Streptococcus pyogenes, namely, PNPase, YhaM, and RNase R. We observe that RNase Y preferentially cleaves after guanosine, generating substrate RNAs for the 3′-to-5′ exoRNases. We demonstrate that RNase Y processing is followed by trimming of the newly generated 3′ ends by PNPase and YhaM. Conversely, the RNA 5′ ends produced by RNase Y are rarely further trimmed. Our strategy enables the identification of processing events that are otherwise undetectable. Importantly, this approach allows investigation of the intricate interplay between endo- and exoRNases on a genome-wide scale.RNA degradation is an essential process that allows bacteria to control gene expression and adapt to various environmental conditions. It is usually initiated by endoribonucleases (endoRNases), which produce intermediate fragments that are subsequently degraded by exoribonucleases (exoRNases). However, global studies of the coordinated action of these enzymes are lacking. Here, we compare the targetome of endoRNase Y with the targetomes of 3′-to-5′ exoRNases from  Streptococcus pyogenes, revealing the interplay between RNase Y and 3′-to-5′ exoribonuclease PNPase and YhaM.Bacterial RNA degradation is typically initiated by endoribonucleases and followed by exoribonucleases. Here the authors report the targetome of endoRNase Y in  The ability to modulate gene expression enables bacteria to rapidly adapt to their environment. Ribonucleases (RNases) regulate transcript abundance, leading to RNA maturation , stabilization or degradation. Eventually, all transcripts—even the most stable—are degraded by RNases, leading to the renewal of the nucleotide pool.1. The main endoRNases that have been demonstrated to initiate RNA decay are RNase E in Gram-negative bacteria and its functional orthologue RNase Y in Gram-positive bacteria3. However, in many Gram-positive bacteria, RNA degradation can also be initiated by the complex of RNases J1/J2, which displays both endo- and 5′-to-3′ exoribonucleolytic activities6. The initial processing of a transcript is the limiting step of the RNA decay and the access of endoRNases to transcripts is usually restricted. For instance, RNase E favours 5′ monophosphorylated (5′ P) transcripts and cleaves 2 nt upstream of a uridine (U) in A/U rich regions8. RNase Y also prefers 5′ P transcripts9 and additional requirements have been described depending on the orthologue studied. In Staphylococcus aureus, RNase Y processes transcripts preferably downstream of a guanosine (G)10. In Streptococcus pyogenes, a G is required for the in vivo processing of the speB transcript, encoding a major virulence factor11. In Bacillus subtilis and S. aureus, RNase Y processing relies on proximal RNA secondary structures12.As a general rule, RNA degradation starts with an endonucleolytic processing in the RNA body, leading to the generation of decay intermediates. Those are further digested by exoRNases and, finally, by oligoRNase/nanoRNases13. In E. coli, the decay intermediates are mainly degraded by the 3′-to-5′ exoRNases II, R and PNPase15. In B. subtilis and S. pyogenes, the major 3′-to-5′ exoRNase is PNPase17. In addition, in B. subtilis, the 3′-to-5′ exoRNases PH and YhaM participate in RNA decay, albeit with lower efficiency than the main 3′-to-5′ exoRNase16. S. pyogenes YhaM exhibits a very short processivity (3 nt on average) on a large number of RNA 3′ ends, the impact of which on mRNA decay is currently unknown17.The decay intermediates, once generated by endoRNase(s), are cleared immediately from the cell by 3′-to-5′ exoRNasesS. aureus and B. subtilis18. To date, the activity and specificity of 3′-to-5′ exoRNases towards decay intermediates produced by a given endoRNase have never been studied on a global scale.With the emergence of RNA sequencing techniques allowing the global detection of RNase cleavage sites, several targetomes of endoRNases have been determined, such as those of RNase Y in S. pyogenes. We study the interplay of endoRNase Y with 3′-to-5′ exoRNases in S. pyogenes, a pathogen causing a wide range of diseases in humans. We determine the first targetome of RNase Y in this bacterium and compare it with three 3′-to-5′ exoRNase targetomes, previously characterized by our laboratory17. We show on a global scale that RNase Y mainly acts in concert with PNPase during RNA degradation. In this regard, we demonstrate a role of the RNase Y-PNPase interplay in the control of the differential stability of polycistronic mRNAs and the decay of 5′ regulatory elements. This strategy allows us to elucidate the interplay and dynamics of endoRNase- and exoRNase-mediated RNA processing events otherwise not detectable when RNases are studied separately.Here, we present a comparative RNA-seq based approach that allows us to dissect the complex landscape of RNA ends in S. pyogenes wild type (WT), RNase Y deletion mutant (∆rny) and complemented RNase Y deletion mutant (∆rny::rny) strains, as described previously for other RNases in this bacterium19  in the ent Fig. . Therefoent Fig. . The 5′ nds Fig. . Second,nds Fig. . This stnds Fig. . Third, ons Fig. . The obsrny_ends originated from 3′-to-5′ exoribonucleolytic activity, we compared the RNase Y targetome with the targetomes of three 3′-to-5′ exoRNases  recently characterized by our laboratory17  strains. The RNA 3′ ends more abundant in the ∆exornase strain and the 3′ ends more abundant in the WT strain were annotated as exoRNase trimming start and stop positions, respectively  around PNPase stop positions using 25 nt-long sequences, which are sufficient for detection of terminator structures, and we could not predict any structure17. Here, we used 50 nt sequences for the calculation, allowing us to detect variations in ∆G that are indicative of structures longer and weaker than the structures of terminator regions. Indeed, we observed a decrease in the MFE upstream of the PNPase stop positions , as the PNPase stop positions were not detected  grew slower than both the ∆rny strain and the YhaM-RNase Y double-deletion strain (∆yhaM∆rny)  Fig. , we calcnds Fig. . We obseart Fig. . ExampleyhaM strain, by neither northern blot analyses nor RNA sequencing 17  were generated by RNase Y 17 Fig. . Here, w)17 Fig. 17. Basede Y Fig.  and b. I17. Here, we observed that some of these decay intermediates were produced by RNase Y rsmC-cdd-bmpA operon, described below , a cytidine deaminase involved in pyrimidine metabolism (cdd) and a lipoprotein (bmpA)  Fig. . RNase Yend Fig. . This obrsmC-cdd-bmpA transcript (~2900 nt) was greatly increased in the ∆rny strain , which was barely detectable in the WT, was stabilized in the ∆pnpA strain , which is consistent with previous reports for the S. aureus and B. subtilis RNase Y proteins, describing ~100 processing positions (identified by 5′ end sequencing)18. In S. aureus, the limited impact of RNase Y on global transcript stability is consistent with the low number of detected direct targets10. In contrast, for RNase E, the major endoRNase initiating RNA decay in Gram-negative bacteria, ~22,000 processing positions were identified in Salmonella enterica8. A possible explanation for the high number of RNase E processing events detected in this bacterium is the absence of RNase J1, which is found mainly in Gram-positive bacteria27 and performs degradation from the 5′ end of the RNAs. In the present study, it is likely that we underestimated the number of RNase Y processing sites. First, because the method used relies on the detection of at least one RNA end (5′ or 3′), we did not identify RNase Y processing events when both generated ends were subsequently degraded by exoRNases  in addition to RNase Y. For example, the RNase J1/J2 complex could play an important role in RNA decay in S. pyogenes, as both enzymes are essential in this bacterium5.Although we showed that RNase Y is involved in RNA decay, we believe that, due to the limited number of direct targets identified, RNase Y might not be the major initiator of mRNA decay in ses Fig. . Second,ses Fig. . Comparises Fig. . It is pB. subtilis, although this interaction was not required for the degradation of all the studied targets29. When PNPase targeted the RNA fragments generated by RNase Y processing, we detected more PNPase trimming stop positions than start positions  and maturation of polycistronic mRNA. A role of RNase Y in the turnover of regulatory elements was previously observed in both ron Fig. .17. Therefore, it was expected that YhaM would nibble the RNA 3′ ends produced by RNase Y. This activity complicated the identification of the original processing positions of RNase Y, as the G characterizing RNase Y activity was removed from the RNA 3′ end by YhaM  and isogenic gene deletion strains  supplemented with 3% sheep blood was used as a solid medium.s SF370 MGAS and i2), diluted to an OD620 nm of 0.02 in 5 ml of medium and centrifuged at 3200 × g for 5 min. The resuspended pellet was used to inoculate flasks containing 25 ml of THY supplemented with 0.2% yeast extract. The growth was monitored by measuring the OD620 nm using a microplate reader (Biotek PowerWave XS2). Growth curve experiments were performed in triplicate and the standard error of the mean was calculated.Bacterial cultures were grown overnight  was added when the bacteria reached the mid-logarithmic phase of growth, and samples were taken either after 0, 5, 10, 20, 30 and 45 min or after 0, 1, 2, 4 and 8 min. The cells were rapidly harvested by mixing the 25 ml of cultures with 25 ml of 1:1 ice-cold acetone/ethanol solution and by centrifugation (3500 × g for 10 min at 4 °C). The pellets were thoroughly resuspended in 5 ml of TE buffer . The cells were lysed by adding 100 µl of lysis buffer  supplemented with 2.5 mg/ml of lysozyme and 0.5 µg/µl of mutanolysin and incubated for 5 min on ice. The samples were mixed with the lysis executioner buffer (2% sodium dodecyl sulfate (SDS), 1 mg/ml Proteinase