Lingulodinium polyedrum using the Luciferin Binding Protein (LBP) as a specific example. LBP is a good candidate for regulation by miRNA since mRNA levels are constant over the daily cycle while protein synthesis is restricted by the circadian clock to a period of several hours at the start of the night phase. The transcriptome contains a potential DICER and an ARGONAUTE, suggesting the machinery for generating miRNAs is present. Furthermore, a probe directed against an abundant Symbiodinium miRNA cross reacts on Northern blots. However, L. polyedrum has no small RNAs detectable by ethidium bromide staining, even though higher plant miRNAs run in parallel are readily observed. Illumina sequencing of small RNAs showed that the majority of reads did not have a match in the L. polyedrum transcriptome, and those that did were almost all sense strand mRNA fragments. A direct search for 18–26 nucleotide long RNAs capable of forming duplexes with a 2 base 3’ overhang detected 53 different potential miRNAs, none of which was able to target any of the known circadian regulated genes. Lastly, a microscopy-based test to assess synthesis of the naturally fluorescent LBP in single cells showed that neither double-stranded nor antisense lbp RNA introduced into cells by microparticle bombardment prior to the time of LBP synthesis were able to reduce the amount of LBP produced. Taken together, our results indicate that circadian control of protein synthesis in L. polyedrum is not mediated by miRNAs.Dinoflagellates have been shown to express miRNA by bioinformatics and RNA blot (Northern) analyses. However, it is not yet known if miRNAs are able to alter gene expression in this class of organisms. We have assessed the possibility that miRNA may mediate circadian regulation of gene expression in the dinoflagellate  Lingulodinium polyedrum involves control over the synthesis of two proteins involved in the light emitting reaction, luciferase and luciferin binding protein (LBP) [lbp RNA during the day phase [The circadian rhythm of bioluminescence in the marine dinoflagellate in (LBP) . Both prin (LBP) . The numin (LBP) , as wellin (LBP)  and LBP in (LBP)  all corrin (LBP) , 6. The ay phase , 8. No sIn many different eukaryotes, including plants , animalsIn addition to recognition of dsRNA, RISC-mediated gene silencing can also involve small regulatory RNA termed micro-RNA (miRNA). These miRNA are transcribed from the genome and also bind RISC, although their effect lies principally in an inhibition of protein synthesis . The mecSymbiodinium microadriaticum revealed that 21 high quality miRNA candidates were present [S. kawagutii, 367 miRNAs were predicted from the genome sequence and 102 were identified by small RNA sequencing [Alexandrium tamarense [Several studies have now reported the presence of miRNAs in dinoflagellates. Small RNA sequencing of  present . In S. kquencing . This laL. polyedrum, all the bioluminescence substrate (luciferin) is bound to LBP [L. polyedrum using Northern blots and RNA sequencing. We also confirmed that both Dicer and Argonaute are found in the L. polyedrum transcriptome. Lastly, we tested if double stranded lbp RNA, introduced into cells using biolistics, would inhibit the nightly bout of LBP synthesis and thus block the synthesis of fluorescent scintillons. Our results show no evidence for inhibition of LBP synthesis by RNA interference.In d to LBP , which td to LBP . Since ld to LBP . We testLingulodinium polyedrum cultures  were obtained from the Provasoli-Guillard National Center for Culture of Marine Phytoplankton  and grown in f/2 medium [2/sec cool white fluorescent light) and 12 hours dark. ZT 0 (Zeitgeber Time 0) corresponds to the beginning of the subjective day (lights on) and ZT 12 to the onset of darkness.2 medium  at a temin vitro transcription reactions were performed with a Riboprobe in vitro Transcription Systems kit (Promega Biotech) using 10 μg linearized plasmid DNA purified using a QIAprep kit (Qiagen). To prepare transcripts from a full-length LBP cDNA, a previously described 2.3 kb LBPa cDNA in pBluescript [BamHI and transcribed using T7 polymerase to generate the antisense strand, or linearized with KpnI and transcribed using T3 polymerase to generate the sense strand. To prepare smaller 1 kb transcripts, a 1080 bp PstI fragment corresponding to the 5' end of the clone was first cloned into both SK and KS versions of pBluescript. The sense strand transcript was prepared using T7 RNA polymerase to transcribe a BamHI digested SK vector, while antisense strands were synthesized using T7 to transcribe a HindIII digested KS vector. In addition, a 45 base oligoribonucleotide corresponding to the first 45 nucleotides of the LBPa and labeled with fluorescein at the 5’ end was synthesized in an antisense orientation (Sigma Genosys). Full length GFP transcripts in the sense orientation were obtained following linearization with SpeI and transcription using T7 polymerase of a pGEM-T Easy vector (Promega) into which a PCR insert of the mGFP6 sequence from the previously described pMDC107 vector [All uescript  was eith7 vector  was liga2O and finally resuspended in 1 ml 50% glycerol. For each experiment, 2–5 μg of purified RNA was added to a mixture of 40 μl of prepared beads, 40 μl 2.5 M CaCl2 and 16 μl 0.1 M spermidine (Sigma), and vortexed for 5 min. The microparticles were washed first with 70% ethanol, centrifuged, then washed again with 100% ethanol, centrifuged and finally resuspended in 20 μl 100% ethanol. Each bombardment used all of the final resuspended microparticles. This method is essentially similar to that used to introduce purified viral RNA into dinoflagellates [Bombardment was performed using sterile M-17 tungsten particles . A bead stock was prepared from 60 mg of microparticles that were vortexed vigorously for 5 min in 1 ml 70% ethanol, left standing for 15 minutes at room temperature, centrifuged at 2000xg for 2 sec, washed twice with DEPC treated Hgellates  except f4 cells/mL was harvested by filtration on Whatman 541 paper either prior (LD12) or subsequent (LD18) to the daily bout of LBP synthesis. Care was taken to remove the filters when the cells had the appearance of a wet paste, as cells that were too dry did not survive the vacuum required for bombardment and cells that were too wet never contained microparticles after bombardment. Cells still on the filter paper were placed in a homemade particle delivery system 7–8 cm below a metal grid supporting the RNA coated beads. Air was pumped from the chamber, and when a vacuum 25 mm Hg was reached, a solenoid releasing a 5 msec pulse of 100 psi Helium was activated. After bombardment, cells were resuspended in 50 ml of fresh f/2 medium and returned to the culture room until observation the following night.To prepare cells for bombardment, 50 ml of cell culture at roughly 10Swimming cells were taken from night phase cell cultures exposed to culture room lights for thirty minutes prior to sampling and concentrated by a brief (15 sec) centrifugation at 13,000 x g. Exposure to light inhibits mechanical stimulation of bioluminescence which would otherwise reduce luciferin fluorescence . With a Solanum chacoense, and from three dinoflagellates  were electrophoresed on 5% acrylamide gels. For Northern blots, 10 μg samples of RNA from the higher plant Arabidopsis thaliana and two dinoflagellates  were transferred to a nylon membrane after electrophoresis on a 5% acrylamide gel and hybridized to an end-labeled radioactive probe prepared from the sequence 5’-acagattgcggcaaccgtgcag-3’ using the StarFire miRNA detection kit (IDT DNA). This probe corresponds to the most abundant S. kawagutii miRNA (scaffold 763_8250) previously shown to react on Northern blots with S. kawagutii RNA [For the ethidium bromide staining of small RNAs, 10 μg samples of Trizol-extracted RNA from the higher plant utii RNA . This miL. polyedrum as described [L. polyedrum rRNA sequences and S. kawagutii tRNA. Size selection of the remaining reads produced a set of reads ranging from 18–26 nucleotides. The collapse function in Galaxy was used to collapse these into a final set of 848,129 unique reads in order to determine the most abundant small RNAs. We removed from this collapsed read dataset all those containing fewer than 5 reads, leaving 133,861 collapsed reads. Among the 133,861 collapsed reads, 51,530 could be assembled to form 11,313 contigs with a 100% sequence identity over at least 14 bases. Of these, 948 had a length greater than the median length of 26 bases and contained only reads in the same direction. These 948 assemblies indicated the small RNAs contributing to them did not have the defined ends expected for bone fide miRNAs, so as with the miRPlex pipeline [Samples of total RNA were isolated from escribed , and senescribed . FastQC escribed  was usedpipeline . To lookpipeline .Acyrthosiphon pisum, Agaricus bisporus, Ajellomyces capsulatus, Aspergillus niger, Caenorhabditis elegans, Cricetus griseus, Daphnia pulex, Glycine mas, Gorilla gorilla, Homo sapiens, L. polyedrum, Medicago trunculata, Mus musculus, Nannochloropsis gaditana, Oxytricha trifallax, Pan troglodytes, Paramecium tetraurellia, Penicillium digitatum, Peromyscus maniculatus, Phaseolus vulgaris, Physcomitrella patens, Phytophtora infestans, Pongo abelii, Populus trichocarpa, Rhodnius prolixis, Ricinus communis, Strigamia maritima, Tetrahymena thermophilia and Trichoplax adhaerens.The domain structure of Dicers and Argoanutes was determined using the conserved domain function of the Blastp package . For phySymbiodinium [L. polyedrum, a species where the circadian clock regulates gene expression post-transcriptionally [S. kawagutii as a sensitive test for miRNAs revealed L. polyedrum extracts contained a cross-reacting RNA of the correct size but of lower abundance  corresponded to the nce logo , is highAs a more direct test for the presence of miRNA, we examined the 4.9 million reads for sequences able to form duplexes with 2 base 3’ overhangs. To facilitate this analysis, we first collapsed the 4.9 million reads to yield a dataset of 848,129 sequences and removed those present in less than 5 copies. We also removed sequences that could be assembled together to form longer contigs, mimicking what is done by the miRPlex pipeline . The remlbp RNA. Cells were examined at ZT18 either the same day (LD18.1) or the following day (LD18.2). Cells that contain beads are readily visible with the microscope  in single cells. LBP binds the bioluminescence substrate luciferin with high affinity , and, sicroscope , and whe, Chromalveolata (Tetrahymena), Archaeplastida (Chlamydomonas), Amoebozoa (Dictyostelium) and Opisthokonts (Neurospora). Furthermore, miRNA complementary to RNA encoding heat shock protein has a decreased abundance in heat-shocked S. kawagutii [As a more direct test for RNA-mediated interference with LBP synthesis, we developed a test for evaluating if LBP synthesis could be inhibited ht phase , 48 whicht phase  and an iht phase , and incht phase . The reght phase , thus leht phase  includinawagutii , consistlbp RNA preparations to block the normal circadian increase in scintillon numbers that occurs during the night phase of the dinoflagellate Lingulodinium as a result of LBP synthesis [We tested the ability of several different antisense or double stranded ynthesis . We predynthesis . Since mHeterocapsa circularisquama [L. polyedrum.In these studies, RNA has been introduced into dinoflagellates by microparticle bombardment, a technique known to induce gene silencing in both plants  and animrisquama  is functrisquama . This surisquama , 56, we L. polyedrum can form without LBP? Some bioluminescent dinoflagellates, such as Pyrocystis fusiformis, do contain scintillons [L. polyedrum scintillons lacking LBP. In contrast, scintillons from L. polyedrum have been purified and characterized. These contain only luciferase and LBP in roughly equimolar amounts [Is it possible that the scintillons in ntillons  yet haventillons . However amounts , and it  amounts , and eveChlamydomonas [Plasmodium and Cryptosporidium [Plasmodium and Cryptosporidium. However, the apicomplexan Toxoplasma expresses a truncated Ago that has been implicated in an effect on gene expression [S. microadriaticum [lbp gene expression in L. polyedrum.It is noteworthy that in the protists where RNA-mediated gene silencing has been demonstrated, the genome sequence has now been shown to encode Dicer-like, Argonaute-Piwi and RdRP proteins . Furtherydomonas , 60. Theporidium , 61, 62.pression , so thatriaticum  althoughS1 FigThe domain structures of animal, plant, diatom and protist Argonautes as determined by the conserved domain function in the blastp package. Conserved domains in the dinoflagellate sequences include PAZ (Piwi-Argonaute-Zwille), Argonaute N terminal (ArgoN) and PIWI PIWI-like. Animal sequences also contain an Argoanute-like (ArgoL) and a GAGE domain of unknown function.(PNG)Click here for additional data file.S2 FigFive Dicer domains from a range of animal plant fungal and protist sequences were aligned and concatenated. RAxML was used to generate the phylogeny from the concatenated alignment. Sequence names correspond to the first five letter of the genus as provided in the methods; sequences were numbered when more than one example was recovered.(TIF)Click here for additional data file.S3 FigThree different samples, taken from three different times, were processed by adapter addition and assessed by electrophoresis. The expected size of miRNAs after adapter ligation is ~150 nucleotides.(DOCX)Click here for additional data file.S4 FigThe number of sequences is reported as a function of the sequence length for the 4.9 million small RNAs sequenced.(PNG)Click here for additional data file.S5 FigUntransformed pollen has no background GFP fluorescence (arrow).(PNG)Click here for additional data file.S1 Table(PDF)Click here for additional data file.S2 Table(PDF)Click here for additional data file.


