Pat1, known as Pdc2 in fission yeast, promotes the activation and assembly of multiple proteins during mRNA decay. After deadenylation, the Pat1/Lsm1–7 complex binds to transcripts containing oligo(A) tails, which can be modified by the addition of several terminal uridine residues. Pat1 enhances Lsm1–7 binding to the 3′ end, but it is unknown how this interaction is influenced by nucleotide composition. Here we examine Pat1/Lsm1–7 binding to a series of oligoribonucleotides containing different A/U contents using recombinant purified proteins from fission yeast. We observe a positive correlation between fractional uridine content and Lsm1–7 binding affinity. Addition of Pat1 broadens RNA specificity of Lsm1–7 by enhancing binding to A-rich RNAs and increases cooperativity on all oligonucleotides tested. Consistent with increased cooperativity, Pat1 promotes multimerization of the Lsm1–7 complex, which is potentiated by RNA binding. Furthermore, the inherent ability of Pat1 to multimerize drives liquid–liquid phase separation with multivalent decapping enzyme complexes of Dcp1/Dcp2. Our results uncover how Pat1 regulates RNA binding and higher order assembly by mRNA decay factors. A dense network of protein–protein interactions regulates 5′–3′ mRNA decay, which is important for gene expression and many physiological processes including development, microRNA-mediated decay, and quality control mechanisms . Bulk 5′Many mRNA decay factors, including Pat1, are enriched in processing-bodies (P-bodies), which are a class of membraneless organelles that may function in mRNA storage or decay . At the Pat1 uses a combination of disordered and globular domains to interact with and activate multiple mRNA decay factors. The disordered amino terminus contains a conserved FDF motif that interacts with Dhh1 (DDX6 in humans) and potentiates P-body formation, but is largely dispensable for function . The unsWhile it is known how Pat1 activates different mRNA decay factors, much less is understood about how it affects specific RNA recognition by Lsm1–7. In vitro, budding yeast Pat1/Lsm1–7 shows a preference for oligoadenylated RNAs compared to those containing poly(A) tails; however, genome-wide CLIP studies indicate Pat1/Lsm1 co-occupy the 3′ untranslated region (UTR) of budding yeast transcripts without enriching a specific sequence motif . In fissSchizosaccharomyces pombe Pat1/Lsm1–7 complex binding to a series of oligonucleotides of different A/U content. Lsm1–7 alone has a binding preference for U-rich RNAs. Addition of Pat1, however, broadens the specificity of the Lsm1–7 complex by enhancing binding to A-rich targets. Furthermore, Pat1 increases cooperative binding of Lsm1–7 to oligonucleotides, which drives multimerization of the heterooctamer on RNA in a sequence independent manner. Oligomerization is an inherent property of Pat1 that permits higher order assembly with multivalent Dcp1/Dcp2 complexes, which can recruit additional mRNA decay machinery. Taken together, this work reveals how Pat1 broadens the specificity of Lsm1–7 and promotes the assembly of higher order decapping complexes.In this work, we evaluate recombinant purified S.pombe Lsm1–7 complexes alone or coexpressed with PatMC for their ability to bind different oligo-RNAs by fluorescence polarization . Furthermore, the Lsm1–7 complex alone strongly favored binding to U-rich 15mers, which was not affected by PatMC. In the absence of PatMC, Lsm1–7 preferentially bound A-tailed RNAs with similar uracil content, suggesting that nucleotide context may be an additional factor for Lsm1–7 binding RNA  are sufficient to support cell growth in yeast . To underization A,B. Becarization . As seenrization . The folding RNA C. BecausIn addition to the differences in affinities for the oligonucleotides, we also observed a difference in Hill coefficients, a measure of cooperativity that places a lower bound on the number of Pat1/Lsm1–7 complexes binding RNA. The binding isotherms of the PatMC/Lsm1–7 complex