K) incubated at 95 °C for 1.5 min. Seven hundred and fifty microlitres of TRIzol reagent (Life Technologies) were added to the samples, which were subsequently inverted three times. After incubation for 5 min at room temperature, 200 µl of chloroform were added and the samples were mixed by vortexing. The samples were then incubated at room temperature for 10 min prior centrifugation at 11,300 × g for 10 min at 4 °C. The upper aqueous phase (~700 µl) was gently collected and the RNAs were precipitated with ice-cold 100% isopropanol at a 1:1 ratio at −20 °C for at least 1 h. After centrifugation at 11,300 × g for 10 min at 4 °C, the RNA pellets were washed with 1 ml of 70% ethanol, air dried for 10 min and dissolved in autoclaved Milli-Q H2O. RNA integrity was assessed on 1% agarose gels.The overnight 17. After treatment with TURBO DNase (Ambion), the RNA quality was assessed using a bioanalyzer system (Agilent 2100). Subsequently, 4.5 µg of RNA was depleted of rRNAs ) and treated with 10 U of RppH (New England Biolabs) at 37 °C for 1 h 30 min to convert the 5′ triphosphate RNAs in 5′ monophosphate RNAs. The RNAs were purified using standard extraction with phenol:chloroform:isoamylalcohol  and precipitated using ice-cold ethanol. The obtained RNAs were treated with T4 polynucleotide kinase (Thermo Scientific) according to the manufacturer’s instructions to allow the subsequent ligation of the sequencing adaptors. After a purification step using the RNA Clean & Concentrator kit (Zymo Research), the RNAs were fragmented (Covaris M220) in a microTUBE AFA Fiber Pre-Slit Snap-Cap tubes for 140 s. cDNA libraries were prepared using the NEXTflex® Small RNA Sequencing Kit v3 (Bioo Scientific) according to the manufacturer’s instructions until step G. The purification step was performed using Agencourt AMPure XP beads (Beckman Coulter). The cDNA libraries were sequenced on a HiSeq3000  at the Max Planck-Genome-centre Cologne. The data were deposited in the National Center for Biotechnology Information (NCBI) sequence read archive (SRP149896). The numbers of sequencing reads obtained are listed in Supplementary Table S. pyogenes reference genome (NC_002737.2) and the total and end-specific (5ʹ and 3ʹ) coverage profiles were visualized using the Integrative Genomics Viewer (IGV)34. Differentially expressed (DE) genes were identified using featureCounts (v1.5.2)35 and edgeR (v3.20.6)37 with absolute log2-fold change (log2 FC) ≥ 1 and false discovery rate (FDR) < 0.05.The RNA sequencing was performed in biological triplicates using the workflow previously published by our laboratory Agilent 100. Subs17. In brief, the genome coverage data was prefiltered with a count per million (cpm) value ≥0.05 and only the RNA ends displaying a cpm ≥5 were further analysed. We carried out differential expression analysis of normalized 5′ and 3′ read ends by comparing the following datasets (triplicates): WT vs. ∆rny and ∆rny vs. ∆rny::rny. RNase Y ends were identified using edgeR (v3.20.6)37 with absolute log2 FC ≥ 1 and FDR < 0.05, and kept only if present in both comparisons. These identified RNA ends were named 5′ or 3′ rny_ends (Supplementary Data rny proportion of ends”) that were previously described17. First, the “proportion of end parameter”, proportions of RNA ends at one position over the total RNA abundance, from the WT strain was set as ≥2%. Second, the “ratio of WT and ∆rny proportion of ends”, the WT proportion of ends over the ∆rny proportion of end, was set as ≥3%. This parameter allowed that the identification of RNA ends depending of an RNase Y processing event was independent from the RNA abundance in the WT and ∆rny strains.RNase cleavage positions were identified following the previously published procedurerny_ends) was compared to the PNPase, YhaM and RNase R targetomes , which were previously identified (SRP149886)17   using a sliding window of 50 nt sequences, with 100 or 200 nt centred on the position of interest. The average MFE at each nucleotide was then calculated. WebLogolib (v3.5.0) was used to generate the sequence logos39, with sequences of 20 nt centred on the processing site with a GC content of 38.5%. The plots were generated using Python (v3.6.3) and matplotlib (v2.1.0).RNAfold (v2.4.3)32P labelled using T4 Polynucleotide Kinase (T4 PNK Fermentas) in presence of 2 µl of T4 PNK buffer (10×) and 2 µl of gamma-32P ATP 40. The labelled oligonucleotides probes were subsequently purified over G-25 columns  following the manufacturer’s instructions. The membranes were prehybridized in the Rapid-hyb buffer  for 1 h at 42 °C. The oligonucleotides were denatured and added to the membranes for an overnight hybridization at 42 °C. The membranes were washed first with 5X SSC-0.1% SDS buffer and then with 1X SSC-0.1% SDS buffer for 15 min at 42 °C, respectively. RNA sizes were estimated using the RNA DecadeTM Marker (Ambion) or the ΦX174 DNA/HinfI Marker (Fermentas).For the short RNAs, 10 µg of RNA was separated on 8% or 10% polyacrylamide/8 M urea gels Figs. , and 7 i11. The RNAs were crosslinked to the membranes using UV . The membranes were pre-hybridized in Rapid-hyb buffer for 1 h  and subsequently incubated with denatured probes, which were 32P labelled as described above. The hybridization was conducted overnight at 50 °C. The membranes were rinsed twice with pre-warmed 1X SSC + 0.1% SDS and subsequently with pre-warmed 0.5X SSC + 0.1% SDS. All washes were performed for 20 min at 50 °C. Long RNA sizes were estimated using the RiboRuler High Range Ladder (Thermo Scientific). A Typhoon Fla 9500 phosphorimager (Fujifilm) was used to visualize the radioactive signal for all northern blotting assays. As a loading control, 5S or 16S rRNAs were probed on the same membranes. Each northern blot was performed in at least triplicate. The uncropped blots are supplied in the Source Data file.For the long RNAs Fig. , 20 µg o19. RNAs were annealed to the radiolabelled primer (Supplementary Table g for 4 °C for 10 min. The pellet was washed with ice-cold 70% ethanol at 20,000 × g for 4 °C for 10 min and resuspended in 5 µl of 2X RNA loading dye. The cDNA products were resolved on 10% polyacrylamide/8 M urea/TBE gels and the size of the products was estimated using the AFLP 30–300 bp ladder labelled according to the manufacturer’s instructions.Primer extension was conducted on 10 µg of total RNA in biological triplicatesFurther information on research design is available in the Supplementary InformationPeer Review FileReporting SummaryDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Supplementary Data 3Supplementary Data 4Supplementary Data 5Supplementary Data 6Supplementary Data 7


The ubiquitous redox coenzyme nicotinamide adenine dinucleotide (NAD) acts as a non-canonical cap structure on prokaryotic and eukaryotic ribonucleic acids. Here we find that in budding yeast, NAD-RNAs are abundant (>1400 species), short (<170 nt), and mostly correspond to mRNA 5′-ends. The modification percentage of transcripts is low (<5%). NAD incorporation occurs mainly during transcription initiation by RNA polymerase II, which uses distinct promoters with a YAAG core motif for this purpose. Most NAD-RNAs are 3′-truncated. At least three decapping enzymes, Rai1, Dxo1, and Npy1, guard against NAD-RNA at different cellular locations, targeting overlapping transcript populations. NAD-mRNAs are not translatable in vitro. Our work indicates that in budding yeast, most of the NAD incorporation into RNA seems to be disadvantageous to the cell, which has evolved a diverse surveillance machinery to prematurely terminate, decap and reject NAD-RNAs. NAD (nicotinamide adenine dinucleotide) acts as a non-canonical RNA cap structure in bacteria and eukaryotes. Here the authors demonstrate the whole landscape of budding yeast NAD-RNAs which are subject to diverse surveillance pathways, suggesting that NAD caps in budding yeast are mostly dysfunctional. The cap is hydrolyzed by various decapping enzymes3, thereby triggering RNA degradation. Recently, another type of 5′-cap structure was discovered, both in prokaryotes6 and eukaryotes10, which is derived from the ubiquitous redox coenzyme NAD. Although in human and plant cells a diverse landscape of NAD-RNAs was found, only 37 RNA species and low abundance were reported in budding yeast8, questioning the biological significance of NAD capping in this organism. As the protocol used in this work excluded the small RNA fraction, which had been particularly rich in NAD-RNAs in prokaryotes4, we address here the whole landscape of NAD transcripts in yeast using the original NAD captureSeq protocol11. We find that NAD-RNAs are ubiquitous (1400 in wild type (WT), several thousands in mutants), most of them being short species (<170 nt). Only very few RNAs are detected with lengths over 250 nt. Ab initio incorporation by RNA polymerase (RNAP) II is found to be the predominant mechanism for NAD incorporation, and for about half of the transcripts, RNAP II uses transcription start sites (TSSs) different from those utilized for m7G-capped RNAs of the same gene. A YAAG core promoter motif is found to correlate with efficient transcriptional NAD incorporation. On average, NAD-RNAs are shorter than non-NAD-RNAs. By deleting the (putative) NAD-RNA decapping enzymes Npy1, Dxo1, and Rai1, we identified overlapping populations of RNAs likely decapped by these enzymes. Sequence analysis of total RNA from each mutant strain supports a hierarchical order of NAD-RNA processing, in agreement with the subcellular locations of the enzymes. NAD-mRNAs that escape decapping do not support translation by cytosolic ribosomes in vitro. We propose that (at least for the nuclear transcripts studied here) NAD incorporation into yeast RNA is largely accidental, due to competition of NAD and ATP in transcription initiation. We speculate that the NAD modification is in most cases undesirable to the cell, which first