Caenorhabditis elegans transcriptome and its modifications and use them as reference standards to characterise the RNA modification profile in this multicellular organism through an untargeted liquid‐chromatography tandem high‐resolution mass spectrometry (LC‐HRMS) approach. We furthermore show that several of these RNA modifications have a dynamic response to environmental stress and that, in particular, changes in the tRNA wobble base modification 5‐methoxycarbonylmethyl‐2‐thiouridine (mcm5s2U) lead to codon‐biased gene‐expression changes in starved animals.More than a hundred distinct modified nucleosides have been identified in RNA, but little is known about their distribution across different organisms, their dynamic nature and their response to cellular and environmental stress. Mass‐spectrometry‐based methods have been at the forefront of identifying and quantifying modified nucleosides. However, they often require synthetic reference standards, which do not exist in the case of many modified nucleosides, and this therefore impedes their analysis. Here we use a metabolic labelling approach to achieve rapid generation of bio‐isotopologues of the complete  Canonical nucleobases, especially those of ribonucleic acids, are naturally subject to diverse modification. The number of identified modified ribonucleosides has grown to over a hundred, and more than half of them can be found in eukaryotic RNA.5C), pseudouridine (ψ), N1‐methyladenosine (m1A) and N6‐methyladenosine (m6A).The recent development of sequencing‐based detection methods has facilitated the identification of modifications in mRNAs. Examples include 5‐methylcytidine  lead to codon‐biased gene‐expression changes in starved animals. We based our method on the powerful software tool MetExtract, originally developed for the automated extraction of metabolite‐derived LC‐MS signals from LC‐HRMS full‐scan data obtained from isotopically labelled biological samples.C. elegans. d‐[13C6]Glucose had previously been used to isotopically label the bacterial transcriptome as it feeds into the pentose phosphate pathway .C. elegans in the presence of isotopically labelled bacteria, serving as food, to transfer the isotopes from the Escherichia coli transcriptome to the C. elegans transcriptome and to generate bio‐isotopologues of every modified ribonucleoside present in the animal. We first grew HT115 E. coli in the presence of d‐[13C6]glucose by adaption of previously reported methods.13C incorporation into amino acids involved in de novo nucleotide synthesis and thus to achieve more uniform 13C6‐labelling rather than 13Cn‐labelling .In this approach, we use metabolic labelling to generate bio‐isotopologues of the C. elegans in the presence of the labelled bacteria for three generations, after which the nematodes were harvested for total RNA isolation . Having established the 13C‐labelling of C. elegans RNA, we aimed to determine its most abundant RNA modifications. We thus fractionated total RNA into small (<200 nt) and large (>200 nt) RNAs  to increase the sensitivity for the less abundant—primarily tRNA—modifications in the small‐RNA fraction.We then cultured n Figure . The degm/z range  was then measured by HPLC‐HRMS in the 250–500 13C6 isotopologues.  For this reason some residues were found more than once: the mass of methylcytidine, for instance, was found three times, presumably m5C, 3‐methylcytidine (m3C) and 2′‐O‐methylcytidine (Cm). Altogether, we identified 21 and 26 modifications in the C. elegans large‐ and small‐RNA fractions, respectively uridine (acp3U) and its corresponding 13C6 isotopologue, a modification that has not been previously described in eukaryotic RNA, thus demonstrating the potential of our approach for the discovery of modified ribonucleosides in a completely untargeted fashion.Thus, at this stage we had identified RNA modifications by their accurate masses and those of their corresponding, co‐eluting, C. elegans under physiological stress to improve understanding of the relationship between modified RNAs and stress response pathways. Heat stress and starvation are two stress conditions that lead to large‐scale gene‐expression changes in animals.C. elegans larvae to heat shock by shifting adult animals from 20 to 37 °C for 4 h or we starved the animals by removing their food source for 4 h at 20 °C. Within 4 h either of heat stress or of starvation, adult C. elegans animals showed gene‐expression changes that were specific to each stress condition: upon heat stress several heat‐shock factors were strongly upregulated , whereas geneexpression changes upon starvation strongly overlapped with previous starvation data on C. elegans .Next, we aimed to quantify the RNA modification landscape of C. elegans before and after stress induction. This was achieved by spiking in digested 13C6‐labelled RNA at 5 %, v/v into unlabelled digested RNA obtained from stressed (heat‐shocked or starved) or control animals and subjecting the mixture to targeted LC‐MS/HRMS analysis. Where possible, standards from commercial and synthetic sources were used to identify the presence of most residues unequivocally , m5C, Cm, Gm, N2,N2‐dimethylguanosine (m22G), 7‐methylguanosine (m7G), I, Um, 5‐methyluridine (m5U), mcm5s2U); moreover, with the added selectivity of tandem mass spectrometry (MS/HRMS), other residues for which standards are not available could also be assigned by taking the co‐elution of their fragments masses into account . We were unable to resolve putative m1G and m2G. However, because these residues do not change we did not proceed further in structurally identifying them and refer to these residues as mG (base‐methylated guanosine). The peak areas of each modified nucleoside and its corresponding 13C6 bio‐isotopologues, present in the spike‐in, were then determined in each sample, and their ratio was calculated  to obtain a normalised peak area for every modification under three different sets of conditions  RNA modification changes in heat‐shocked animals: the N2,N2,7‐trimethylguanosine  residue showed a strong reduction in the large‐RNA fraction, and mcm5s2U levels showed a modest decrease in the small‐RNA fraction. On the other hand, starvation induced numerous RNA modification changes in both the large‐ and the small‐RNA populations. Interestingly, in the large‐RNA fraction in particular, we observed that several base methylations increased upon starvation; these included m5C, m1A, m7G, m5U, m2,2,7G and m22G. Observable RNA modification changes upon starvation in the small‐RNA fraction were constrained to two known tRNA wobble base modifications: ac4C and mcm5s2U levels both decreased in starved animals. We validated our results by absolute quantification of two of the RNA modifications by a method previously reported by us5C levels in the large‐RNA fraction were indeed found to be significantly increased upon starvation , whereas m6A levels in the large‐RNA fraction showed no change between control and starved animals . Both these absolute quantitations are in line with the relative quantification method based on bio‐isotopologues presented here. Overall, this allowed us to determine the relative abundance of multiple RNA modifications in whole animals upon stress induction.Next, we compared the normalised peak areas of stressexposed nematodes with those of control animals for every modification Figure  A, B. We5C and m5U in the large‐RNA fraction  and mcm5s2U in the small‐RNA fraction reversing significantly. As shown in detail for mcm5s2U in Figure p value <0.01) its levels no longer differed from those in the control population. Absolute quantification of m5C and m6A in the large‐RNA fractions of rescued animals and control animals  were again in line with our relative quantification measurements: that is, we measured no difference between control and rescued populations for these modifications. Our results show that several RNA modifications not only respond to stress conditions, but are also dynamically regulated between stress and normal growth conditions of C. elegans. We were particularly interested in the tRNA wobble base modification mcm5s2U, which showed levels that were significantly reduced in starved animals but were then restored to wild‐type levels upon reintroduction of food. mcm5s2U is found in tRNA‐LysUUU, tRNA‐GluUUC and tRNA‐GlnUUG at position U34 of the anticodon loop and it is required for cell viability and animal development.5s2U has been linked to inefficient translation, ribosome stalling and protein misfolding in cases of genes that are enriched for codons AAA, GAA and CAA.To gain a full picture of the dynamic nature of RNA modifications, we tried to determine whether or not the stress‐induced changes in RNA modifications are reversible. To this end we rescued starved animals by reintroducing food and compared the changes in RNA modifications to those in age‐matched control animals. All RNA modifications that were significantly altered upon starvation showed a general trend of reversal upon starvation rescue, with, for example, m5s2U levels leads to codon‐specific changes in gene expression, we analysed the codon enrichment of AAA, GAA and CAA in comparison with all other codons, among genes that show differential expression upon starvation . Surprisingly, the codons AAA and GAA show significant enrichment in differentially expressed genes after starvation  and MgSO4 (1 mm). Starter cultures were used for growing fresh E. coli HT115 cultures in M9 minimal medium to an OD600 of 0.8–1.0. Bacterial cultures were pelleted for RNA isolation or C. elegans culture. Resuspended bacteria were seeded to NGM‐N agarose plates, and the plates were left to dry overnight. Ten larval stage 1 animals were placed on seeded NGM‐N agarose plates with the labelled bacteria and left to grow for two generations. Adult F1‐generation animals were bleached to obtain a synchronous population of F2‐generation animals. F2 animals were placed on freshly seeded NGM‐N plates with labelled bacteria and grown to adult stage. Adult animals were washed off the plates and pelleted before DNA and RNA isolation. Synchronised populations of L1 animals were grown until the young adult stage. For heat stress, animals were transferred to 37 °C incubators for 4 h with food. After 4 h, animals were washed off the plates, cleaned from bacteria by washing in M9 buffer (3×) and stored in TRIsure (Bioline) reagent for subsequent RNA isolation. For starvation experiments, young adult stage animals were washed off the plates using M9 buffer and plated either on food plates for control or on empty plates for starvation. Animals were left at 20 °C for 4 h. After 4 h, animals were washed off the plates, cleaned by washing in M9 buffer (3×) and stored in TRIsure reagent for RNA isolation. For rescue experiments, control and starved animals were plated on plates with food and left for 8 h at 20 °C.Starter cultures of Zymo Research) according to the manufacturer's instructions.Isolated total RNA (see the Supporting Information) was fractionated into “small” and “large” RNAs with the aid of a Quick‐RNA MiniPrep kit (m/m based on RNA amounts). For the <200 nt fraction the unlabelled RNA (1.5 μg) was added to the 13C SIL small RNA (3 μg). For the >200 nt fractions, 3 and 6 μg, respectively, were used. LC‐HRMS was performed with a Thermo Ultimate 3000 UHPLC system equipped with a Waters HSS‐T3 column and coupled to a Thermo Qexactive hybrid mass spectrometer. LC conditions were as follows: H2O/MeCN solvent system ; HRMS was performed in Full MS‐SIM mode, resolution 35 000, scan range 250–500 m/z.Enzymatic digests of size‐fractionated RNA were mixed in a 1:2 ratio  was added as the internal reference standard. With the same experimental setup, LC‐MS/HRMS was then performed with the machine operated in MRM mode; resolution 35 000 and inclusions as listed in Table S3.Using the absolute concentration of rC as a reference for the amount of RNA in each sample (Table S2) we adjusted the samples' RNA concentrations. Complete experimental procedures are provided in the Supporting Information.The authors declare no conflict of interest.As a service to our authors and readers, this journal provides supporting information supplied by the authors. Such materials are peer reviewed and may be re‐organized for online delivery, but are not copy‐edited or typeset. Technical support issues arising from supporting information (other than missing files) should be addressed to the authors.SupplementaryClick here for additional data file.


