigated the binding of wild-type ERα and its deletion mutant lacking the AF-1 domain ERαΔ2) Fig. A to TNPO Fig. A t11. Given that TNPO2 could bind to two PY-motifs of ERα in the cytoplasm and competitively bind to importin-α in its basic NLS region in an E2-dependent manner, we speculated that TNPO2 may function in the cytoplasmic retention of ERα to induce cytoplasmic signalling pathways. To further investigate the role of TNPO2 in the regulation of ERα localisation, we co-transfected HeLa cells with 3 × HA-ERα and 3 × Flag-TNPO2 expression vectors. Immunofluorescence staining showed that 3 × HA-ERα was mainly localised in the nucleus of untreated HeLa cells, with or without co-expression of 3 × Flag-TNPO2. However, upon E2 treatment for 10 min, 3 × HA-ERα shifted more to the cytoplasm when the 3 × Flag-TNPO2 was co-expressed, as compared with 3 × HA-ERα only  forms heterodimers with other importin-α, and proposed that importin-α8 can suppress cNLS binding through hetero-dimerization52. Here, we found that TNPO2 could competitively inhibit the nuclear import of ERα by importin-α/β1. The transportin subfamily was first suspected to act as a nuclear bidirectional receptor; however, subsequent studies refuted this hypothesis. It is thought that the transportin subfamily in nucleocytoplasmic transport is restricted to nuclear import45. As a typical example, TNPO1 and TNPO2 mediate nuclear import of heterogeneous nuclear ribonucleoprotein (hnRNP) A1 through the interaction of the M9 transport signal. However, even though M9 is known to serve as a bidirectional signal, a nuclear export receptor that recognises M9 is yet to be identified54. Interestingly, similar to our results, transportin can bind to Deleted in breast cancer 1 (DBC-1) with and without NLS55. Therefore, the transportin subfamily may play a role in the regulation of cytoplasmic signalling as well as in nuclear transport.So far, it has been reported that some importins could inhibit the nuclear transport mediated by other importins. For example, importin-α inhibits the nuclear import of Snail mediated by importin-β19, Shc56, HER-210, and EGFR11. In the future, an accurate understanding of the cytoplasmic roles of the transportin subfamily may provide insight into the novel regulatory mechanism of the biological function of transportin-cargos.Of note, our data show that TNPO2 is localised on the plasma membrane and colocalises with ERα in MCF-7 cells, both of which were diminished in TNPO2 knockdown cells  facilitates translocation of ERα to the cytoplasm and repopulation of cancer cells that have acquired resistance to this treatment61, pcDNA3.1/Zeo(+)-EGFP DEST62, and pcDNA3.1/Zeo(+) DEST, pGEX-6P-2 DEST, which were generated by a method similar to63 using pcDNA3.1/Zeo(+) (Thermo Fisher Scientific), pGEX-6P-2  and the Reading Frame Cassette of the Gateway Conversion System (Thermo Fisher Scientific). pCold II Ran Q69L was constructed by inserting the human Ran Q69L mutant into pCold II DNA .Human ERα, importin-α1, importin-α2, importin-α4, importin-β1 and TNPO2 were generated by polymerase chain reaction (PCR) amplification from total RNA from MCF-7 and HeLa cells using appropriate primers (Supplementary Table ), and subsequently subcloned into the pENTR vector . ERα mutants and importin-α ΔIBB were constructed by site-dirHeLa and MCF-7 cells were cultured in Dulbecco’s modified Eagle’s medium  supplemented with 10% foetal bovine serum (FBS). The cells were maintained in a 37 °C incubator with 5% CO2-humidified air.For E2 treatment, cells were maintained in 5% Charcoal/Dextran-treated FBS (Thermo Fisher Scientific) in phenol red-free D-MEM  for 2 days, and then serum-free media before experimentation.66. Briefly, HeLa cells were grown on multiwell plates  for 2 days. After washing with ice-cold transport buffer , the cells were permeabilised with 20 µg/mL