disfavors the synthesis of full-length NAD-RNAs, then decaps them rapidly using a multi-tiered machinery localized in different compartments and—even if they reach full length and escape decapping—ultimately rejects them from ribosomes.In eukaryotes, the 5′-terminus of messenger RNAs is protected by a m4, in which the enzyme adenosine diphosphate-ribosylcyclase (ADPRC) tags NAD-RNAs at the NAD moiety, followed by click-chemistry biotinylation and selective isolation by streptavidin binding. After adapter ligation, reverse transcription (RT), and PCR amplification, amplicons with sizes between 150 and 300 bp were selected by gel electrophoresis; thus, this library represented mostly RNA species with sizes between 20 and 170 nt present in the original sample. Enrichment was determined by quantitative comparison with a minus ADPRC negative control. In this unfragmented library, 1460 RNAs were found to be enriched, with changes reaching up to 1200-fold , mainlyold Fig. . Five ouTo test whether highly expressed transcripts are generally more likely to be enriched in NAD captureSeq, we compared the enrichment levels observed in the unfragmented NAD captureSeq library with the transcript abundance determined by transcriptome sequencing. This analysis revealed no correlation , the 5′-NAD modification was directly identified and quantified by mass spectrometry (MS) after pulldown Fig. , confirmEscherichia coli, the Nudix hydrolase NudC acts as an efficient decapping enzyme for NAD-RNA13. The yeast homolog Npy1 is known to hydrolyze the pyrophosphate bond in NAD to yield nicotinamide mononucleotide (NMN) and adenosine monophosphate (AMP)14, and was recently suggested as an NAD-RNA decapping enzyme13. The only support for this claim was, however, its in vitro processing of a synthetic NAD-RNA 12mer into a product that migrated on HPLC like a 12mer-5′-monophosphate RNA (p-RNA) and the inactivity of an active-site mutant to produce this product13. To characterize the in vitro activity of Npy1, we purified the protein from E. coli and analyzed its reaction kinetics with an in vitro-transcribed NAD-RNA (a 98 nt 5′-fragment of TDH3 RNA) on acryloylaminophenyl boronic acid (APB) gels, which separate NAD-RNA from p-RNA15. Purified Npy1 decapped NAD-RNA without inducing nucleolytic degradation and had no effect on m7G-RNA in vitro  were NAD-capped (relative to WT), which were almost half of all detected RNA species , removal of Npy1 reduced the total amount of NAD attached to RNA by ~60%  and modification ratio (change). Plotting the modification ratio of npy1∆ mutant vs. WT confirmed that the global reduction of NAD modification is not caused by few strongly reduced species that override the effects of many weakly increased ones. The slope <1 (0.61) of this plot confirms that, on average, the modification in the npy1∆ mutant is lower than in the WT  had the strongest negative effect on growth in normal medium and in the presence of increasing concentrations of ethanol , and the strongest global effect on RNA expression was noticed for removal of RAI1 , indicative of their modification with NAD. No significant further increase was observed in the double- and triple- deletion strains  and small RNAs (35%) were detected by NAD captureSeq  four enzymes as follows: RNAP II, Rai1, Dxo1, and Npy1. Using the deletion mutants, we first analyzed transcriptional preferences. Although the sequencing read profiles of some RNAs revealed homogenous 5′-ends (indicative of a defined TSS), others showed irregular patterns suggesting pervasive transcription or multiple TSSs  < 0.00002) and for appropriate control groups . In the enriched fraction, we observed a highly conserved motif YAAG , followed by an A-rich stretch of lower significance, whereas in the non-enriched fraction no preferences were found  were analyzed, only a YA motif22 was identified  in the top 100 and 200 enriched RNAs, and its prominence decreased with decreasing NAD captureSeq enrichment values  being A/T-rich between positions −30 and +10, (2) a switch from T-rich to A-rich in the coding strand around position −8, (3) a pyrimidine at position −1, and (4) an A at position +1. In addition, two specific features distinguish good NAD-incorporating promoters, namely a slightly increased probability for an A at position +2 and a strongly conserved G at +3.We supposed that analysis of the und Fig. . The motped Fig. . When foTDH3, a highly enriched NAD-RNA observed in every strain, and added a low-copy plasmid in which we inserted a DNA fragment containing the 600 bp upstream of the TDH3 gene, containing the entire promoter region, plus the first 54 bp after the experimentally observed TSS of the TDH3 RNA , followed by the ORF of superfold-GFP to monitor gene expression  reduction of relative NAD incorporation upon mutating positions −1 and +3, whereas for positions +2 and the A-rich tail the observed effects were not statistically significant  and negative control . After normalization to non-enriched species, in all eight libraries the sample group contained less full-length reads than the negative control and more abortive fragments  length of their 5′-ends (TL). This phenomenon was observed for ~20% of all RNA species and occurred in both directions, namely (apparent) TL lengthening and shortening upon knockout. For example, among the 1100 enriched sequences in common between the WT and TLs Fig. . For allTLs Fig. . This ph28, whereas for Dxo1 both nuclear and cytosolic locations were claimed29. Npy1 was described as a peroxysomal protein30. Localization microscopy using three different C-SWAT fluorescent protein fusions31 for each candidate gene revealed strong localized fluorescence in the nucleus for Rai1, whereas Dxo1 showed only a very weak and ubiquitous fluorescence   Fig. . These f7 had reported that NAD-RNA is not translated in human (HEK293T) cell extracts, based on a single mRNA luciferase construct with a single fixed TL sequence, while a recent study in the model plant Arabidopsis thaliana demonstrated that NAD-capped mRNAs are enriched in the polysomal fraction, associate with translating ribosomes, and can probably be translated9. No data for yeast have been reported yet. For seven different mRNAs, we prepared luciferase fusion constructs with long and short TLs by in vitro transcription, followed by the removal of the accompanying ppp-RNA by treatment with polyphosphatase and exonuclease Xrn-1. Although the control constructs harboring an m7G-capped 5′-end were efficiently translated in a yeast in vitro extract and showed significant differences in luminescence depending on the TL length20, NAD-capped RNA was not translated to any significant extent, even less than ppp-RNA and p-RNA of the same sequence . Our work confirms that there are hardly any full-length NAD-mRNAs, but additionally reveals a rich landscape of thousands of short NAD-mRNA fragments whose purpose is apparently not to encode for proteins.Taken together, our results indicate that in budding yeast, NADylation of RNAs is a very common phenomenon. A previous study reported only 37 species enriched in NAD captureSeq in budding yeast grown in the same medium32. Second, individual mRNA species in this organelle were found to be highly 5′-NAD modified, comprising up to 60% of the respective transcript pools. Third, the redox state of mitochondrial NAD caps was observed to vary, depending on the metabolic growth conditions of the cell32. These findings may indeed indicate a regulatory role of the NAD cap in this organelle, which harbors redox-intensive energy conversion pathways highly dependent upon the coenzyme, and are contrasting sharply with our discovery of a tightly policed landscape of nuclear-derived NAD-mRNA fragments.This conclusion may not apply in yeast mitochondria, however, where several lines of evidence suggest that transcriptional incorporation of the coenzyme is, in fact, an evolved feature. First, the mitochondrial transcription machinery exhibits NAD-mediated RNA initiation efficiencies that are at least tenfold higher, compared to the nuclear RNAP II34.In addition, the identity of a dedicated surveillance machinery within the mitochondrial matrix is of considerable interest. The three decapping enzymes investigated here, however, are unlikely to encounter mitochondrial transcripts, as they lack the required targeting sequence and are absent in yeast mitochondrial proteomic data36, the promoter sequence determines the efficiency of NAD incorporation, which is for most yeast nuclear transcripts between 1% and 5%. We observe that a YAAG motif supports efficient NAD incorporation by RNAP II in vivo, with the G at position 3 being particularly important. This finding does not rule out the existence of unknown alternative post-transcriptional pathways for NAD incorporation, e.g., for enriched snoRNAs or rRNAs, or for enriched mRNAs without the YAAG motif. In contrast to the promoter requirements determined here, a preference for HRRASWW was reported for E. coli RNAP36, WARR for B. subtilis RNAP5, and RA for yeast mtRNAP32, with the underlined A always indicating the TSS. It should, however, be noted that the E. coli RNAP and yeast mtRNAP consensus motifs were established using an entirely different methodology, making a direct comparison difficult. We find that, as a consequence of this promoter dependence, for many RNAs the NADylated species originate from different TSSs and have therefore different (shorter or longer) 5′-UTRs than the canonical ones. This phenomenon may modulate the secondary structure of these RNAs and hence their stability, molecular interactions, and biological fate. Of note, the discovery of alternative TSS selection and the YAAG core promoter motif have been made possible by the combination of 5′-end selection by ADPRC treatment and ligation-based attachment of the 5′-adapter, which allowed the determination of NAD-RNA 5′-ends with single-nucleotide