Influenza viruses have a segmented viral RNA (vRNA) genome, which is replicated by the viral RNA-dependent RNA polymerase (RNAP). Replication initiates on the vRNA 3′ terminus, producing a complementary RNA (cRNA) intermediate, which serves as a template for the synthesis of new vRNA. RNAP structures show the 3′ terminus of the vRNA template in a pre-initiation state, bound on the surface of the RNAP rather than in the active site; no information is available on 3′ cRNA binding. Here, we have used single-molecule Förster resonance energy transfer (smFRET) to probe the viral RNA conformations that occur during RNAP binding and initial replication. We show that even in the absence of nucleotides, the RNAP-bound 3′ termini of both vRNA and cRNA exist in two conformations, corresponding to the pre-initiation state and an initiation conformation in which the 3′ terminus of the viral RNA is in the RNAP active site. Nucleotide addition stabilises the 3′ vRNA in the active site and results in unwinding of the duplexed region of the promoter. Our data provide insights into the dynamic motions of RNA that occur during initial influenza replication and has implications for our understanding of the replication mechanisms of similar pathogenic viruses. Negative-sense RNA viruses include a number of important human and animal pathogens; examples of disease caused by these viruses include measles, rabies, Ebola and influenza. Influenza A and B viruses are responsible for seasonal respiratory infections and influenza A viruses also cause occasional severe pandemics in humans resulting in substantial morbidity and mortality worldwide. Both type A and B viruses have eight viral RNA (vRNA) segments, while influenza C viruses have seven segments. The conserved 13 nucleotides at the 5′ end and 12 nucleotides at the 3′ of each vRNA segment are partially complementary and form a partly double-stranded promoter created by base-pairing between the two ends  assay to measure distances between fluorescent dyes located on the viral RNA promoter upon RNAP binding ; this asRNA corresponding to the 3′ and 5′ conserved ends of the neuraminidase vRNA or cRNA gene segment were custom synthesized by IBA  and were labelled with the fluorophores Cy3 or Cy3B  and ATTO647N (Sigma) as previously described , before Recombinant A/NorthernTerritories/60/68 (H3N2) viral RNAP with a protein-A tag on the PB2 subunit was expressed using the Multibac system  in Sf9 i2, 100 μg/μl BSA, 1 mM DTT and 5% glycerol) before being diluted into buffer B (50 mM Tris–HCl (pH8), 100 mM KCl, 10 mM MgCl2, 100 μg/μl BSA, 1 mM DTT and 5% glycerol) to a final RNA concentration of 100 pM for confocal analysis. For RNAP labelling, 2–3 nM of Cy3- or ATTO647N-labelled trisNTA was incubated with 100 nM RNAP and 1 nM fluorescently labelled RNA in buffer A for 15 min at 28°C, before being diluted into buffer B for confocal analysis. For nucleotide addition, 500 μM ApG was incubated with 100 nM RNAP and 1 nM fluorescently labelled RNA in buffer A for 15 min at 28°C, before being diluted into buffer B for confocal analysis. The average excitation intensities were 250 μW at 532 nm and 60 μW at 635 nm. Custom-written LabVIEW software was used to register and evaluate the detected signal.A custom-built confocal microscope was used for single-molecule FRET experiments as previously described –17 and tE*, were calculated for each fluorescent burst above a certain threshold, yielding a two-dimensional histogram  (A model which incorporated the influenza A polymerase  and resictively) . The ATTctively) ,23. DistTranscription was carried out using double-stranded RNA templates in the presence of ApG as a primer, as previously described . Sequencin vitro replication assay . Single-molecue FRET was measured on complexes diffusing in solution, and apparent FRET efficiency values (E*) reporting on the donor-acceptor distance between the fluorophores on the RNA were depicted as merged histograms from three independent experiments and fitted with Gaussian functions to determine the mean FRET efficiency of the distributions   with short, partially complementary RNAs corresponding to the conserved 5′ and 3′ termini of the vRNA, fluorescently labelled with a donor dye at position 18 on the 5′ strand and an acceptor dye at position 1 on the 3′ strand Figure  10,13).,13.10,13ibutions ,25.E* values of 0.48 and 0.79 , can be explained by the increased distance of the dye at 3U1 to the dye at 5U18 as the template RNA moves through the active site, while the shift of the high-FRET population to a lower mean E* value (from 0.79 to 0.72) may reflect movement of the 5U18 dye during initial replication. ApG addition resulted in an increase in the low-FRET population  and a decrease in the high-FRET population (from 42% to 22%) compared to when no ApG was present . In order to confirm that the changes in FRET efficiency that we observed were due to changes in distance, rather than restricted orientation of the dyes, we used an RNA construct where the dyes were swapped so that a Cy3 dye was placed at position 3U1 and an ATTO647N dye was placed at position 5U18. The accessible volume modelling of the dyes showed that the Cy3 dye in the RNAP active site was constrained in a very different way to the ATTO647N dye , however we observed a similar result to that shown in Figure E* values of 0.58 and 0.81)  Figure , middle.) Figure , bottom.E* ∼ 0.85) , suggesting an increase in the distance between the two dyes  Figure , middle,s Figure , bottom,tris-nitrilotriacetic acid (trisNTA) (10 tag placed at amino acid 200 within a ß-ribbon of the PB1 subunit of the RNAP (PB1 his200). Labelling was carried out in situ, resulting in a large background from unbound dye-conjugated trisNTA , however a clear FRET signal could be observed. FPS modelling of the dye positions provided an estimation of the number of FRET populations to expect from each labelling scheme; however, accurate donor-acceptor distance calculations were not possible due to the additional histidine residues not being present in the structure  –30 was be Figure . The Hisn Figure  or in viy Figure . When wed values of 44 nM, >1 μM and 13 nM, respectively . Similar experiments carried out previously with vRNA showed that the Kd of the RNAP for the 5′ vRNA terminus, the 3′ vRNA terminus, and the vRNA promoter was 2.2 nM, >1 μM and 0.4 nM, respectively . The promoter alone gave a single high-FRET population with a mean The presence of a bimodal FRET distribution using the 5U5 and 3U1 labelling strategy showed that the cRNA 3′ end adopts two different conformations when bound by RNAP, possibly corresponding to pre-initiation and initiation states similar to that of the vRNA promoter. ApG addition resulted in a decrease in the high-FRET population Figure . An RNA E* value of 0.56 for the RNA only, and a single FRET distribution centred at an E* of 0.78 when RNAP was added  upon ApG addition, consistent with the dyes moving further apart, suggesting that dinucleotide addition destabilised the distal region . The 5′ and 3′ termini in this region must separate during processive elongation, when the 3′ end of the template vRNA translocates through the active site while being copied into nascent cRNA. However, it is unclear at which point the two termini separate. To detect changes in the distal region, we first placed a dye pair at position 3 on the 5′ vRNA and position 17 on the 3′ vRNA. This resulted in a single FRET distribution centred at an n Figure , bottom.n Figure . MeasureE* ∼ 0.83), rather than the fluorescence quenching that was expected  , consistent with the dyes moving further apart, indicating that dinucleotide addition can act to destabilise this early distal region  Figure , middle.n Figure , bottom.E* ∼ 0.85  and was hence sufficient to destabilise this region , forming ribonucleoprotein (RNP) complexes. The pre-initiation state may represent the vRNA conformation found within vRNP complexes packaged into progeny virions, which are not actively replicating. The addition of ApG, mimicking an initial replication product, appears to bias the RNAP complex towards the initiation state, supporting the idea that this state represents an active form. The binding of the 3′ vRNA on the RNAP surface could also represent a ‘waiting’ conformation that the 3′ end adopts after it has been copied and exits from the active site, ensuring that it is located nearby to allow subsequent rounds of transcription to occur with a minimal delay after copying of the gene segment . The 3′ We have also shown that the 3′ terminus of the cRNA promoter is likely to form a pre-initiation state. Negative-stain EM has revealed that cRNPs are structurally similar to vRNPs , and seqOur results support a model in which the RNAP-bound 3′ end of the template vRNA adopts two conformations, consistent with predictions of pre-initiation and initiation states, and whereby translocation of the 3′ vRNA through the active site during initial replication results in destabilisation and processive unwinding of the double-stranded duplex Figure . This moSupplementary Data are available at NAR Online.