were consistently approximately twofold more cooperative than Lsm1–7 alone for all tested RNAs, suggesting that PatMC/Lsm1–7 may be engaging A15 and U15 RNA in a different manner D. SpecifTo directly test the number of PatMC/Lsm1–7 complexes bound to short oligonucleotides, we used size exclusion chromatography coupled to multiangle light scattering (SEC-MALS). The SEC step fractionates protein complexes by hydrodynamic radius and molar mass is concurrently detected by light-scattering and differential refractometry . The PatSupplemental Fig. S2A–C). This suggests that the tetrameric PatMC/Lsm1–7/RNA complex is less stable than the dimeric species.To evaluate the stability of the oligomeric assemblies, we performed sedimentation velocity analytical ultracentrifugation (SV-AUC) on PatMC/Lsm1–7 with stoichiometric amounts of A15 RNA over the course of 12 h. While we could detect a strong dimeric peak in the sedimentation distribution, there was a minimal amount of tetrameric assemblies C. FurtheWe next asked how PatMC/Lsm1–7 assembled on U15 RNA. As seen with A15 RNA, addition of stoichiometric amounts of U15 to the PatMC/Lsm1–7 complex resulted in two peaks by SEC-MALS with molar masses corresponding to dimeric and tetrameric assemblies A,B. Thisd. Binding of labeled U15 was followed by fluorescence anisotropy. Titration of Lsm1–7 alone or the PatMC/Lsm1–7 complex results in saturation of the anisotropy signal at one equivalent of RNA, indicating 1:1 binding between PatMC/Lsm1–7 and the U15 oligonucleotide . This indicates that each PatMC/Lsm1–7 heterooctamer binds a single oligo-RNA, though we cannot exclude the possibility that PatMC/Lsm1–7 can co-occupy RNA sequences longer than 15 nt tested here. Therefore, we conclude that RNA ligands of different sequences and lengths induce stable dimerization of the PatMC/Lsm1–7 complex.It is possible that each PatMC/Lsm1–7 complex binds an individual RNA or that multiple PatMC/Lsm1–7 complexes co-occupy a single RNA to promote oligomerization. To test these possibilities, we determined the stoichiometry of PatMC/Lsm1–7 binding to RNA. Experiments were performed under saturating conditions, where concentration of the oligo-RNA was far above the Kcleotide D. SimilaAs PatMC/Lsm1–7 binds to RNA as a higher order assembly, we asked if the complex had the intrinsic ability to multimerize in the absence of RNA. While PatMC/Lsm1–7 initially purified as a monomer, concentration and subsequent SEC-MALS of the complex in the absence of RNA revealed two peaks with molar masses corresponding to monomeric and dimeric PatMC/Lsm1–7 complexes A–C. ThisSupplemental Fig. S4). Because Lsm1–7 was monomeric in the absence of PatMC, the above results indicate PatMC drives multimerization of Lsm1–7, which may be enhanced by RNA binding  .Previous studies demonstrate that the monomeric, globular carboxy-terminal domain of Pat1 can bind helical leucine motifs (HLMs) in the disordered carboxyl terminus of Dcp2 . Known dExt) A. ThoughExt for droplet formation, we queried how individual regions of both complexes contribute to phase separation. The carboxy-terminal domain of Pat1 is monomeric and binds HLMs, but did not promote phase separation of Dcp1/Dcp2Ext (HLM1) (To understand the requirements of PatMC and Dcp1/Dcp2/Dcp2Ext B. Additit (HLM1) B. These Ext droplets were able to incorporate A15 RNA, though A15 RNA did not change the critical concentration required for phase separation . However, neither short A15 RNAs nor poly(A) RNA promoted droplet formation with either PatMC  . The Pater PatMC C. RNA coExt condensates, indicating that PatMC can bridge both 5′ (Dcp1/Dcp2) and 3′ (Lsm1–7) decay factors in the context of the phase separated droplet . Furthermore, Dcp1/Dcp2Ext/PatMC/Lsm1–7 condensates recruited RNA . These observations suggest that Pat1 may bridge both 5′ and 3′ activities in the context of a phase separated droplet.We next asked if other mRNA decay factors could be incorporated in PatMC/Dcp1/Dcp2 droplets. Lsm1–7, but