precision, in contrast to random-primed library preparation methods that create heterogeneous ends. An increased affinity of RNAP II for the YAAG motif, while the polymerase transiently harbors NAD in its catalytic site, could potentially also explain the enrichment of non-canonical TSSs upon execution of NAD captureSeq. An in-depth biochemical investigation should explore the possibility of NCIN-mediated guidance of RNAP II, and other RNAPs, to distinct TSSs.NAD is initially incorporated into several thousands of transcripts by transcription initiation by RNAP II in a largely statistical manner reflecting the competition of NAD with the canonical initiator ATP. As in prokaryotes37. Alternatively, NAD-RNAs might be subject to accelerated degradation after transcription is complete, but it is unclear how 5′-NAD can accelerate degradation at the 3′-end.NAD-RNAs are—on average—shorter than non-NAD-RNAs and only rarely reach the size of a typical primary mRNA transcript. The most likely explanation is that some unidentified QC mechanism detects 5′-NADylation of RNA as an error early during transcription and prevents efficient elongation, as it does with uncapped or incompletely capped transcripts28 and to act in RNA surveillance by assisting the 5′- to 3′- exonuclease Rat1 in the co-transcriptional degradation of uncapped transcripts39, such an order appears plausible. The Rat1-Rai1 complex is hereby believed to play an important role by mediating 5′-end cap QC (5′-QC) in the yeast RNAP II transcription cycle, following the transcription checkpoint pause stage, whereby RNAP II enters transcription elongation upon phosphorylation of distinct serines within the C-terminal repeat domain of the polymerase40. Surveillance and hydrolysis of the accidentally incorporated 5′-NAD cap could be enacted in a mechanistically similar manner, mirroring the clearance of unmethylated, aberrantly capped mRNAs by the Rat1-Rai1 heterodimer41.The discovery that budding yeast maintains at least three different, partly redundant, pathways for NAD cap removal, using enzymes with different chemistry and cellular localization, implies that decapping unwanted NAD-RNAs is important for the cell. Our data are in agreement with the hypothesis that Rai1 acts earlier than the other two enzymes. As the nuclear protein Rai1 is known to associate with RNAP II during elongationSachharomyces cerevisiae Npy1 is not entirely restricted to NAD, but also includes the redox cofactor flavin adenine dinucleotide among others30, warrants a thorough biochemical characterization to define the set of RNA 5′-metabolite caps, targeted by this enzyme. A corresponding study should hereby follow the example set by the systematic and meticulous elucidation of RNA 5′-cap specificities of mammalian DXO and Schizosaccharomyces pombe Rai143.An earlier report, providing evidence that dinucleotide hydrolysis mediated by 7, hardly translate synthetic NAD-mRNAs, suggesting that the ribosomal machinery contains additional safeguards against NAD-mRNAs. Thus, budding yeast protects itself at different stages of gene expression against NAD-RNA.The observed combination of the low efficiency of RNAP II transcription initiation by NAD, the reduced length of NAD-RNAs, and the abundance of NAD-RNA decapping enzymes warrants that hardly any NAD-RNAs occur in the cell that could give rise to translation into proteins. Our data indicate, however, that yeast ribosomes, such as mammalian onesS. cerevisiae strains were grown in yeast extract/peptone/dextrose media (YPD). All strains used in this study except of YDK587-1 and its derivatives, YDK53-7, and C-SWAT mNeonGreen (mNG-I) strains were derivatives of the S288C strain BY4742 (MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0) and are listed in the key resource table. YDK53-7 and C-SWAT mNeonGreen (mNG-I) strains were derivatives of the S288C strain BY4741 (MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0) and YDK587-1 was a derivate of the S288C strain ESM356-1 (MATa ura3-52 leu2Δ1 his3Δ200 trp1Δ63). Strains were cultivated in YPD medium at 30 °C according to standard protocols. Antibiotics were used at the following final concentrations: 200 μg mL−1 geneticin (G418), 300 μg mL−1 hygromycin B, and 100 μg mL−1 nourseothricin. Gene deletions were performed using standard PCR-based recombination methods as described45, followed by PCR-based confirmation. Single, double, and triple mutants were generated by mating and sporulation followed by random spore isolation. Plasmid transformations were performed using standard methods47 and transformants were selected on synthetic complete (SC) medium lacking leucine for p415-based plasmids.Unless otherwise stated, 48 with minor changes. Cells were collected (OD600 0.8) from 0.5 L YPD medium, quickly frozen in liquid nitrogen, and stored at −80 °C until all samples were ready. Cell pellets were thawed on ice and washed with dH2O, resuspended in 8 mL TES solution  and were added to 8 mL phenol. The mixture was vortexed thoroughly for 1 min, incubated for 60 min at 65 °C with occasional shaking , and placed on ice for 15–20 min. Then the samples were centrifuged at 14,000 × g, 4 °C. The aqueous supernatant was subjected to phenol extraction, P/C/I purification, and chloroform purification. RNA was precipitated with 0.1 volumes 3 M NaOAc pH 5.5 and 2.5 volumes ethanol at −20 °C overnight. Precipitated RNA was dissolved in 2 mL dH2O. RNA concentration was determined by Nanodrop spectrometry and its integrity analyzed by 1.2% formaldehyde-denaturing agarose gel electrophoresis. RNA (1 mg) was treated with 100 U DNase I in 1X DNase I buffer (Roche) for 40 min at 37  °C. DNase I was removed by P/C/I extraction (twice), followed by ethanol precipitation of the RNA . The pellet was dissolved in 100 µL dH2O, the RNA concentration determined by Nanodrop, and its integrity analyzed by both denaturing agarose gel electrophoresis and Bioanalyzer RNA integrity analysis.Total RNA from yeast BY4742 strains was isolated by the hot phenol method11 with minor modifications, as outlined in the following. Total RNA, obtained in biological triplicates, was used as starting material for library preparation. More specifically, 100 µg total RNA was commonly supplemented with 5 ng NAD-RNAI (“spike-in” control) and treated with ADPRC , in the presence of 10 µL 4-pentyn-1-ol at 37 °C and for 60 min in 100 µL total reaction volume, including 50 mM Na-HEPES pH 7.0, 5 mM MgCl2. The same reaction mixture, omitting the ADPRC enzyme, served as background control (negative control). The reactions were stopped by adding 100 µL dH2O and 200 µL P/C/I reagent. This was followed by performing P/C/I extraction twice and three additional ether extractions. Subsequent copper-click reactions, captured by streptavidin bead, preadenylated-3′-adapter ligation, RT, Cytidine triphosphate(CTP)-tailing, anchored-5′-adapter ligation, and cDNA PCR amplification with barcode oligos were performed as standard library preparation11. Library QC was conducted by Sanger sequencing. Specifically, the amplicon derived from the NAD-RNAI spike-in control and other cDNA amplicons were amplified using corresponding adapter-ligated sequences. Resulting cDNA libraries were then cloned into plasmid(s) for Sanger sequencing. After the successful pre-sequencing, PCR products were purified and size-selected within a range of 150–300 bp by 10% native polyacrylamide gel electrophoresis (PAGE). The absence of primer-dimers was ensured by Bioanalyzer 2100 (Agilent) analysis using the Agilent High Sensitivity DNA Kit. Concentrations were measured using the Qubit approach. Primer-dimer-free libraries were then multiplexed at a final concentration of ~20 nM. Based on the cDNA length distribution, the library pools were either supplemented with 20% v3 Phix Control (Illumina) or custom Illumina sequencing primers, which bear three Gs at their 5′-end, preceded by the Illumina standard sequencing, to mitigate library imbalances before NextSeq 500 75 bp single-end (SE) sequencing.Standard, unfragmented NAD captureSeq: Total RNA was subjected to the NAD captureSeq protocol2CO3, 90 mM NaHCO3 pH 9.3) at 94 °C (5 min for WT strain and 20 min for npy1Δ strain to approach similar fragment size). Sheared RNA fragments were visualized on a 1.2% formaldehyde-denaturing agarose gel. Next, the sheared RNA was precipitated, in a total volume of 200 µL dH2O, by addition of 600 µL ethanol, 20 µL 3 M NaOAc pH 5.5, and 1 µL glycogen at −20 °C overnight. The precipitated RNA was washed with 75% ethanol and subsequently dissolved in 50 µL dH2O. An equivalent of 100 µg sheared RNA was then treated with 100 U T4 PNK, along with 0.1 mM ATP, 100 mM imidazole-HCl pH 6.0, 10 mM MgCl2, 10 mM β-mercaptoethanol, and 20 µg ml−1 RNase-free bovine serum albumin (BSA) in a total volume of 200 µL at 37 °C for 5.5 h. RNA extraction was performed twice, employing the P/C/I approach, and followed by triple ether extraction and ethanol precipitation. Precipitated RNA was the washed again, using 75% ethanol, and ultimately dissolved in 20 µL dH2O, yielding the library input for the standard NAD captureSeq protocol11. PCR products were size-selected within a range of 150 to 300 bp  and of 300–500 bp (referred to as “large fragmented NAD captureSeq library”), enabled by 10% native PAGE. Bioanalyzer QC, library multiplexing, and the overall sequencing strategy were executed in a similar manner, as done for the unfragmented NAD captureSeq. Again, the NextSeq 500 75 bp SE approach was chosen for sequencing.Fragmented NAD captureSeq: Biological triplicates of total RNA (gDNA free) were used as library starting material. Total RNA (100 µg), supplemented with 5 ng NAD-RNAI , were randomly sheared in a 65 µL reaction volume that contained 32.5 µL alkaline fragmentation solution  served as library input material. Total RNA (1 µg) was subjected to rRNA depletion by Ribo-Zero rRNA Removal