Nematostella vectensis and Hydra magnipapillata) and one poriferan (Amphimedon queenslandica). Our analysis revealed that nonbilaterian miRNAs frequently undergo modifications like the bilaterian counterparts: the majority are expressed as different length isoforms and frequent modifications of the 3′ end by mono U or mono A tailing are observed. Moreover, as the factors regulating miRNA modifications are largely uncharacterized in nonbilaterian animal phyla, in present study, we investigated the evolution of 3′ terminal uridylyl transferases (TUTases) that are known to involved in miRNA 3′ nontemplated modifications in Bilateria. Phylogenetic analysis on TUTases showed that TUTase1 and TUTase6 are a result of duplication in bilaterians and that TUTase7 and TUTase4 are the result of a vertebrate-specific duplication. We also find an unexpected number of Drosophila-specific gene duplications and domain losses in most of the investigated gene families. Overall, our findings shed new light on the evolutionary history of TUTases in Metazoa, as they reveal that this core set of enzymes already existed in the last common ancestor of all animals and was probably involved in modifying small RNAs in a similar fashion to its present activity in bilaterians.In bilaterian animals the 3′ ends of microRNAs (miRNAs) are frequently modified by tailing and trimming. These modifications affect miRNA-mediated gene regulation by modulating miRNA stability. Here, we analyzed data from three nonbilaterian animals: two cnidarians ( MicroRNAs (miRNAs) are produced through a series of complex enzymatic processes prior to taking their mature form. In general, primary miRNA (pri-miRNAs) processing is initiated by cleaving the pri-miRNAs by an RNAse III enzyme called Drosha and released into cytoplasm as premiRNA . In the Schizosaccharomyces pombe the TUTase-related Cid1 protein family regulates mRNA turnover by targeting specific mRNAs and promoting their decapping  is involved in uridylation of histone mRNAs and promotes their decay  and Cnidaria   , it is pllyfish) .Nematostella vectensis and Hydra magnipapillata) and Porifera (Amphimedon queenslandica). Additionally, we analyzed the phylogeny of TUTases family throughout Metazoa and their holozoan relatives to unravel the evolutionary history of these RNA modifying enzymes.To assay whether miRNA modifications such as nontemplated addition and trimming occur in nonbilaterian animals, we performed an in-depth analysis of small RNA sequence data from Cnidaria  on all available metazoan species (under the term Metazoa) as well as on Choanoflagellata and Capsaspora using Homo sapiens protein sequences as queries. In case we found no matches we carried out TBLASTN searches with the same query sequences, using the Transcriptome Shotgun Assembly (TSA) database of NCBI and restricted it to Cnidaria and Porifera. In addition, for nonbilaterian species we carried out species-specific searches through Compagen     , Joint G4, 2017)  and the 4, 2017)  databaseNematostella vectensis polyps were grown in 16 ‰ artificial sea water at 18 °C in the dark and fed three times a week with freshly hatched Artemia salina nauplii. Induction of spawning was performed as previously described  using Tri-Reagent (Sigma-Aldrich) following the manufacturer’s instructions. Three biologically independent animal pools were used for each developmental stage. From each stage 20 µg of total RNA was used and the RNA integrity was analyzed with a Bioanalyzer . The small RNA size selection was performed using 15% denaturing urea polyacrylamide gel, RNA elution from gel was performed overnight and precipitated using ethanol. Small RNA libraries were constructed as described in Phillip Zamore Lab Illumina TruSeq Small RNA Cloning Protocol April, 2014 . Small RNA libraries were sequenced in the Center for Genomic Technologies of the Hebrew University on the NextSeq 500 platform (Illumina) with 50 nt read length. The raw data have been deposited at NCBI GEO Submission (GSE94526)   online.Nematostella genome from the NCBI database was used as a reference, and the mature miRNA and miRNA precursor were retrieved from miRBase Release 21 (Hydra magnipapillata and Amphimedon queenslandica were retrieved from data deposited at NCBI Sequence Read Archive (SRA050926) .The sequencing data were preprocessed to remove the adapters using Cutadapt  and sequlease 21 . The smaA050926)  and (SRPA050926) , respectNematostella vectensis, Amphimedon queenslandica, and Hydra magnipapillata. In Nematostella we analyzed small novel RNA sequencing data we generated from distinct RNA preparations of late planula, primary polyp and adult male. The small RNA sequencing data of Hydra and Amphimedon were retrieved from previously published data  (Hydra magnipapillata (SRA accession number SRP037736)   and HydrP037736) . We founP037736)  online. P037736)  online, Drosophila and humans we have noticed changes in protein domain assembly. Thus, we decided to perform classification of all TUTases in human, zebrafish and Drosophila through phylogenetic analysis to understand the relationship of TUTases among these three species.From an initial comparative analysis on TUTases in both Drosophila into GLD2 and Wispy. In addition, we noted that TUTases6 duplication in the Drosophila lineage resulted in Tailor and Monkey King . This is probably the result of a loss of TUTase2 that is specific to demosponges as we were able to find TUTase2 homologs in the Calcarea and Homoscleromorpha sponge classes (Drosophila into TUTase2 (GLD2) and Wispy  and Wispy from most major phyla of animals and other holozoans. Initially we did not find exact matches for TUTase2 in Choanoflagellata or  classes . Howevernd Wispy , in ordend Wispy . Furthernd Wispy . In our nd Wispy .Capsaspora. As we observed that TUTase4 is absent in Diptera we continued searching in other Classes of Hexapoda. Unlike Diptera, other subphyla of arthropods, including the other groups of Neoptera (winged insects) do carry TUTase4 in their genome. This strongly suggests a dipteran-specific loss of TUTase4  polymerases (PAPs) associated domain  and one poriferan (Amphimedon queenslandica). Our analysis revealed that in all three species miRNAs frequently undergo modifications like in bilaterians: the majority of miRNAs are expressed as different length isoforms and frequent modifications of the 3′ end by mono U or mono A tailing are observed  took over the roles of TUTase4 that was lost.Based on our analyses of miRNA 3′ nontemplated modifications in nonbilaterian animals, we observed that mono-uridylation is the most frequent modification C and 2C.psaspora . Anotherpsaspora . Despitee for it . Taking In contrast to TUTase4, TUTase7 is found only in subgroups of vertebrates  and 7 anModification of certain miRNAs at their 3′ end by addition of mono (A) has been noted in several organisms and TUTase2 was found to responsible for this modification . FurtherDrosophila (Drosophila and its possible overlap with Wispy is currently unknown.In the case of Drosophilidae, we found unprecedented number of modifications in TUTase family of proteins  and 7, tosophila . Additioosophila . GLD2 fuDrosophila could originate from a TUTase6 duplication. As mentioned earlier, the Drosophila Tailor is directly involved in catalysis of the miRNA 3′ uridylation. Deletion of Tailor in Drosophila S2 cells increases the levels of mirtrons, a class of noncanonical splicing-mediated miRNAs. This demonstrates that Tailor has a role in negatively regulating mirtrons accumulation (Drosophila, while being an extremely important model organism for the functional study of RNA metabolism in animals, probably possess a highly derived RNA tailing mechanism.Beside the TUTase2 duplication in Drosophilidae, we found that TUTase6 is also duplicated into Tailor and Monkey King specifically in this lineage . TUTase6mulation . To the mulation . These fmulation . The numIn addition to TUTase6 duplication in Drosophilidae, we noted that TUTase1 probably has been derived from a gene duplication of TUTase6 in Urbilateria . StudiesCapsaspora and Metazoa. Further, other major transitions in the TUTase family that our study reveals include the emergence of TUTase2 and TUTase4 at the base of Metazoa, the loss TUTase2 in demosponges, and the emergence of TUTase1 is bilaterians (In overall, our study reveals that three members of the TUTase family, TUTases 3, 5, and 6, were most likely already present in the last common ancestor of aterians .We found that miRNAs in nonbilaterian animals often undergo modifications by tailing like in bilaterians, suggesting that this RNA stability control system was already present in the last common ancestor of all animals. However, variation in the components of this system exists in some animal lineages, with Drosophilidae being the most striking example for substantial divergence. By phylogenetic studies on TUTases, we revealed several important events in the evolution of this protein family in the animal kingdom . Lastly,Genome Biology and Evolution online.Supplementary DataClick here for additional data file.


Chara vulgaris, a multicellular alga found within the Streptophyta clade and one of the closest extant algal relatives of land plants. In this study, the organelle transcriptomes of C. vulgaris and Chlamydomonas reinhardtii were deep sequenced for a comprehensive assessment of RNA editing. Initial analyses based solely on sequence comparisons suggested potential edit sites in both species, but subsequent high-resolution melt analysis, RNase H-dependent PCR (rhPCR), and Sanger sequencing of DNA and complementary DNAs (cDNAs) from each of the putative edit sites revealed them to be either single-nucleotide polymorphisms (SNPs) or spurious deep sequencing results. The lack of RNA editing in these two lineages is consistent with the current hypothesis that RNA editing evolved after embryophytes split from its ancestral algal lineage.Nearly all land plants post-transcriptionally modify specific nucleotides within RNAs, a process known as RNA editing. This adaptation allows the correction of deleterious mutations within the asexually reproducing and presumably non-recombinant chloroplast and mitochondrial genomes. There are no reports of RNA editing in any of the green algae so this phenomenon is presumed to have originated in embryophytes either after the invasion of land or in the now extinct algal ancestor of all land plants. This was challenged when a recent in silico screen for RNA edit sites based on genomic sequence homology predicted edit sites in the green alga  Triticum aestivum) and evening primrose (Oenothera berteriana), this phenomenon appears to repair deleterious mutations and preserve protein function in organelles that are limited to clonal reproduction and prone to a relatively high rate of mutation [Plant organelle RNA editing is a post-transcriptional change in the nucleotide composition of an RNA and in most cases involves a change in a pyrimidine by the addition or removal of an amino group to convert a uracil to or from a cytosine . First dmutation ,3,4. An Arabidopsis, tobacco, and O. berteriana mitochondria [The majority of editing occurs within coding regions  and theschondria ,8,9,10. Marchantia polymorpha, which appears to have lost the ability [Ceratium horridum, Heterocapsa triquetra, Lingulodinium polyedrum, and Symbiodinium minutum [Extensive organelle RNA editing occurs among almost all embryophytes  with the ability ,13,14. T ability ,16. RNA  ability . Dinofla ability  whose pl ability ,20,21. E minutum ,22,23,24Chara vulgaris [In addition to the sequence-derived evidence for RNA editing, in silico methods have been developed that use evolutionarily conserved mRNA edit sites among embryophytes to predict sites in newly produced genomic sequences ,28,29,30vulgaris .Chara, plus the potential for easily overlooked non-coding region edits led us to undergo a complete survey of organelle transcriptomes of two green algae, C. vulgaris and Chlamydomonas reinhardtii, chosen to represent the Streptophyta and Chlorophyta algal lineages [The lack of published evidence for the absence of RNA editing in green algae, the presence of editing in two photosynthetic