Digitonin, High Purity  in TB on ice for 5 min. Cells were then washed and immersed in ice-cold TB for 10 min. Preincubated 400 nM GFP-cargo-substrate, 600 nM importin-α, 400 nM importin-β1 and 4 µM RanGDP were mixed with 500 µM GTP and an ATP-regeneration system  in TB containing 1% BSA. After applying the mixture, the digitonin-permeabilized cells were incubated at 30 °C for 30 min. Finally, the cells were rinsed with ice-cold TB and fixed with 3.7% formaldehyde in TB for 15 min. The cells were then stained with DAPI.An in vitro transport assay was performed as described previouslyFect A plus  according to the manufacturer's instructions. siRNA duplexes were reverse transfected into HeLa and MCF-7 cells using Lipofectamine RNAiMAX Transfection Reagent (Thermo Fisher Scientific) as recommended by the manufacturer.Transfections were performed using Lipofectamine 2000 Transfection Reagent (Thermo Fisher Scientific) and ScreenERα Antibody (F-10) , Anti-HA-tag mAb , Monoclonal ANTI-FLAG M2 antibody produced in mouse , Ran BP-1 Antibody (M-45) , Actin Antibody (C-11) , Purified Mouse Anti-Karyopherin β , Purified Mouse anti-Transportin , TNPO2 Polyclonal antibody , IPO4 Rabbit pAb , Anti-Importin 7 antibody , Anti-Cellular Apoptosis Susceptibility/CSE1L antibody , Purified Mouse Anti-Ran , Phospho-Akt (Ser473) (D9E) XP Rabbit mAb , Akt (pan) (C67E7) Rabbit mAb  were used.Importin-β family, Hikeshi and CALR Silencer Select (Pre-designed) siRNAs were synthesized by Thermo Fisher Scientific: importin-β1 (ID: s7917), TNPO1 (ID:s7933), TNPO2 (ID:s26881), TNPO3 (ID:s24032), importin-4 ID:s36153), importin-5 (ID:s7935), importin-7 (ID:s20640), importin-8 (ID:s20637), importin-11 (ID:s27652), importin-13 (ID:s18609), RanBP6 (ID:s25641), RanBP9 (ID:s19529), exportin-7 , exportin-2 , exportin-4 (ID:s34638), exportin-5 (ID:s33190), exportin-6 (ID: s23302), exportin-T (ID:s22232), RanBP17 (ID:s35075), Hikeshi (ID:s28222), Calreticulin (ID:s114).Escherichia coli strain BL21 and RosettaBlue (Merck Millipore), and purified with Glutathione Sepharose 4B . GST cleavage was performed using Turbo3C Protease . Hexahistidine (6 × His)-RanQ69L was expressed in BL21 and purified with TALON Metal Affinity Resin (Takara Bio Inc.). The GTP-bound RanQ69L was purified as described previously64.GST-fusion proteins were expressed in 2, 0.05% Tween-20, 1 μg/mL each of aprotinin, leupeptin, and pepstatin A). These lysates were centrifuged at 20,400×g for 30 min, and the supernatants were collected and quantified using a DC Protein Assay kit . GST pulldown assays were performed by mixing cell lysates with Glutathione Sepharose 4B charged with 50 pmol GST fusion proteins with or without 2 µM 6 × His-RanQ69L. The beads were washed with lysis buffer and resuspended in Laemmli sample buffer, and subsequently analysed by SDS-PAGE and western blotting. Quantification of western blot band signals was performed with CS Analyzer software and statistical analysis was performed using GraphPad Prism.At 24 h after transfection, HeLa cells were washed with cold PBS. The cells were lysed with lysis buffer , the cells were incubated with primary antibodies. After washing with PBS, Alexa Fluor Plus 488- and 555-conjugated secondary antibodies (Thermo Fisher Scientific) were used. Nuclei were stained with 0.1 μg/mL DAPI . Fluorescence was observed using an Olympus FV-1200 confocal microscope . Quantification of nuclear signal was performed with ImageJ software, and statistical analysis was performed in GraphPad Prism.HeLa cells and MCF-7 cells grown on glass cover slides in a 12-well culture plates were fixed with formaldehyde for 10 min at room temperature, permeabilized with PBS containing 0.2% TritonX-100 and washed with PBS. After blocking with blocking buffer . The amplicons were designed to amplify fragments of > 150-bp (Supplementary Table Supplementary Information.