Kit (yeast). rRNA-depleted RNA was randomly sheared in 10 µL dHn and 3′-end adapters (C NNNNNN AGATCG) were trimmed (minimum length 12 nt) by in-house scripts. Reads that mapped to the spike-in internal standard (IS) RNAI sequence  were then counted first. IS unmapped reads were subsequently classified as “small RNA reads” (12–17 nt) or “normal RNA reads” (>18 nt).Unfragmented NAD captureSeq analysis: Original reads were demultiplexed, based on the PCR barcode, not allowing for mismatches and subjected to standard QC procedures. Further 5′-end leading (G/N)S. cerevisae BY4742 strain  (bowtie -v 2). Normal RNA reads that mapped to rRNA genes or tRNA genes were separated and counted individually. Remaining reads that could not be mapped to rRNA genes were remapped to the yeast reference genome (bowtie –best --strata -M 1 -m 20 -v 2). The.sam files were then converted to.bam files and sorted . The.bam files were used to generate.wig files, which could be normalized by reads per million mapped reads, for single-nucleotide resolution-based analysis. Sorted.sam files were further filtered by strand-specific selection and used for three different types of analysis as follows:Normal RNA reads were mapped to the reference genome 49 to perform an alternative TSS study. Briefly, the position of the first nucleotide of the mapped reads within the range of the 5′-UTR18 with additional 50 nt upstream extension from each gene was collected. A sliding window of 18 nt was applied for searching for TSS clusters from the highest density one to the lowest one. Then a 2 × n table was made, whereby n is the number of unique TSS clusters found in the sample group  and/or the negative control group . Values in the table represent the number of reads for these clusters in the S or N group , and are sorted from the shortest (left) to the longest UTR (right). For covariance analysis, the top row (S) was assigned a weight value of 2, whereas the bottom row (N) was assigned 1. Column  weight was set to be the 5′-UTR length of each cluster. The Pearson’s correlation r was calculated by cov /(δx × δY). The operator M was calculated based on the equation M2 = (n − 1) r2 (n ≥ 30), following the χ2-distribution (freedom 1). The corresponding p-value was used for FDR calculation, employing the Benjamini–Hochberg method.On the one side, the filter-passed reads from sorted.sam files were used for linear trend testsk) was the signal intensity from the ultra performance liquid chromatography (UPLC)/MS measurement of total RNA. The distribution of all-cap-RNA in total RNA was simulated by their TPM in transcriptome data. The distribution of NAD-RNA was simulated by their TPM in NAD captureSeq data. The relative individual NAD-ratio was calculated by k*TPMNAD-RNA/TPMall-cap-RNA.On the other side, the filter-passed reads were counted for RNA hits  based on the annotation.gff3 file  was defined as TSS (reference as +1) but this position was annotated as “A” in the genome. The sequence flanking the TSS from the −10 to +10 position, within the reference genome, was further analyzed for a consensus motif. The sharp value was defined as the ratio between the number of nucleotides accumulating at the +1 position and that at the −1 position, revealed by the generated.wig file. The sharpA feature was screened, based on the premise that the TSS constitutes an A and possesses a sharp value bigger than 4.In addition, the filter-passed reads, which mapped to 5′-UTRs of mRNAs, were collected for For the small RNA reads group, reads were purged by additional genome mapping (bowtie -v 0). Next, small RNA clusters mapping to mRNA 5′-UTRs were assembled together . Differential abundance of small RNA reads was calculated by the enrichment of reads in S group to N groups.Fragmented NAD captureSeq was analyzed in the same way, as done for the unfragmented NAD captureSeq library.Transcriptome analysis: Raw reads were demultiplexed and QC was performed using the same parameters, already employed for unfragmented NAD captureSeq analysis. Reads with 5′-/3′-end adapters were properly trimmed, leveraging an in-house script. Then the sequence of raw reads was converted into its reverse complement. Reads were then mapped to the yeast reference genome . rRNA genes and tRNA gene reads that mapped independently, as well as rRNA-free hits counting (same annotation region on 5′-UTR of mRNA), were treated in a similar way as done in the NAD captureSeq analysis, described above. Also, .wig files and differential expression statistics were generated and performed according to the same procedure.Function clusters were generated by David (version 6.7) and FGNet (version 3.16.0). Enriched promoter motifs were analyzed by MEME (version 5.0.3) and WebLogo (version 3). Meta-analysis of function clusters and pathways was enabled by Metascape (metascape.org) and cytoscape (version 3.7.0).2O, to remove unspecifically binding RNAs. RNA was eluted by adding four times 200 µL 2 mM EDTA  under 10 s per min shaking (350 r.p.m.) at 75 °C for 10 min. The eluate was precipitated with 0.5 M ammonium acetate pH 5.5 and 50% isopropanol. Precipitated RNA was dissolved in dH2O for UPLC/MS analysis.Streptavidin Sepharose High Performance beads (250 µL) were loaded on Mobicol Classic columns. The column was washed three times with 1× phosphate-buffered saline (PBS) buffer, then five times 25 µL of 25 µM biotin-DNA probe , resulting from the NudC treatment of NAD-RNA and d4-NR as IS. It was collected and dried under vacuum. The amount of NR was determined by UPLC-MS/MS and reflects the exact same amount of original NAD-RNA in the digested sample. The employed UPLC-MS/MS setup contained a triple stage quadrupole mass spectrometer  system coupled with an Acquity UPLC system. A BEH Amide column  was used with an eluent A (0.01% (v/v) aqueous formic acid with 0.05% ammonia and 5% acetonitrile) and B (acetonitrile included 0.01% formic acid), at a flow rate of 0.8 mL min−1. The gradient started to change from 12.5% A/87.5% B to 95% A/5% B within 1.8 min. The ratio was changed back to starting conditions within the following 1.0 min. The column was had been pre-equilibrated under starting conditions for 1.0 min. Electrospray ionization was performed with a 1500 V capillary voltage, 11 V cone voltage, 150 °C source temperature, 200 °C desolvation temperature, 150 L h−1 cone nitrogen gas flow, and 800 L h−1 desolvation gas flow (N2). The Xevo TQ-S was automatically tuned to d4-NR and NR using the MassLynx V4.1 system software (Waters) with the IntelliStart standard procedures. Multiple reaction monitoring measurements were conducted, using collision gas  for collision-induced decomposition and MS/MS transitions were monitored in the positive ion mode .To determine the amount of NAD that is covalently linked to RNA, the RNA samples were washed three times with 400 μL of 8.3 M urea, one time with dH7G-RNA, and p/ppp-RNA from each other. Briefly, 0.5% (w/v) APB (Lab stock), 10% acrylamide/Bis solution (19:1), 0.1% (w/v) APS, and 0.1% (v/v) TEMED with 2.5× Tris-acetate-EDTA (TAE) buffer in 50 mL volume were mixed and poured between glass plates (Bio-Rad). Gels were run at a stable current (15–29 mA per gel) in 1× TAE buffer. Gels were then stained with SYBR Gold or gels, containing 32P-labeled nucleic acids, were exposed to storage phosphor screens  and visualized using a Typhoon FLA 9500. Signal quantification was performed using the ImageQuant software .Native PAGE (10%) was utilized to size select cDNA for NGS. Briefly, 10% acrylamide/Bis solution (19 : 1), 0.1% Ammonium persulfate(APS) (w/v), and 0.1% N,N,N′,N′-Tetramethylethylenediamine(TEMED) (v/v) along with 1× Tris-borate-EDTA (TBE) in 50 mL volume were mixed and poured between glass plates (19 cm × 27 cm). Gel mixtures were polymerized at room temperature for 45 min. The electrophoresis conducted at a stable 27 mA current for 2.5 h. The gel was then stained with SYBR Gold (Thermo Scientific) in 1× TBE buffer, 5 min. The signal intensities were read out by scanning the gel at 400 V, 50 or 100 µm resolution using a Typhoon FLA 9500. The printout picture of the gel  was used for excision of desired size ranges within the corresponding gel lanes. APB gel electrophoresis was utilized to separate NAD-RNA, m2, 0.01% Triton X-100, 10 mM dithiothreitol (DTT), 5% dimethyl sulfoxide, along with 60 µCi α-32P-ATP, 10 µCi α-32P-CTP, 4 mM each CTP/GTP/UTP, 2 mM ATP, 6 mM NAD (NAD-RNA) or without NAD (ppp-RNA), 0.7 µM T7 polymerase (self-prepared) within a total reaction volume of 100 µL. The in vitro transcription reaction was incubated at 37 °C for 3 h. Subsequently, 10 U DNase I were added and the reaction incubated for an additional 30 min at 37 °C. ppp-RNA was purified by 10% denaturing PAGE, as described above. NAD-RNA was first purified by 10% denaturing PAGE, followed by a 10% PAGE, supplemented with 0.5% APB, to remove ppp-RNA. For p-RNA preparation, 1 µg ppp-RNA was treated with 20 U RNA 5′-polyphosphatase and 1× Reaction Buffer (Epicentre) in a reaction volume of 20 µL for 1 h at 37 °C. The desired p-RNA species was then obtained by performing P/C/I extraction twice, followed by ethanol precipitation, as described. To generate m7G-RNA, ~1.2 µM denatured ppp-RNA  was first added to a mixture of 1× Scriptcap capping buffer, 1 mM GTP, fresh 0.1 mM S-adenosyl methionine (SAM), and 1 U µL−1 Script Guard RNase Inhibitor. The mixture was then supplemented with 0.4 U µL−1 Scriptcap Capping Enzyme, in a final reaction volume of 50 µL and incubated for 50 min at 37 °C. The modified RNA reaction was then purified by 10% APB-PAGE as described above, to separate uncapped, ppp-RNA, and m7G-capped RNA.Radio-labeled RNA: 400 nM double-stranded DNA (dsDNA) served as a template in a transcription buffer containing 40 mM Tris-HCl pH 7.9, 1 mM spermidine, 22 mM MgCl2O to remove free NTPs and small molecules. Retained RNA was collected and again precipitated with