lineages, the in silico prediction of editing in lineages . InitialC. vulgaris was dredged from a pond on the UVa-Wise campus , March 2015, and a culture was initiated in a 20 liter aquarium along with mud and water from the same pond. The culture was maintained in the UVa-Wise greenhouse with natural light. The tank water was replenished periodically with pond and/or distilled water.C. reinhardtii, strain cc503, was obtained from the Chlamydomonas stock center , January 2015, and maintained on Tris–acetate–phosphate (TAP) medium with Hutner’s trace elements [−2 s−1 using a combination of LED and plant growth lights.elements . Liquid C. vulgaris thalli were cut from the greenhouse grown culture, washed multiple times in distilled water to remove contaminating algae and bacteria, and excess water was blotted from them with paper towels. Thalli were immediately placed in liquid nitrogen and ground into a powder using a mortar and pestle. DNA or RNA was extracted using Qiagen’s DNeasy and RNeasy kits , according to the manufacturer’s instructions, and RNA extraction included the optional DNase treatment. Nucleic acid yields were determined using a Nanodrop Lite  apparatus. C. reinhardtii cells were pelleted in a clinical centrifuge at 2000× g for 5 min, the growth medium was decanted, and the pellet was immediately resuspended in extraction buffer.Chara samples used for deep sequencing were collected in April and July 2015. Illumina sequencing of the Chara genome and April transcriptome sample libraries was completed by the University of Virginia’s sequencing core . Illumina sequencing of the July Chara and C. reinhardtii total transcriptome libraries was completed by Genewiz .Chara using GENEious software v. 8 [Plastome and chondriome sequences were assembled for are v. 8  and GenBare v. 8 ,36.Nicotiana tabacum [Chara and C. reinhardtii with at least 10% of the transcripts having a discrepancy from the genomic reference sequence were then manually inspected to determine if any differences were caused by phenomena other than RNA editing.Each transcriptome was aligned to chondriome and plastome genomic sequences and putative edit sites were initially determined using GENEious’ “Find Variations/SNPs” function set at a minimum variant frequency of 0.1. This very low editing frequency threshold was chosen to minimize the possibility of overlooking edits due to a low editing frequency. This was based on the deep transcriptome alignment and analysis of edit sites found in the whole chondriome of  tabacum  where a High resolution melt (HRM) has been used previously to define edit sites  and usesChara was removed from the greenhouse culture, rinsed with distilled water, and placed in 100 ml of liquid medium containing ¼× TAP salts and ¼× Hutner’s trace elements [Chara was incubated in triplicate in each temperature + pH regimen for 3 days in a temperature controlled growth chamber on a 16 h light/8 h dark cycle and 200 μE m−2 s−1 of light.One gram of elements  in 250 mThalli were harvested from the growth medium, blotted dry, immediately placed in liquid nitrogen, and ground into a powder using a mortar and pestle. DNA and RNA were extracted separately using Qiagen’s DNeasy and RNeasy kits, according to the manufacturer’s instructions, and RNA extraction included the optional DNase treatment. Nucleic acid yields were determined using a Nanodrop Lite apparatus. Complementary DNA (cDNA) was produced from DNase treated RNA using Bio-Rad’s iScript cDNA synthesis kit .psbI  and psbA (negative control) using the melt analysis protocol of Bio-Rad’s CFX96 Real-Time PCR system programmed to heat from 65 °C to 95 °C at 0.1 °C increments. Data were collected from gDNA and RNA extracted from three Chara replicates and three technical replicates for a total of n = 9 melt temperature differences for each environmental condition. Melt temperature differences were calculated by subtracting the lower melt temperature from the higher one. Two-way analysis of variance (ANOVA) of the melt temperature differences was completed using SigmaStat v. 4.0 .Amplicons were produced as paired PCR reactions, one genomic DNA (gDNA) for each cDNA, harvested from the nine pH + temperature conditions using Bio-Rad’s Precision Melt Supermix with 25 ng of gDNA or cDNA template and 10 pmol of each primer. Primers were designed with the aid of Primer3 : psbI-F Chara’s psbI gene according to the criteria and technical help provided by Integrated DNA Technologies , psbI-rh-F 5′-TTACGTCCTGGATTACGTCuGGGAA-x psbI-rh-R 5′-GTATATACAGTTGTAATATTgTTTAA-x (rhPCR is a test that can distinguish single nucleotide polymorphisms (SNPs) between two otherwise identical template DNAs using qPCR . rhPCR pgTTTAA-x . The -x Chara psbI reactions included 25 nM DNA or cDNA template, 10 pmol of each primer, PerfeCTa SYBR green supermix , 0.01% Triton X-100, and 8 nU RNase H2 (Integrated DNA Technologies). qPCR was performed in Bio-Rad’s cfx96 Real-Time PCR system with the program . A rhPCR N. tabacum positive control was designed based on an edit site in the mitochondrial cytochrome oxidase b (cob) gene  using the primers 5′-TTACTATTGGACAAATTTCTuCTTA-x and 5′-TCTGGGACGAGTTGGAAGAG and amplified using the same strategy described above  chondriome and 1,523,432 reads aligned to the 184,933 bp plastome. Compared to the GenBank archived sequences, the assembled plastome differed by 21 SNPs—nineteen in intergenic regions and introns, one synonymous change in rps3, and one Gly115Val missense in rpl22  gene exons. The newly generated plastome and chondriome sequences were used as templates for transcriptome alignment and analysis.Since SNPs and RNA editing would be indistinguishable in our assay, it was necessary to sequence the plastome and chondriome of the in rpl22 . The choC. vulgaris thalli in April (20 °C) and July 2015 (30 °C) and sequenced as separate replicates with yields of 43,741,464 paired-end reads for the April extraction and 187,760,146 for July.To find potential edit sites, total RNA was extracted from In the mitochondrial transcriptome, 67,949 reads from the April replicate and 12,418,978 reads from the July replicate aligned to the chondriome. One hundred and seventy nucleotide variants were found and investigated. All of them occurred outside of protein coding regions and upon manual inspection, each was found to be the result of misaligned sequences or due to introns directly adjacent to intron/exon junctions. None of them were likely candidates for RNA editing.psbI coding region from the April replicate. These variants did not appear in the July library . There were no significant changes noted in psbI or psbA when the pH of the growth medium differed and no significant interaction between temperature and pH.HRM analysis was used to measure nucleotide changes between eratures A, but twpsbI DNA and mRNA [cob gene  and mitochondrial (EU306622.1) sequences for this strain. For the chloroplast, 32,533,027 RNA derived reads aligned to the 184,933 bp plastome and for the mitochondria, 12,418,978 reads aligned to the 67,737 bp chondriome.To screen for potential editing in Chara, one putative edit site was detected in the mitochondrial nucleotide position, 1182, within the cytochrome oxidase b coding region  and the possibility that C. vulgaris plastome predicted there may be edit sites within coding regions. In this study, the transcriptomes of two green algae were deep sequenced and thoroughly screened for organellar editing. Some putative edit sites were initially detected but subsequent experiments determined these were either spuriously detected, or a single nucleotide polymorphic difference between the clonal isolate that was sequenced versus the archived genome sequence. Based on these results we can say with confidence that there is no organellar RNA editing in these two species.It is assumed that organellar RNA editing does not occur in algae, but there are no published accounts offering empirical evidence of its absence and a recently published in silico screen of the


Lettuce chlorosis virus (LCV), a member in the genus Crinivirus (family Closteroviridae). The YSS is the first in this family of viruses to be determined using Selective 2′-Hydroxyl Acylation Analyzed by Primer Extension (SHAPE). Using luciferase constructs/replicons, in vivo and in vitro assays showed that the 5′ and YSS-containing 3′ terminal regions of LCV RNA1 supported translation activity. In contrast, similar regions from LCV RNA2, including those upstream of the YSS, did not. LCV RNA2 mutants with nucleotide deletions or replacements that affected the YSS were replication deficient. In addition, the YSS of LCV RNA1 and RNA2 were interchangeable without affecting viral RNA synthesis. Translation and significant replication were observed for specific LCV RNA2 replicons only in the presence of LCV RNA1, but both processes were impaired when the YSS and/or its upstream region were incomplete or altered. These results are evidence that the YSS is essential to the viral replication machinery, and contributes to replication enhancement and replication-associated translation activity in the RNA2 replicons.The terminal ends in the genome of RNA viruses contain features that regulate viral replication and/or translation. We have identified a Y-shaped structure (YSS) in the 3′ terminal regions of the bipartite genome of  Crinivirus (family Closteroviridae) are whitefly-transmitted, emerging plant viruses and affiliates of the alphavirus-like supergroup of single-stranded (ss), positive (+)-sense RNA viruses with large (15.3–17.6 kb) and complex genomes1Lettuce chlorosis virus (LCV) have been developed, allowing comparative inferences with Lettuce infectious yellows virus  and viruses in the related genus Closterovirus45785trans58Members of the genus large 15.–17.6 kb The 3′ terminus in the genome of RNA viruses contain secondary structures that, on their own or in combination with others through RNA-RNA interactions, are associated with many important functions, including the initiation and regulation of (−)-RNA synthesis, translation regulation, and virion encapsidation1314in vivo and in vitro translation assays using constructs containing the firefly luciferase (F-Luc) coding sequence flanked with the 5′ NCR and the YSS-containing 3′ NCR of LCV RNAs 1 or 2. We have also investigated the role of the YSS in supporting viral RNA synthesis by assessing the relative amounts of progeny RNA produced when 3′ NCR/YSS mutants of LCV RNA2 are co-inoculated with WT LCV RNA1 to tobacco protoplasts. The implications of our findings for conceptualizing the role of the YSS in viral RNA synthesis and translation for LCV and other criniviruses are discussed.In this study, using Selective 2′-Hydroxyl Acylation Analyzed by Primer Extension (SHAPE)161718Substantial studies have demonstrated that secondary structures of (+)-RNA viruses participate in various regulatory functions15202122232425in vivo translation assays using a series of F-Luc reporter constructs (see Methods), and translation efficiency was determined by the ratio of the F-Luc/R-Luc measurements . In constructs LUC-TMV, LUC-R1 and LUC-R2A, the F-Luc gene was flanked by the 5′ and 3′ NCRs of Tobacco mosaic virus (TMV) RNA, LCV RNA1 and LCV RNA2, respectively (Saguaro cactus virus (SCV) have shown that incorporating into luciferase constructs different number of additional nts from the 5′ proximal ORF of Sindbis virus sgRNA and SCV gRNA, respectively27To determine if the YSS contributes to viral translation, we performed ectively . Relativectively , demonstectively . These rectively , a modifectively , and botectively . Howeverectively . Studiesin vitro assays and the results were consistent with those of the in vivo assays -RNA and (+)-RNA synthesis of 3′Δ4 were comparable to that of the WT -RNA accumulating at approx. 