not a nonspecific protein such as the unrelated Spycatcher protein, was recruited into preformed PatMC/Dcp1/Dcp2Our biochemical reconstitution uncovers how Pat1 broadens the specificity of the Lsm1–7 complex and promotes higher order assembly of multiple mRNA decay factors. First, Pat1 expands the Lsm1–7 complex's sequence preference by enhancing binding to adenine-rich RNAs. Second, PatMC promotes cooperative binding of Lsm1–7 to RNA, which drives oligomerization on nucleic acid. Third, the PatMC/Lsm1–7 complex has the inherent ability to oligomerize, which is dependent on Pat1 and consistent with coimmunoprecipitation data in fission yeast . FinallySupplemental Fig. S1G; The Lsm1–7 ring is the high affinity RNA binding component of the Pat1/Lsm1–7 complex and has a preference for U-rich oligonucleotides . Pat1 brSupplemental Fig. S2). As this work was in review, a crystal structure of S. pombe Lsm1–7 with U5A1 RNA was reported, demonstrating binding to the distal face of the ring, similar to the Lsm2–8 complex  was ordered from IDT and cloned into site two of the pET-Duet vector. Cells were harvested by centrifugation and lysed in appropriate buffer. For Lsm1–7 and PatMC/Lsm1–7 complexes, cells were lysed in Buffer A  by sonication. Lysate was subsequently clarified by centrifugation and the supernatant was bound to Ni-NTA resin (GE) at 4°C for 1 h. The resin was then transferred to a gravity column and washed with 20 column volumes of Buffer A before being eluted in 25 mL of Buffer E . The elution was then loaded directly onto a 5 mL HiTrap Heparin column (GE). The heparin column was run at 2 mL/min from a 0.25–1 M NaCl gradient over 20 column volumes. Fractions containing the appropriate protein complex were concentrated in 30 kD concentrators (Amicon) to ∼2 mL before adding TEV overnight at 4°C. The following day, the sample was filtered and further purified by gel filtration using a Superdex 200 16/60 column (GE). Coexpressed PatMC/Lsm1–7 was purified in 400 mM NaCl, 20 mM HEPES 7.0, 1 mM DTT and Lsm1–7 alone was purified in 150 mM NaCl, 20 mM HEPES pH 7.0, 1 mM DTT. Fractions containing protein were concentrated before being flash frozen and stored at −80°C.All proteins were expressed in BL21(DE3)* (Thermofisher) cells in LB media. Cells were grown to ODAll MBP-Pat1 fusions were purified as described previously . For Spy2 with 0.3 µg/µL acetylated BSA (Promega). All RNAs used were labeled with 5′ FAM (IDT) and were used at final concentration of 500 pM. The binding experiments were conducted at 22°C. All binding curves were fit to the following Hill model for single site binding:G was determined by the relationship ΔG = +RT * ln(Kd). The ΔG from each individual fit was averaged and plotted with standard deviation. Hill coefficients were averaged from fitting three separate binding isotherms and shown with standard deviation.All fluorescent polarization experiments were performed in 200 mM NaCl, 20 mM HEPES pH 7.0, 1 mM DTT, 5 mM MgClFor analysis of stoichiometry, 5′-FAM RNA was kept at 0.5 µM in the same buffer as used for the fluorescence polarization assay. Protein was titrated into solution containing labeled RNA and fluorescence polarization was measured for each concentration. The linear portion was fit to a linear model and the average of the last four points was used to fit a line at the saturation point. The intersection of these two lines was used to determine the binding stoichiometry.Analytical SEC was performed in buffer M , or in the appropriate NaCl concentration. Samples were mixed at ∼30 µM and incubated for 15 min in 350 mM NaCl, 20 mM HEPES pH 7.0, 1 mM DTT on ice before being filtered and injected onto a GE Superdex 200 10/300-Increase analytical size exclusion column. When appropriate, samples were mixed with 1.1-fold molar excess RNA. All samples were run at 0.35 mL/min, and peaks were analyzed by SDS-PAGE (Invitrogen). For experiments involving reinjection of fractions over SEC, 500 µL of fractions were spin