ethanol. Recovered RNA was then washed twice with 75% ethanol and ultimately dissolved in 20 µL dH2O. For further isolation of 5′-NAD-modified RNA, 5 µg partially purified NAD-RNA was treated with 5 U RNA 5′-polyphosphatase in 1× Reaction Buffer (Epicentre) with an overall volume of 10 µL for 40 min at 37 °C. Remaining NAD-capped RNA was then P/C/I extracted twice and ethanol precipitated. Precipitated mRNAs were further treated with 2 U Xrn-1 in 1× NEBuffer 3 (NEB) reaching a total reaction volume of 40 µL. For p-mRNA preparation, 5 µg ppp-mRNA was treated with 10 U RNA 5′-polyphosphatase in 1× Reaction Buffer (Epicentre) and final volume of 20 µL for 40 min at 37 °C. For m7G-mRNA preparation, 1 µM denatured ppp-mRNA  was mixed with 1× capping buffer (NEB), 0.5 mM GTP, fresh 0.1 mM SAM, and 0.5 U µL−1 Vaccinia Capping Enzyme in 20 µL volume, incubate at 37 °C, 40 min. Processed RNA was purified twice by P/C/I extraction and precipitated with ethanol. All mRNA raw concentrations were measured by nanodrop, followed by qRT-PCR to determine the relative abundance of the mRNA middle region, as well as of the 3′-end region. Final concentration of mRNAs was adjusted accordingly.Luciferase mRNA: 140 nM (~6 µg) linear dsDNA template was added into the same transcription buffer as described above, along with 4 mM each CTP/GTP/UTP, 2 mM ATP, 6 mM NAD (NAD-mRNA) or without NAD (ppp-mRNA), 0.7 µM T7 polymerase (self-prepared) with a total reaction volume of 100 µL. The reaction mixture was incubated for 3 h at 37 °C. Similarly, 10 U DNase I were added subsequently to the reaction and incubated for an additional 30 min at 37 °C. RNA integrity was examined by visualization of the reaction products on a 1% formaldehyde-denaturing agarose gel. RNA was purified by performing P/C/I extraction twice and precipitated with ethanol. Precipitated RNA was loaded onto Amicon Ultra-0.5 mL Centrifugal Filter Units 10 kDa and washed four times with 400 µL dH50. For imaging the fluorescence of cell colonies and cell patches on plates, a custom-made fluorescence illumination cabinet was used, equipped with filters for GFP.For fluorescent live cell imaging, a DeltaVision Elite widefield fluorescence microscope  consisting of an inverted epifluorescence microscope  equipped with a light-emitting diode light engine , an sCMOS camera , and a ×60 1.42 NA Plan Apochromat N oil-immersion objective (Olympus) was used. Cells were inoculated in low-fluorescence medium  and grown to mid-log phase. Cells were immobilized for imaging in glass-bottomed 96-well plates  using Concanavalin A and fluorescence filters for GFP, as standard procedure51 with minor modifications. Cell pellets were obtained from 0.5 L YPD yeast (BY4742 strain) culture (OD600 ≈ 15). The cell pellet was washed three times with 50 mL ice-cold Breaking Basic Buffer 2 pH 7.0, 2 mM DTT), supplemented with 8.5% (w/v) mannitol. Then, the wet weight of the cells was determined. The cells were subsequently resuspended in Breaking Basic Buffer with 8.5% mannitol and 0.5 mM phenylmethylsulfonyl fluoride (PMSF) , under addition of 1.5 mL per 1 g wet weight. The solution was supplemented with ice-cold, sterile ~0.5 mm glass beads, whereby the equivalent of six times of the cell wet weight was added. The cells were raptured by manual shaking , in overall five rounds, each consisting of 1 min shaking interrupted by 1 min cooling on ice. Glass beads were removed by low speed centrifugation , then the cleared supernatant was obtained by two consecutive rounds of centrifugation . The solution was filtered through a 0.45 µm filter and then 2 mL of the supernatant were subjected to Fast Protein Liquid Chromatography (FPLC) runs, outlined below. HiPrep 26/10 Desalting Columns (containing Sephadex G-25 Fine resin) were pre-equilibrated with 100 mL Breaking Basic Buffer, supplemented with 0.5 mM PMSF. The injected sample was resolved by the indicated column matrix running on a FPLC system  employing the same equilibration buffer. Absorption values of each fraction were determined at 260 nm (A260) and appropriate fractions, exceeding 75% of the highest A260 value, were pooled together. Next, 1 mM CaCl2 and 50 U mL−1 micrococcal nuclease were added to the pooled fractions, followed by incubation at 26 °C for 15 min. The reaction was stopped by adding EGTA to a final concentration 2.5 mM. Aliquots of 100 µL each were then flash-frozen in liquid nitrogen and stored at −80 °C.Preparation of cell-free extracts for in vitro translation was performed as described−1 creatine phosphokinase, 2.5 mM DTT, 125 mM KOAc, 5 mM MgOAc pH 7.0, 25 µM Amino Acid Mixtures, 1 U µL−1 RNas in Ribonuclease Inhibitors, 80 nM m7G-capped Renilla mRNA, and dH2O in a final volume 80 µL. The solution was gently mixed and 4 µL of the Master Translation Solution aliquoted to individual PCR tubes. Then, 1 µL mRNA (200 ng or accordingly) and 5 µL of the cell-free extract were added to thus prepared reaction volumes. The “ready-for-translation” solution was again mixed gently and incubated at 26 °C for 30 min. Next, 90 µL dH2O were added to each in vitro translation reaction. Seventy five microliters of this dilution were then used to conduct a firefly luciferase activity assay (Bright-Glo Luciferase Assay System). The remaining 25 µL, with additional 50 µL of dH2O, were subjected to a Renilla luciferase activity assay (Renilla-Glo Luciferase Assay System), executed according to the manufacturing instructions. Emitted luminescence was read out using a TECAN plate reader.In vitro translation: Master Translation Solution was freshly prepared by mixing 25 mM HEPES-KOH pH 7.6, 1.25 mM ATP, 0.125 mM GTP, 0.15 U µLn = 4). One milliliter of these cultures, at an OD600 of ~0.8, were pelleted by the centrifugation at 4000 × g, 1 min, 4 °C. The cell pellets were washed four times with 1 mL ice-cold PBS and resuspended in 1 mL NADH/NAD Extraction Buffer. The resuspension (400 µL) were subjected to three freeze–thaw cycles, each consisting of 10 min on dry ice, alternating with 10 min thawing at room temperature. The homogenized cell lysates were vortexed for 10 s and centrifuged at 14,000 × g, 4 °C, 10 min. Twenty microliters of each supernatant were aliquoted for subsequent lysate RNA quantification. The remaining solution was applied onto a 10 kDa spin filter to remove proteins larger than 10 kDa, by centrifugation at 14,000 × g, 4 °C, 10 min. The flow-throughs were collected and NAD/NADH quantification was conducted, as stated by the manufacturer. The determined NAD amounts were normalized by the amount of measured lysate RNA.The general experimental procedure was based on the manufacturer’s protocol (NAD/NADH Quantification Kit) with minor modification. Yeast cells were cultured in 50 mL YPD medium in biological quadruplicates  and incubated at 37 °C for 120 min. Samples that were treated with the E. coli Nudix hydrolase NudC are referred to as “positive control.” Aliquots of 5 µL were taken from the reaction mixtures at indicated time points. Thus, treated RNAs were mixed with the same volume of 2× APB Gel Loading Buffer  and placed on ice for further APB gel electrophoresis. Reaction mixtures, containing NAD, were stored on ice before performing thin-layer chromatography . Resolution of nucleotides via TLC, at room temperature for 5.5 h, was achieved employing a flow phase of 1 M NH4OAc and ethanol (4 : 6).In general, 11. For RT on beads, to each sample of the fully treated group, as well as the background group, were added 2.5 µM random hexamers, 0.5 mM dNTP mix, and dH2O at 65 °C for 5 min. After reaction, the beads were incubated on ice for 2 min. Then, to the reaction was added 1× SSIV Buffer (Thermo Scientific), 5 mM DTT, 10 U µL−1 SuperScript IV Reverse Transcriptase, and 50 ng µL−1 acetylated BSA (Sigma-Aldrich), and the mixture incubated at 23 °C for 10 min, followed by 1 h at 55 °C. This was followed by RNA rebinding to streptavidin beads, washing, NaOH-mediated hydrolysis, and cDNA precipitation, as described in the NAD captureSeq protocol11. Equivalents of 1 µg of diluted non-NAD-RNAs were reverse transcribed, employing random hexamer oligos, as described above. The relative abundance of transcripts was then quantified by RT-qPCR. The enrichment of transcripts, assessed by RT-qPCR of cDNA, from the fully treated group, the background group, and the non-NAD-cap group was normalized by rRNA, RDN5-1. The NAD-modification ratio of each RNA species was determined by the equation NAD-modification ratio = (NAD-RNAfully treated group − unspecific-binding-RNAbackgroud group)/(NAD-RNAfully treated group + other-cap-RNAnon-NAD-cap group).For each assay, 100 µg total RNA (gDNA free) was subjected to ADPRC treatment as fully treated group. An equal amount of total RNA was applied to the same treatment without ADPRC as background group. The subsequent copper-click reaction, capture by streptavidin beads (streptavidin-unbound flow-through RNAs (non-NAD-RNA) were collected and precipitated with ethanol), as in the standard NAD captureSeq procedure−1, Sigma) in 100 µL Immobilization buffer at room temperature for 10 min under gentle agitation. Subsequently, beads were washed three times with 150 µL Immobilization Buffer. Precipitated RNA from the copper-click reaction was dissolved in 100 µL Immobilization Buffer and then added onto the washed beads. The mixture was incubated at room temperature, while shaking at 500 r.p.m. for 1 h. Next, the 96-well plate was placed on a magnetic rack at room temperature for 10 min. Then, the first supernatant was collected. Beads were washed with 100 µL Immobilization Buffer and following that, the second supernatant was collected and pooled with the first one, subsequently referred to as “non-NAD-RNAs.” The RNA from the pooled supernatants was precipitated and 1 µg of diluted non-NAD-RNAs was utilized for RT to determine transcript abundance by RT-qPCR. RNAs captured by magnetic streptavidin beads  