39% compared to that of the WT , ΔSL1, or ΔSL2 inoculations at all time points  or the left arm (SLD1-2) of the lower stem of SL1 were substituted with the complementary sense nts . These miral RNA . Unlike iral RNA , synthesiral RNA .mfold to not affect the formation of the YSS in the resulting chimeric RNAs since 7 of the 9 nts in this region are identical between RNAs 1 and 2 (in vitro transcripts of the two chimeras and that of pCM1 (R1), and pCM2 (R2) were inoculated to tobacco protoplasts in various combinations [#1–#5]  and RNA2 (R2), resulting in chimeras pR1-3′R2 and pR2-3′R1 . Althoug 1 and 2 . The cap [#1–#5] . Viral Rin vivo, this begs the question of how might the cap-dependent translation of P5.6 be achieved. We took the first step towards addressing this question by determining whether or not in the presence of LCV RNA1, RNA transcripts produced from the R2 series of F-Luc constructs could function as replicons to support translation activity. The capped RNA transcripts of constructs: LUC-R2A, -R2B, -R2C, -R2CΔSL1, -R2CΔSL2, -R2D and -R2E were each co-inoculated with that of the internal control R-Luc construct and that of LCV RNA1 into tobacco protoplasts, and the cells were harvested at 72 hpi for analyses. (−)-RNA1 and (+)-RNA1 of LCV were both observed in the Northern blot analyses of all test samples (data not shown). In addition, high levels of (−)-RNA and (+)-RNA synthesis (determined using riboprobes specific to the F-Luc sequence) were observed in the LUC-R2C and -2E samples. In contrast, RNA synthesis in the LUC-R2A, -R2B, -R2CΔSL1, -R2CΔSL2, and -R2D samples were drastically reduced   are expr reduced . Remarka reduced , while t reduced . Interes reduced . These rClosteroviridae, a thorough understanding of these processes has been challenging given their large genomes and complex genome expression strategies9Successful proliferation within the host cell requires the genome of (+)-RNA viruses to undergo translation and replication. In many cases, these processes are aided by RNA structures located near the termini of the gRNAs. For viruses in the family Results from luciferase assays of LUC-R1 suggested that the 5′ and 3′ NCRs of RNA1 are involved in translation. This is not surprising since RNA1 encodes the replicase needed for viral replication. By contrast, similar regions from RNA2 alone, including those upstream of the YSS-containing 3′ NCR, did not support translation in the R2 series of luciferase replicons . On thisBrome mosaic virus (BMV), TMV and other (+)-RNA viruses14333435mfold predictions, deleting the nts in 3′Δ11 does not affect the overall structure of the YSS, while deleting those in 3′Δ24 eliminates the right basal stem of the YSS and disrupts its overall structure but leaves both SL1 and SL2 intact (not shown). This suggests that the YSS tolerates a disruption of the basal stem (S3), but cannot tolerate any SL disruptions. Disruptions of either SLs  abolished viral RNA synthesis Red clover necrotic mosaic virus, cap-independent translation of RNA2, which encodes the movement protein, is linked to its replication in trans by the RNA1-encoded replicaseTobacco rattle virusThe simplest explanation for the association between replication and translation involving LCV RNA1 and the R2 replicons is that the YSS facilitates the replicase produced from RNA151041424344mfold-predicted models generated using the full-length gRNA sequences of criniviruses revealed that structures similar to the YSS are strikingly conserved among other crinivirus genomes  or even trans-activator structural element(s) present in LCV RNA14547Further studies are needed to delineate the regions/nts of the YSS involved in viral RNA synthesis enhancement, and also to investigate the mechanisms underlying the replication-associated translation of replicons, including issues on whether translation is coupled to de novo replicon replication, the possible involvement of Structure cassette plasmids used for SHAPE analysis were engineered with the full-length cDNA sequences of LCV RNA1 (pS-CM1), LCV RNA2 (pS-CM2), LCV RNA2 without SL1 (pS-ΔSL1) and LCV RNA2 without SL2 (pS-ΔSL2). The construction of these plasmids involved using two independent PCR-amplifications. Sequence information of all oligo-primers used for the PCRs is provided in F-Luc reporter constructs for translation analyses were engineered using pCM1, pCM2 and TMV30BGFPAll PCR-amplifications were performed using Herculase II fusion DNA polymerase with high fidelity proofreading capability (Agilent Technologies), and all cloned products derived from PCR-amplification were sequenced in both directions.in vitro synthesis of RNA transcripts using the T3 mMessage mMACHINE kit (Life Technologies). 11 μl of dH2O containing 2 pmol of the RNA transcript was heated at 95 °C for 2 min, and immediately placed on ice for 1 min. 6 μl of 3.3x RNA folding mix and 1 μl of 100 mM MgCl2 was added to the transcript , and the mixture was divided into two 9 μl portions and incubated at 37 °C for 10 min to allow the RNA to renatureet al.® reverse transcriptase (Invitrogen) under three consecutive sets of temperature and time conditions: 42 °C for 1 min, 52 °C for 25 min, 65 °C for 5 min using 5′ end-labeled-VIC and -NED 3′-linker primers , respectively. Note that the structure cassettes contain a linker region that folds into stable SL structures at the 3′ termini of the transcript upon synthesis. The 3′ linker is designed for the binding of the 5′ end-labeled-VIC and -NED 3′-linker primers and reverse transcriptase -treated sample with the sequencing sample containing the VIC 3′-linker primer. Following ethanol precipitation, cDNAs were resuspended in 10 μl of deionized formamide and then resolved by capillary electrophoresis. The electropherogram data of each reaction was processed and analyzed by the Qushape softwareRNAstructure (version 5.4)5 μg each of pS-CM1, pS-CM2, pS-ΔSL1 or pS-ΔSL2 were linearized by restriction enzyme digestion using NgoMIV. The linearized DNA served as a template for the criptase 5253. Twoin vitro synthesis of capped RNA transcripts using the T3 mMessage mMACHINE kit (Life Technologies). The Renilla luciferase (R-Luc) construct was linearized by restriction enzyme digestion using SmaI, and capped RNA transcripts were synthesized using the T7 mMessage mMACHINE kitin vivo translation, 3 pmol of the in vitro produced capped transcript of each F-Luc reporter construct along with 1 pmol of the in vitro synthesized capped R-Luc RNA transcripts were inoculated to half a million N. tabacum var. Xanthi protoplasts. In parallel, 1 μl of water was inoculated along with the R-Luc RNA transcripts as a negative control. Additionally, for inoculations that included LCV RNA 1, 2 μg (0.7 pmol) of in vitro transcripts synthesized using NgoMIV-linearized pCM1 were used. After 72 hr incubation at 26 °C, cells were harvested and processed using the Dual-Luciferase Reporter Assay System (Promega). Procedures for in vitro translation are provided in In vivo translation efficiency was determined by the taking the ratio of luminescence produced by F-Luc to that produced by the internal control R-Luc. Both the in vivo (in vitro (in vivo translation experiments involving the R2 series of luciferase constructs and LCV RNA 1 (F-Luc reporter constructs (5 μg) were linearized by restriction enzyme digestion using NgoMIV or KpnI (in the case of LUC-TMV). The linearized DNA then served as a template for the  in vivo  and in vin vitro  experimeCV RNA 1  were repin vitro transcription of pCM1, pCM2, the pCM1- and pCM2-engineered mutants followed those previously describedProcedures for the in vitro synthesized transcripts of WT RNAs 1 and 2 (or their engineered derivatives) were each inoculated to N. tabacum var. Xanthi protoplasts following the previously described procedure5in vitro transcripts were individually inoculated to 0.5 × 106 protoplasts, and the protoplasts from all three inoculations were combined into a 100 × 20 mm petri dish and incubated at 26 °C. 16 hours post-inoculation (hpi), the combined protoplasts were redistributed into three 60 × 20 mm petri dishes (10 ml per dish) and incubated at 26 °C until the first harvest at 24 hpi. The inoculated protoplasts were also harvested at 48 and 96 hpi. A similar approach was adopted for the in vivo translation experiments involving the R2 series of luciferase constructs and LCV RNA1  according to the manufacturer’s instructions. Approximately 2 μg of total RNA from each sample were analyzed by Northern Blot using DIG-labeled riboprobes II and VIII as previously describedin vitro transcribed with T7 RNA polymerase (Roche Applied Science). R2 (+)-sense replicons were detected using the (−)-sense F-Luc DIG-labeled riboprobe, which was generated by linearizing pFLuc with NcoI and in vitro transcribed with Sp6 RNA polymerase (Roche Applied Science).2 μg of the LCV RNA1 . Duplicain vitro synthesized transcripts of pCM2, pΔSL1 and pΔSL2 were each inoculated to N. tabacum var. Xanthi protoplasts following the procedures as described above, including the normalization step, except the inoculated protoplasts were redistributed at 3 hpi and harvested at 3, 12, and 18 hpi. Approximately 5 μg of the TRIzol®-extracted total RNA from each sample were subjected to DNase treatment . The treated RNA was run on 1% HEPES denaturing gel stained with ethidium bromide to quantitate and normalize the 18S rRNA for equal amounts of rRNA that will be used for cDNA synthesis. 2.5 μg of treated RNA was used to synthesize the first-strand cDNA using Superscript III reverse transcriptase (Invitrogen) and gene-specific oligo-primer LCV-99-AC .


DDX41 gene are identified from myelodysplastic syndrome and/or acute myeloid leukemia (MDS/AML) patients. The review focuses on DDX41, as well as its homolog Abstrakt in Drosophila, which is important for survival at all stages throughout the life cycle of the fly.RNA helicases are involved in almost every aspect of RNA, from transcription to RNA decay. DExD/H-box helicases comprise the largest SF2 helicase superfamily, which are characterized by two conserved RecA-like domains. In recent years, an increasing number of unexpected functions of these proteins have been discovered. They play important roles not only in innate immune response but also in diseases like cancers and chronic hepatitis C. In this review, we summarize the recent literatures on one member of the SF2 superfamily, the DEAD-box protein DDX41. After bacterial or viral infection, DNA or cyclic-di-GMP is released to cells. After phosphorylation of Tyr414 by BTK kinase, DDX41 will act as a sensor to recognize the invaders, followed by induction of type I interferons (IFN). After the immune response, DDX41 is degraded by the E3 ligase TRIM21, using Lys9 and Lys115 of DDX41 as the ubiquitination sites. Besides the roles in innate immunity, DDX41 is also related to diseases. An increasing number of both inherited and acquired mutations in  RNA helicases are enzymes that utilize the energy derived from NTP hydrolysis to unwind double-stranded RNA (dsRNA) molecules  of pathogens can be detected by cells of the innate immune response with the help of germline-encoded pattern recognition receptors (PRRs) to induce type I interferons (IFN)  alter the native DDX41 interactome especially for major components in the U2 and U5 spliceosomes. It was concluded that DDX41 defects lead to loss of tumor suppressor function due to altered pre-mRNA splicing and RNA processing . We found that mutations related to diseases are conserved except M155I mutation  germline heterozygous mutations have been reported associate with familial myelodysplasia syndrome (MDS)/acute myeloid leukemia (AML)  (No. 2014CB910400 and 2013CB911103), the National Natural Science Foundation of China , the Beijing Nova Program (Grant No. Z141102001814020) to S.O., Youth Innovation Promotion Association CAS (Grant No. 2013065) to S.O., and the special project of Ebola virus research from the president foundation of Chinese Academy of Sciences.MDS, myelodysplastic syndrome; AML, acute myeloid leukemia; BTK, Bruton’s tyrosine kinase; D-E-A-D, Asp-Glu-Ala-Asp; HCV, hepatitis C virus; IFN, interferon; IRF3, interferon regulatory factor 3; mDC, myeloid dendritic cells; PAMP, pathogen-associated molecular pattern; PRR, pattern recognition receptor; STING, stimulator of interferon genes; TBK1, TANK-binding kinase 1; TRIF, TIR-domain-containing-adapter-inducing interferon-βYan Jiang, Zhi-Jie Liu, and Songying Ouyang declare that they have no conflict of interest. This article does not contain any studies with human or animal subjects performed by the any of the authors.