were washed five times with 150 µL Streptavidin Wash Buffer  and then washed three times with 150 µL First strand Buffer . Subsequently, the beads were re-blocked using 100 µL First strand Buffer, containing acetylated BSA (100 µg mL−1). Then, equilibrated beads were washed twice with 150 µL 1× SSIV buffer (Thermo Scientific). RT was conducted using random hexamer oligos. The reactions were then transferred into PCR tubes and heated to 65 °C for 5 min to denature the RNA. The tubes were cooled on ice for 5 min, then incubated at 23 °C for 10 min, followed by a heating step at 55 °C for 30 min to enable the RT reaction by Superscript reverse transcriptase IV. Next, 100 µL NaOH (0.15 M) were added into the reaction and the mixture incubated at 55 °C for 15 min. The first RT supernatant was collected. The remaining beads were repeatedly subjected to the same treatment, by adding 100 µL NaOH (0.15 M). Then the second, residual RT supernatant was pooled with the first one and is subsequently referred to as NAD-RNA (S group) and background RNA (N group). Both were ethanol precipitated and directly subjected to RT-qPCR. The abundances of NAD-TDH3 RNA and non-NAD-TDH3 RNA were normalized by transcript abundances of the NAD-RNAIII and ppp-RNAI controls, respectively. The NAD-capping ratio of TDH3 was calculated by dividing NAD-TDH3 by the sum of NAD-TDH3 and non-NAD-TDH3.For the NAD-TDH3 promoter assay, 100 µg total RNA (gDNA free) were supplemented with 5 ng of NAD-RNAIII (self-prepared) and 5 ng ppp-RNAI (self-prepared), and subjected to the same ADPRC treatment and copper-click reaction, as described above. Then, 50 µL of Hydrophilic Streptavidin Magnetic Beads slurry per reaction was utilized to enrich for NAD-modified RNAs in a 96-well plate  format. The beads were hereby pre-washed twice with 150 µL Immobilization Buffer  and then blocked using acetylated BSA  served as reference RNA. The reactions were stopped by adding an equal volume of 2× APB Gel Loading Buffer. Separated and cleaved products of NAD-RNA and ppp-/p-RNA were visualized by APB-supplemented PA gel electrophoresis and subsequent SYBR Gold staining. The gel was scanned using a Typhoon imager for further signal quantification. For those samples with weak signal intensities upon DNAzyme treatment on APB gel, RT-qPCR was performed to quantify RNA abundance.For in vitro-transcribed NAD/ppp-mRNAs, 200 ng of denatured (~375 fmol) NAD-mRNAs were supplemented with 10 mM Tris-HCl pH 8.0, 50 mM NaCl, 2 mM fresh DTT, and 1 µM DNAzyme , 40 cycles of consecutive annealing/extension steps at 95 °C for 7 s and 60 °C for 15 s, respectively. Melting curves were generated by heating from 65 °C to 95 °C with an incremental increase of 0.5 °C s−1 hot-start Q5 high-fidelity DNA polymerase in a total volume 50 µL. The PCR was initiated by heating to 98 °C for 40 s and followed by 5 cycles . The second round of the PCR aimed to specifically amplify the bridged 5′-UTR/CDS, bearing the Luc2 template extension, by additionally adding corresponding 500 nM Fw_5UTR primer, 500 nM Rev_Luc2_polyA primer, and 1× Q5 Reaction Buffer (NEB) to the final volume of 56.2 µL.Two rounds of PCR were carried out to generate linear DNA templates for in vitro translation reactions as follows: the first round aimed to bridge the 5′-UTR/CDS to the firefly Luc2 sequence. The reactions contained 6.4 pM 5′-UTR/CDS template, 6.4 pM bridge region DNA .n ≥ 3) with SDs, and Student’s t-test (single-tail and double tail (only FCM data), unequal variance) was calculated by R/python. For NGS TSS switching relevant math, covariance with two variables, linear trend test, and corresponding p-values, as well as FDR value, were calculated using python scripts. For NGS NAD captureSeq, NAD-RNA enrichment and transcriptome differential expression analysis, DESeq2 was utilized, as described above.All samples for NAD captureSeq, transcriptomics, UPLC-MS, RT-qPCR, and proteomics were prepared as biological triplicates. The outlined in vitro experiments, including NAD kinetics, RNA decapping kinetics, and in vitro translations, were performed as technical triplicates. Mean values (Further information on research design is available in the Supplementary InformationPeer Review FileReporting Summary


Hopefully, it will accelerate the progress to develop medicines or vaccines to control the pandemic of COVID-19.The recent outbreak of COVID-19 caused by a new zoonotic origin coronavirus (SARS-CoV-2 or 2019-nCoV) has sound the alarm for the potential spread of epidemic coronavirus crossing species. With the urgent needs to assist disease control and to provide invaluable scientific information, we developed the coronavirus database (CoVdb), an online genomic, proteomic and evolutionary analysis platform. CoVdb has brought together genomes of more than 5000 coronavirus strains, which were collected from 1941 to 2020, in more than 60 countries and in hosts belonging to more than 30 species, ranging from fish to human. CoVdb presents comprehensive genomic information, such as gene function, subcellular localization, topology and protein structure. To facilitate coronavirus research, CoVdb also provides flexible search approaches and online tools to view and analyze protein structure, to perform multiple alignments, to automatically build phylogenetic trees and to carry on evolutionary analyses. CoVdb can be accessed freely at  Coronaviridae is a group of positive-sense, single-strand RNA viruses with a likely ancient origin, and human coronavirus repeatedly emerged during the past hundred years . CoronavThe aim of the construction of CoVdb is to provide coronavirus knowledge, to contribute to global coronavirus research, especially for the investigation of the emerging SARS-CoV-2. For previous works, ViPR  and ViraCoronavirus sequences and annotations are downloaded from the NCBI nucleotide database . We chosE-value <0.05 and only kept the one that has the highest score in alignments. Using matched UniProt accession numbers, we retrieved detailed proteomic information from UniProt. We looked for each gene’s possible protein 3D structure counterparts in the Protein Data Bank .We did subcellular localization prediction for coronavirus gene clusters using an online tool MSLVP . We usedWe utilized CUDA ClustalW  to perfoA semi-automatic pipeline was developed to update the database, considering the fast increase of coronavirus’ sequences in NCBI. For specifics, the pipeline first refreshes the strain list according to NCBI’s coronavirus list (requiring complete genome), then updates corresponded annotation and finally make revision for some overall analysis results.Similar with what we have done for SGID  and ASFVRhinolophus affinis, a species of bat in the Rhinolophidae family. Strains isolated from pangolins are also in the vicinity of SARS-CoV-2. Pangolin was once considered as a potential intermediate host of SARS-CoV-2  into 628 clusters . This number indicates that the differentiation or diversity within coronavirus strains is not low. For these, we still performed a subcellular localization analysis for the 628 clusters to predict their roles in infection, although the structure of coronavirus is not complex. Based on prediction only, 21% (133 items) are predicted to be located in the host nucleus or host cytoplasm, while 40% (250 items) are predicted to be membrane proteins . CoVdb hFor all coronavirus strains, using nine representative human coronavirus genomes as the reference, we did sliding window analyses on Pi , Tajima’The genome browser (GBrowser) in CoVdb follows a style with gene segments followed by analysis tracks . Users can view population genetic tracks of strains belonging to one or more hosts, such as avian, bat and so on. They can also view that of strains belonging to one or more specific diseases, such as COVID-19, SARS and 229E. Users can select by checkboxes. At the top of GBrowser, there are browsing tools, such as search by inputting a chromosome position, zoom in/out and position movements. It provides notes for whether a gene of ORF is already annotated in GeneBank or newly predicted in the top of the genome browser page. In addition to basic information, CoVdb shows a gene in function, subcellular localization, topology and protein structure. The search engine in CoVdb is powerful and supports fuzzy search, search with taxonomy , filtering, sorting, BLAT and BLAST. CoVdb also allows to search by cell location. For personalized analyses, CoVdb is able to provide gene links if inputting a list of chromosome positions or gene accessions.CoVdb has tools to facilitate some specific use in coronavirus research, such as tracing origination, vaccine or drug design. In the tool ‘Protein’, the protein structure information is listed, where users can view the overlapped amino acids of a coronavirus protein in the 3D structure counterpart and do online protein structure analysis by an embedded application iCn3D  Figure . Users aThe spike glycoprotein (S protein) plays a key role in the infection of COVID-19 while the receptor-binding domain (RBD)  is the rDedicated to assist researchers to combat the pandemic of COVID-19 and to provide a more specialized platform for coronavirus, we comprehensively gathered data and systematically constructed the coronavirus database, CoVdb. In the database, researchers can conveniently retrieve genomic or gene information of coronavirus and do online analyses in comparative genomics, protein structure and evolutionary biology. With the help of this database, we have successfully developed test strips able to detect SARS-CoV-2 (unpublished). With the increase of the number of sequenced coronavirus genomes, we will provide continuous update and maintenance of the database in the future. Hopefully, this database will play more important roles in fighting against the infection of coronavirus in the future.baaa070_SuppClick here for additional data file.


Tomato bushy stunt virus (TBSV) replication. However, it is currently unknown what features make a particular DEAD-box helicase either pro-viral or antiviral. In this work, we succeeded in reversing the viral function of the antiviral DDX17-like RH30 DEAD-box helicase by converting it to a pro-viral helicase. We also turned the pro-viral DDX3-like RH20 helicase into an antiviral helicase through deletion of a unique N-terminal domain. We demonstrate that in the absence of the N-terminal domain, the core helicase domain becomes unhinged, showing altered specificity in unwinding viral RNA duplexes containing cis-acting replication elements. The discovery of the sequence plasticity of DEAD-box helicases that can alter recognition of different cis-acting RNA elements in the viral genome illustrates the evolutionary potential of RNA helicases in the arms race between viruses and their hosts, including key roles of RNA helicases in plant innate immunity. Overall, these findings open up the possibility to turn the pro-viral host factors into antiviral factors, thus increasing the potential antiviral arsenal of the host for the benefit of agriculture and health science.Positive-stranded (+)RNA viruses greatly exploit host cells to support viral replication. However, unlike many other pathogens, (+)RNA viruses code for only a limited number of genes, making them highly dependent on numerous co-opted host factors for supporting viral replication and other viral processes during their infections. This excessive dependence on subverted host factors, however, renders (+)RNA viruses vulnerable to host restriction factors that could block virus replication. Interestingly, cellular ATP-dependent DEAD-box RNA helicases could promote or inhibit the replication of  The largest group of eukaryotic viruses, the positive-strand RNA viruses, depends greatly on co-opting host components to support their replication. This dependence on host factors by these viruses also makes them vulnerable to antiviral factors. This is well-illustrated in case of tombusviruses, a small RNA viruses of plants. Tombusviruses co-opt many host factors to support various steps in their replication. Among these host factors are cellular DEAD-box helicases, which help remodeling viral RNA structures during the RNA replication process. However, similar cellular helicases remodel the viral RNAs incorrectly, making them antiviral or restriction factors. To gain insights into what makes a particular DEAD-box helicase pro-viral or antiviral, in this work, we converted the antiviral plant RH30 helicase into a pro-viral helicase through modifying the N-terminal sequences. We also succeeded to turn the originally pro-viral plant RH20 helicase into an antiviral helicase using a similar strategy. By characterizing the newly acquired functions of these helicases, we obtained valuable insights into what features make these helicases either pro-viral or antiviral. These discoveries have implications to better understand the arms race between viruses and hosts. In addition, it opens up the opportunity to generate new antiviral tools by converting pro-viral host factors into antiviral factors, thus enhancing our molecular tools against the ever-evolving RNA viruses. Positive-stranded (+)RNA viruses exploit host cells by co-opting many cellular factors to facilitate viral replication. In addition, many (+)RNA viruses take advantage of metabolic pathways, subcellular membranes and intracellular trafficking to build elaborate membranous viral replication compartments , which are the sites of viral replication, in the cytosol of the infected cells –8. The eTomato bushy stunt virus (TBSV) and other tombusviruses infecting plants. UTP (NEB) in a total of 20 μl reaction volume. In addition, about 3.8 μM of purified recombinant proteins was also added in the reaction. The reaction was performed at 25°C for 3 h and then stopped by the addition of 5 volumes of 1% SDS and 5 mM EDTA, followed by phenol-chloroform extraction and RNA precipitation. Then the repRNA products and dsRNA intermediates were analyzed by electrophoresis in 0.5X Tris-borate-EDTA (TBE) buffer in a 5% polyacrylamide gel (PAGE) containing 8 M urea.The yeast cell-free extracts (CFE) that support escribed ,54. The in vitro transcription. On the other hand, 32P-labeled ss RI(-), RII(+),RIII(-) or RIV(+) RNAs was synthesized by T7-based in vitro transcription along with 32P-labeled UTP, respectively. To prepare partial dsRNA duplexes , 2 pmol of 32P-labeled ssRNA were annealed to 6 pmol of unlabeled repRNA in STE buffer  by slowly cooling down the samples  from 94°C to 25°C in 30 min in a PCR machine. An increasing amounts  of purified MBP fusion proteins or GST fusion proteins were added to the partial dsRNA duplexes in the RNA binding buffer  supplemented with 1 mM ATP, followed by incubation at 25°C for 25 min. The reaction mixtures were then treated with Proteinase K (2 μg/ per reaction) at 37°C for 20 min, followed by analysis with nondenaturing PAGE containing 5% polyacrylamide.To study if the purified recombinant proteins could separate partial dsRNA duplexes, the dsRNA separation assay was performed as described . First, in vitro T7 transcription supplemented with 1.5 mM of rATP, rCTP and rGTP as well as 0.75 mM rUTP along with 0.07 mM biotin16-UTP (Roche). Approximately 5.7 μM of recombinant MBP-RH30, MBP-RH30ΔN/ΔC, GST-RH20 and GST-RH20ΔN2-96, respectively, were incubated with 0.1 μg of biotinylated RNA, 0.1 μg of tRNA, 2 U RNase inhibitor, and 1 mM ATP or no ATP in the presence of biotin-RNA binding buffer  in a total of 10 μL reaction volumes at 25°C for 15 min. Non-biotinylated RNA composed of the RII(+) of DI-72, MBP protein and GST protein were used as controls, respectively. After that, about 3.8 μM of the recombinant MBP-p33C was added to the reactions and incubated at 25°C for 15 min. The reaction mixtures were then incubated with 20 μL of streptavidin magnesphere paramagnetic particles (VWR) at room temperature for 20 min. The particles were captured by a magnetic stand and washed with biotin-RNA binding buffer for five times. The ribonucleoprotein complexes were eluted with 20 μL of SDS-PAGE loading dye containing β-mercaptoethanol by boiling for 15 min. The obtained samples were analyzed by western blot analysis with anti-MBP or anti-GST antibodies.To test if the RNA helicases and their mutants could interfere with viral template recognition by TBSV p33, we performed a biotinylated RNA-protein interaction assay as previously described  with a m6-tagged p92 from the chromosome  Ypic for 3 h at 4°C. The supernatant of detergent-solubilized membranes was obtained by centrifugation at 35,000 g for 20 min at 4°C and then was incubated with anti-FLAG M2-agarose affinity resin (Sigma) in columns for 16 h at 4°C. The columns were washed with high-salt buffer for three times. For elution, the protein samples in the columns were incubated with SDS-PAGE loading dye at 85°C for 6 min and were collected by centrifugation at 500 g for 3 min. The collected samples were then boiled with β-mercaptoethanol for 20 min. Affinity-purified Flag-p33 was analyzed by western blot with anti-FLAG antibody, and co-purified 6xHis-tagged proteins was analyzed by western blot with anti-His antibody  yeast protease inhibitor cocktail, followed by washing with cold column buffer containing 0.5% NP-40 for four times. The protein complexes bound to MBP resins were eluted with cold column buffer containing 0.36% [W/V] maltose and 1 mM DTT. For those bound to the GST bind resin were eluted with a column buffer containing 10 mM glutathione and 1 mM DTT in pH 7.5. The amounts of proteins were analyzed by western blot assay with anti-MBP or anti-His antibodies [To test direct binding between proteins, we performed an in vitro pull-down assay as previously described ,64. The tibodies .32P-labeled RNA probes along with different concentrations  of purified recombinant MBP-fusion proteins or MBP in the presence of RNA binding buffer , 2 U of RNase inhibitor, as well as 0.1 μg of tRNA in a total of 10 μl reaction volume. Two different amounts (2 and 4 pmol) of unlabeled RNAs together with 5.7 μM of either MBP-RH30 or MBP were used for template competition.The conditions for the EMSA experiments were described previously . Briefly32P-labeled single-stranded RI(-) or RII(+) RNAs were synthesized by T7-based in vitro transcription using 32P-labeled UTP. To prepare partial dsRNA duplexes, consisting of either RI(-)/DI-72 (+) or RII(+)/DI-72 (-) (see 32P -labeled RI(-) or RII(+) were annealed to 6 pmol of unlabeled DI-72(+) or DI-72 (-) in STE buffer  by slowly cooling down the samples  from 94°C to 25°C in 30 min. To test if the purified recombinant proteins could separate the partial dsRNA duplex, 1.9 and 5.7 μM purified MBP fusion proteins or MBP as a negative control were added separately to the partial dsRNA duplex in the RNA binding buffer  along with 1mM ATP, followed by incubation at 25°C for 25 min. The reaction mixtures were then treated with Proteinase K (2 μg/per reaction) at 37°C for 20 min, followed by loading onto 5% nondenaturing polyacrylamide gel with 200V for 1 h.To study if purified proteins could unwind partial dsRNA duplex, the dsRNA strand-separation assay was performed as described . Firstly (-) see , 2 pmol S1 Text(DOCX)Click here for additional data file.S1 FigThe helicase core domains are boxed. Note that the N-terminal and C-terminal domains are the least conserved between the two helicases.(DOCX)Click here for additional data file.S2 FigN. benthamiana plants transiently expressing truncated RH30 helicases were inoculated with TBSV. Top panel: Northern blot analyses of tombusvirus gRNA using a 3’ end specific probe shows the accumulation of gRNA and subgenomic RNAs in plants. Plants agroinfiltrated with the empty vector and expressing full-length RH30 were used as negative and positive controls, respectively. Bottom panel: northern blot shows 18S ribosomal RNA as a loading control. See further details in (A) (DOCX)Click here for additional data file.S3 FigΔN helicase is localized in the nucleus in the whole N. benthamiana plant and protoplasts marked with RFP-H2B. No CNV component was expressed. (B) Limited re-targeting of RH30ΔN helicase into the VROs marked by the BFP-tagged p33 replication protein in N. benthamiana plants infected with CNV. (C-D) Partition of RH30 and RH30ΔN/ΔC helicases between the cytosol and nucleus in the absence of viral components. The nucleus is marked by a histone protein (transgenic plants expressing nucleus marker RFP-H2B). Scale bars represent 10 μm. See further details in (A) Confocal microscopy images show that most of RH30(DOCX)Click here for additional data file.S1 Table(DOCX)Click here for additional data file.S2 Table(DOCX)Click here for additional data file.