Here we provide a detailed laboratory protocol for in vitro dual RNA-Seq that is readily adaptable to any host-bacteria system of interest.Dual RNA-Sequencing leverages established next-generation sequencing (NGS)-enabled RNA-Seq approaches to measure genome-wide transcriptional changes of both an infecting bacteria and host cells. By simultaneously investigating both organisms from the same biological sample, dual RNA-Seq can provide unique insight into bacterial infection processes and reciprocal host responses at once. However, the difficulties involved in handling both prokaryotic and eukaryotic material require distinct, optimized procedures. We previously developed and applied dual RNA-Seq to measure prokaryotic and eukaryotic expression profiles of human cells infected with bacteria, using  Chlamydia-infected eukaryotic cells as proof-of-principle , requiring rRNA depletion or mRNA enrichment strategies to ensure sufficient non-rRNA can be sequenced at reasonable cost  Total host-bacteria RNA extraction and purification; (2) next-generation sequencing (NGS) of total RNA; and (3) bioinformatic processing and statistical analysis of the host-bacteria transcriptome. Here we present a laboratory methodology for stage (1) of a typical  Total hon Figure . The proFor dual RNA-Seq, a typical workflow includes RNA extraction, eukaryotic, and prokaryotic rRNA depletion, library preparation, sequencing, and analysis Figure . HoweverThe choice of organism, both host and bacteria, is central to the question(s) of interest and will inform much of the experimental design. Host cells should be selected for their biological relevance in relation to the bacterium. The identification of host differentially expressed genes (DEGs) or isoforms relative to mock-infected controls is the standard approach; additional controls should also be considered to differentiate specific from non-specific host responses to the bacterium. The choice of bacterium is dependent on the infection system under study, and can be expanded to compare  transcriptional differences in the presence of different bacterial species, virulent vs. avirulent strains, or mutant vs. wild-type strains.The ability of a dual RNA-Seq experiment to accurately determine differential gene expression between conditions is contingent on obtaining minimal biological and technical variation, which can be addressed by managing the trade-off between the number of replicates and sequencing depth  and technical  reasons. Additionally, enough RNA should be available to complete the QC stages of the experiment, including RT-PCR, agarose gel electrophoresis, and Bioanalyzer analysis. At the sequencing stage, the method of library preparation will dictate how much RNA is required, which usually lies within the microgram to picogram range. However, a useful convention is that more RNA input will require less amplification during sequencing, resulting in superior library complexity. As a guide, for mammalian cell culture-based dual RNA-Seq experiments, one well of a six-well plate results in ~100 ng of host RNA and ~500 pg bacterial RNA.8 host reads and >1 × 106 bacterial reads are required for adequate coverage  or paired-end (PE) libraries should be prepared and whether reads should be strand-specific. For dual RNA-Seq we prefer paired-end reads where each RNA fragment is sequenced from both sides, as it doubles the number of reads, reduces the rate of alignment ambiguity, and enables the identification of novel splice isoforms (host). Strand-specific libraries are also recommended, as the usually high gene density and presence of overlapping genes on opposite strands seen in bacterial genomes can more accurately be measured, while allowing the identification of potential antisense transcripts. The capture of the entire transcriptome, including coding, non-coding, anti-sense, and intergenic RNAs enables the discovery of complex and global transcriptional events.When infecting host cells, the bacterial MOI is an important consideration—a suitable infection rate should be selected to maximize the transcriptional response from both organisms while minimizing the number of uninfected cells. Here we describe the use of an MOI of ~1.5 which allows for a ~100% infection; if additional measures are required to enrich the selection of infected cells the researcher can consider increasing the MOI, the sequencing depth, or physically separating infected from infected cells by FACS or laser microdissection , is not used, the host cells are seeded at ~60% confluency at the time of infection to ensure continued viability throughout the time-course of the study. Given the time-sensitive nature of the experiment, it is crucial to synchronize the initial bacterial infection by centrifugation, followed by the removal of dead or non-viable bacterial cells by washing twice with DPBS.The method of infection depends on the host cell and this protocol provides guidance for infecting mammalian cell lines. While the methodology is largely consistent for different cells, the type of media and the required supplements will differ and should be selected according to the host cell. Table To ensure high-quality data, dual RNA-Seq typically requires a relatively large amount of input RNA. Additionally, RNA integrity is critical to the success of the protocol, as degraded samples will result in low yields that are insufficient for sequencing  with optimized steps to be the most effective at extracting quality RNA from several host-bacteria pairings.There are several commercial kits available for performing the RNA extraction Table , but theTM kit is most effective for lysing both host and Gram negative bacteria. For Gram positive bacteria, we include an initial sonication step to break open the cell wall prior to extraction. In each case, both the host and bacterial cells are lysed, releasing endogenous RNases so it is critical to minimize the delay between host cell lysis and RNA extraction to avoid degradation. Host cells are lysed, host proteins digested, and total nucleic acid precipitated with isopropanol. One caveat of the MasterPure extraction kit is the residual contamination of genomic DNA, however we have found that two treatments with TURBO DNA-freeTM DNase (Thermo Fisher) is most effective, compared to phenol:chloroform and precipitation-based methods.The method of bacteria and host lysis is an important consideration and will depend on the infection system being investigated. We find that the enzymatic lysis included in the MasterPureWe recommend performing three real-time RT-PCR assays for human targets and one endpoint PCR assay for bacterial targets to confirm DNA removal and the presence of RNA. These assays are based on TaqMan® Gene Expression assays (Applied Biosystems) with primer and probe sets targeting beta actin, mitochondria-encoded ATP synthase 6, and eukaryotic 18S rRNA , enabling simultaneous reduction of both host and bacterial rRNA. It is important to note that this method will not enrich immature mRNAs or non-coding RNAs; specific target enrichment techniques that are outside the scope of this protocol should be considered if these are of experimental interest. Aliquots of rRNA-reduced samples may be then subjected to poly(A) depletion to further enrich host mRNA transcripts and separate mRNA from rRNA. Poly(A)-depleted and rRNA-depleted eluates can also be further purified before being combined for library construction. As a guide, 1 μg of input RNA will yield <80 ng of rRNA-depleted RNA. The remaining RNA is concentrated and purified with a RNA Clean & ConcentratorTM-5 kit (Zymo Research).As rRNA typically constitutes more than 95% of total RNA, rRNA depletion should be considered to maximize the recovery of mRNA and reduce sequencing depth. It is possible to avoid depletion steps by instead sequencing far more deeply and subsequently removing rRNA reads during analysis. However, the sequencing costs of this strategy are substantial, as most sequence reads will align to rDNA genes and be discarded. Further reductions of sequencing costs per base may permit this strategy in the coming years, but it is currently more cost-effective to deplete rRNA prior to sequencing. There are several commercial kits available for nuclease digestion and size-selection; this protocol utilizes both a hybridization-based rRNA depletion and poly(A)-based depletion step. For hybridization, cDNA oligonucleotides attach to complementary rRNA that is immobilized on magnetic beads; always ensure that the oligonucleotides are compatible with your organism(s) of interest. For this, we combine an equivalent volume of Ribo-Zero beads from both a Human/Mouse/Rat-specific and Gram-negative bacteria-specific Ribo-ZeroA260/A280 readings are measured. As a guide, an A260 of 1.0 is equivalent to about 40 μg/mL of RNA. The A260/A280 ratio is a measurement of protein contamination in the sample, where pure RNA should have a ratio of ~2.0. Note, however, that this method can suffer from low accuracy and biases introduced by protein contamination  is usually regarded as the gold standard for estimating RNA quality by calculating an RNA integrity number (RIN), an estimation of RNA degradation which ranges from 1 to 10, with 10 being the least-degraded and most quality RNA. However, the mixture of eukaryotic and prokaryotic RNA in a dual RNA-Seq sample confounds the RIN measurement and a reliable number cannot be obtained. Instead, the Bioanalyzer electropherogram is useful to detect the presence of carry-over rRNA and RNA degradation. For RNA quantification with the Bioanalyzer we generally use an RNA 6000 ladder as the reference and prefer to use RNA 6000 pico chips as the nanochips tend to suffer from reduced sensitivity. However, note that the accuracy of the assay will decrease if the maximum RNA concentration is exceeded for the chip. Please consult Table When preparing RNA for NGS, the broad steps include fragmentation and/or size selection, conversion to cDNA, adapter attachment, and quantification. NGS is a rapidly expanding field and there are currently a number of sequencing platforms currently available, including Illumina, SOLiD, Ion Torrent, Roche 454, Oxford Nanopore, and Pacific Biosciences, with specific advantages and library preparation kits associated with each. When deciding on a sequencing technology, the defining objective of the experiment should be considered; i.e., is the focus on all transcriptional events , just the coding mRNA transcripts, or just small RNAs  and enriched via polymerase chain reaction (PCR) amplification. Paired-end reads >50 nucleotides will promote increased fragment randomization; longer reads will enable greater coverage, reduced multi-mapping, and improved transcript identification  to ensure that the majority of reads are derived from the two organisms of interest. This is followed by an assessment of read quality with FASTQC (www.bioinformatics.babraham.ac.uk/projects/fastqc/) and the use of Trimmomatic  method for both host and bacterial reads, which provides between-sample normalization while correcting for variations in sequencing depth and sample variation .Bacterial species of interest CAUTION Highly flammable. Causes skin and serious eye irritation. Handle using appropriate safety equipment.Ethanol  Isopropanol Poly(A)Purist Mag Purification Kit freeTM kit TURBO DNA-DEPC-treated water TM Complete DNA and RNA Purification Kit MasterPureTM-5 RNA Clean & ConcentratorTM rRNA Removal Kit (Human/Mouse/Rat) Ribo-ZeroTM rRNA Removal Kit (Gram-negative bacteria) Ribo-ZeroSonicator S-4000  (for use when lysing Gram positive bacteria)TaqMan® Gene Expression Assay High-Capacity cDNA Reverse Transcription Kit Beta actin probe Mitochondrially encoded ATP synthase 6 probe Eukaryotic 18S rRNA probe MicroAmp® optical 96-well reaction plates MicroAmp® optical adhesive film MicroAmp® optical film compression pad Flask rocker DNase and RNase-free 1.5 mL microcentrifuge tubes Refrigerated microcentrifuge Centrifuge Vortex Heating block (x2) IceIce bucketTimerReal-time qPCR machine Temperature cycler 2 Incubator, 37°C, 5% COBioanalyzerRNA 6000 Pico Kit 6-well plates Cell scrapers 15 mL centrifuge tubes 70% ethanol Combine 350 mL ethanol with 150 mL nuclease free water. Store at room temperature for up to 6 months.TM rRNA Removal KitRibo-Zero The Ribo-Zero kits are composed of two parts: the magnetic core kit and the rRNA removal reagents. Store the magnetic core kits at 4°C and Mouse/Human/Rat and Gram-negative bacteria rRNA removal reagents at −80°C.TM-5RNA Clean & Concentrator Add 48 mL of 100% ethanol to the 12 mL RNA Wash Buffer before use.5 mammalian host cells per well in all wells of a six-well plate. Ensure there are two plates per time-point (one infected plate and one non-infected control plate). Incubate plates overnight at 37°C, 5% CO2.1. Seed × 10Following seeding, sit plates on a bench at room temperature for 15 min to allow cells to settle, ensuring an even distribution of cells.The next day, the cells should be evenly distributed throughout the well.2. The next day, infect all plates (except mock-infected control plates) at an MOI of 1.5 to ensure that 100% of the host cells will be infected.Rock plates for 30 mins on shaker.2, according to time-point.3. Centrifuge plates at 500 × g for 30 min at room temperature. Incubate plates at 37°C, 5% COCentrifugation is important to synchronize the infections and time-points and ensure an MOI of 1.5.2.4. Wash cells twice with DPBS and overlay with warm, fresh media. Incubate at 37°C, 5% CO5. At each time-point, wash cells twice with ~3 mL DPBS and add 1 mL DPBS to each well. Harvest cells with a cell scraper and dispense solution into a 15 mL centrifuge tube.Ensure all cells are collected from each well. It may be beneficial to add a second 1 mL of DPBS to collect any remaining cells.Store tubes at −80°C until all time-points are complete. Cells can be stored for up to 6 weeks at −80°C.6. Remove centrifuge tubes from −80°C freezer and thaw at room temperature.7. Pre-set heating block to 65°C.TM Complete DNA and RNA Purification Kit; Epicenter) to 300 μL of Tissue and Cell Lysis Buffer  for each sample.8. Add 1 μL of 50 μg/mL Proteinase K  to each 25 μL sample and mix thoroughly by vortexing.For systems containing Gram positive bacteria:12. Set sonicator to an amplitude setting of 10 and perform six treatments of 30 s each to disrupt the cells. Place tubes on ice for 30 s between each treatment.Incomplete sonication can lead to a reduction in bacterial RNA yield.For all systems:13. Incubate tubes in a heating block at 65°C for 15 min, vortexing briefly every 5 min.The solution should become cloudy as cells are lysed and proteins are digested.14. Place samples on ice for 3–5 min.TM RNA Purification Kit; Epicenter) to each 300 μL of lysed sample and vortex vigorously for 10 s.15. Add 175 μL of MPC Protein Precipitation Reagent  to a clean 1.5 mL microcentrifuge tube and discard the pellet.18. Add 500 μL of isopropanol to the recovered supernatant and invert the tube 30–40 times. Do not vortex.Isopropanol will desalt and precipitate the nucleic acid. It may be beneficial to mark the side of the tube prior to centrifugation as pellets from isopropanol treatment have a glassy appearance that may be difficult to see.19. Pellet the total nucleic acid by centrifugation at 10,000 × g for 10 min at 4°C.20. Carefully pour off the isopropanol without dislodging the pellet.Pellets from isopropanol treatment are loosely attached and care should be taken when pouring.21. Rinse the pellet twice with 70% ethanol, being careful not to dislodge the pellet. Centrifuge briefly if pellet is dislodged.Ethanol is more volatile than isopropanol and will make the DNA easier to dissolve. Perform at room temperature.TM RNA Purification Kit; Epicenter). Samples can be stored in TE Buffer overnight at 4°C.22. Remove residual ethanol with a pipette and resuspend the pellet in 35 μL of TE Buffer  to each 35 μL sample.24. Add 4 μL of 10x TURBO Dnase Buffer  and incubate tubes at room temperature for 5 min. Mix occasionally by manually inverting.27. After incubation, add 8 μL of DNase Inactivation Solution  to a fresh 1.5 mL microcentrifuge tube.The pellet should be visible; if not, vortex briefly and repeat centrifugation for 5 mins. Take care not to dislodge the pellet when pouring.30. Remove 5 μL from each RNA sample and place in a clean 1.5 mL microcentrifuge tube.Keep all samples and reactions on ice.31. Add 95 μL of nuclease-free water (1:20 dilution).32. Prepare a 2X RT master mix for each sample in triplicate. Include a non-template control and an RT- control.10X RT buffer: 2 μL25X dNTP mix (100 mM): 0.8 μL10X RT random primers: 2 μLRNase inhibitor: 1 μL2O: 3.2 μLRNA-free dH33. Prepare 1X reactions  by combing the 2X master mix with the following in individual wells of a 96-well reaction plate (Thermo Fisher):2OReaction 1: 10 μL sample + 2 μL RT + 1 μL dH2OReaction 2: 10 μL sample + 3 μL dH2O + 2 μL RTReaction 3: 11 μL dHGently pipette up and down to mix.Seal the plate.34. Briefly centrifuge plate to spin down the solution and eliminate any air bubbles.35. Perform the reverse transcription reaction according to the following conditions:25°C, 10 min7°C, 120 min85°C, 5 mincDNA can be stored at −20°C. Create small aliquots to prevent degradation resulting from frequent freeze-thawing.36. Run 20 μL of each reaction on a 1% agarose gel and examine under UV fluorescence.There should be large bands in the lane containing Reaction 1, indicative of cDNA. The lane containing Reaction 2 should be free of bands, otherwise contamination with gDNA has occurred. The lane containing Reaction 3 should also be free of bands. See Table 37. Make a 1:20 dilution of cDNA, prepared above.38. Prepare enough RT-PCR master mix to assay each sample in triplicate, as well as controls.20x TaqMan® Gene Expression Assay: 1 μL2x TaqMan® Gene Expression Master Mix: 10 μLRNase-free water: 5 μL39. Cap the tube and invert the tube several times to mix the reaction components. Pulse vortex.40. Aliquot 15 μL of master mix into individual wells of a 96-well reaction plate (Thermo Fisher). Include a triplicate reaction for each sample and non-template controls.41. Add 4 μL of each diluted template to appropriate wells and gently tap plate on benchtop to distribute contents to the base of the well.42. Place adhesive film (Thermo Fisher) over the plate and seal with compression pad (Thermo Fisher).If any bubbles are visible in the wells or liquid is present on the sides of the wells, centrifuge plate at 500 × g for 2 min. Do not touch the film with ungloved hands at any point.43. Place plate in RT-PCR machine and run assay according to the following cycling conditions:Hold: 95°C, 10 minCycle (40x): 95°C, 15 s60°C, 1 min.Amplification curves should be present for all targets, representing the accumulation of product throughout the RT-PCR experiment. Ensure that the amplification signal is at least ten times the baseline standard deviation to indicate amplification over noise. See Table 44. Pre-set one heating block to 68°C and one at 50°C.TM rRNA Removal Magnetic Core Kit from 4°C and allow to warm to room temperature. Remove Human/Mouse/Rat and Gram-negative bacteria components of the Ribo-ZeroTM rRNA Removal kits from −80°C and thaw on ice.45. Remove Ribo-ZeroTM Magnetic Core Kit on ice as this can damage them and reduce the efficiency of the procedure.Do not place the Ribo-ZeroTM rRNA Removal kit; Illumina) for 20 s by vortexing.46. Vigorously mix magnetic beads ; two tubes per sample.47. Carefully pipette 65 μL of magnetic beads into 2 mL microsphere wash tubes  to each tube. Vortex vigorously.50. Wash the microsphere wash tubes by adding 130 μL of microsphere wash solution  to each tube and vortex vigorously until a homogenous suspension is produced.52. Add 65 μL of microsphere resuspension solution  to each tube. Mix by vortexing for 10 s and set aside (at room temperature).53. Add 1 μL of RiboGuard RNase inhibitor  according to the following preparation :54. Treat two aliquots of each sample with Ribo-Zero rRNA removal solution : 1 μLRNase-free water : 5 μLRibo-Zero rRNA Removal Solution (Human/Mouse/Rat kit): 5 μLFully mix the samples by pipette-mixing 10-15 times.The beads may clump at this stage due to the biotin on the probes binding to multiple beads. If so, vortex for at least 10 s.55. Incubate at 68°C for 10 min in heating block. Return the Ribo-Zero reaction buffer to −80°C.56. Remove the microsphere wash tubes from the heating block and incubate at room temperature for 15 min.57. Vortex the microsphere wash tubes at medium speed for 20 s to ensure a homogenous slurry.58. Pipette hybridized RNA sample to the resuspended microsphere wash tubes, pipette-mixing 10–15 times to mix. Immediately vortex the microsphere wash tubes at medium speed for 5 s.to the magnetic beads) is critical for rRNA removal efficiency.The washed magnetic beads must be at room temperature for use in this step. The order of the addition  and centrifuge at 12,000 × g for 1 min at room temperature. Save the eluate and discard the removal unit.62. Transfer the RNA-microsphere suspension to a Microsphere Removal Unit  to each volume of RNA sample and mix well.63. Add 2 volumes of RNA Binding Buffer  in a collection tube and centrifuge at 12,000 × g for 1 min. Discard the flow-through.65. Transfer the mixture to a Zymo-Spin IC column  to the column and centrifuge at 12,000 × g for 1 min. Discard the flow-through.67. Add 400 μL of RNA Prep Buffer  to the column and centrifuge at 12,000 × g for 1 min. Discard the flow-through.68. Add 800 μL of RNA Wash Buffer  molecular weight smearing (> ~400 bp) will be the eluted RNA and can be expected. See Table A260/A280 ratio.74. Measure the quality and quantity of eluted RNA by assaying a 1 μL on a Nanodrop instrument. Observe absorbance readings at 260 nm, as well as the A260 of 1.0 is equivalent to about 40 μg/mL of RNA. The A260/A280 ratio is a measurement of protein contamination in the sample, where pure RNA should have a ratio of ~2.0.The RNA concentration should be sufficient for the sequencing requirements (check the sample submission guidelines for the sequencing technology that will be used). An http://www.genomics.agilent.com/en/Bioanalyzer-System/2100-Bioanalyzer-Instruments/?cid=AG-PT-10675. Measure the quantity and integrity of RNA on a Bioanalyzer instrument. Consult the manufacturer's guidelines for the assay: If carryover rRNA is present, the electropherogram will produce two distinct peaks corresponding to 18S and 28S eukaryotic rRNA , indicating that rRNA-depletion was not successful. Degraded RNA can be identified by any peaks appearing at the lower part of the electropherogram, which highlights a distribution shift toward smaller RNA fragments. See Table The method of library preparation is dependent on the sequencing platform used and thus is outside the scope of this protocol. Commercial sequencing enterprises and sequencing centers will provide detailed guidelines on the library preparation and sample submission guidelines that begin with the isolation of pure mRNA as described above. Following sequencing, the user will be provided with a series of raw FASTQ sequence files that serve as the input for the subsequent computational data analysis.JM, MH designed the protocol. JM, MH, GM wrote the manuscript.The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.