filtered before reinjecting over SEC.For SEC-MALS, 165 µg of sample was filtered through a 0.1 µm spin filter (Amicon) before being injected onto a preequilibrated KW-804 column (Shodex). For samples with RNA, stoichiometric amounts of RNA were added prior to spin filtration. Data was acquired with an inline DAWN HELEOS MALS and Optilab rEX differential refractive index detector (Wyatt Technology). All analysis was performed using ASTRA VI software (Wyatt Technology). Data was then exported and plotted with R.Sample was buffer exchanged into buffer M using Zeba spin columns (Thermofisher) and diluted to 9.3 µM. When appropriate, stoichiometric amounts of RNA were added to the sample after buffer exchange. AUC cells were assembled according to manufacturer's protocol and 100 µL of sample was loaded into the cell. The sample was incubated at 22°C for >2 h prior to centrifugation. Samples were run at 30,000 rpm for 10–12 h in a Beckman XL/A analytical ultracentrifuge. Scans for samples containing only protein were collected at 280 nm, and samples containing RNA were scanned at both 280 and 260 nm. Sedimentation velocity analysis was performed in SEDFIT (NIH) and plots were generated with GUSSI . ExperimFor labeling with dyes, proteins were buffer exchanged into appropriate labeling buffer using Zeba spin columns (Thermofisher). Lsm1–7 and SpycatcherKCK were labeled with fivefold molar excess Alexa Fluor 555 maleimide for 1 h at room temperature in 150 mM NaCl, 20 mM HEPES pH 7.5, 0.5 mM TCEP. Reactions were quenched by addition of βME to a final concentration of 10 mM. MBP-PatMC was labeled with fourfold molar excess NHS-Fluorescein (Thermofisher) for 1 h at room temperature in 150 mM NaCl, 150 mM Sodium bicarbonate pH 8.4 before being quenched by adding TRIS-HCl pH 8.0 to a final concentration of 50 mM. All quenching steps were performed at room temperature for 20 min. Free dye was separated from labeled protein by Illustra NICK columns (GE) according to the manufacturer's instruction. Labeling efficiency and concentrations were calculated by UV-vis spectroscopy.i equipped with a 40× dry lens. Samples were prepared in a 384 well plate (Greiner) that was cleaned with 0.1 M NaOH and passivated with PEG-silane and 100 mg/mL BSA (Sigma-Aldrich) before being washed with water to remove residual BSA. Proteins or RNA were mixed at specified concentrations in a final buffer concentration of 60 mM NaCl, 20 mM HEPES pH 7.0, 1 mM DTT. When appropriate, dye-labeled protein or RNA were added to 100 nM. Samples were incubated at room temperature for 20 min prior to imaging. Images were analyzed in FIJI (All images were acquired with Nikon Eclipse T in FIJI .Supplemental material is available for this article.


A simple method for extraction of high quality RNA from cells that have been fixed, stained and sorted by flow cytometry would allow routine transcriptome analysis of highly purified cell populations and single cells. However, formaldehyde fixation impairs RNA extraction and inhibits RNA amplification. Here we show that good quality RNA can be readily extracted from stained and sorted mammalian cells if formaldehyde is replaced by glyoxal—a well-characterised fixative that is widely compatible with immunofluorescent staining methods. Although both formaldehyde and glyoxal efficiently form protein-protein crosslinks, glyoxal does not crosslink RNA to proteins nor form stable RNA adducts, ensuring that RNA remains accessible and amenable to enzymatic manipulation after glyoxal fixation. We find that RNA integrity is maintained through glyoxal fixation, permeabilisation with methanol or saponin, indirect immunofluorescent staining and flow sorting. RNA can then be extracted by standard methods and processed into RNA-seq libraries using commercial kits; mRNA abundances measured by poly(A)+ RNA-seq correlate well between freshly harvested cells and fixed, stained and sorted cells. We validate the applicability of this approach to flow cytometry by staining MCF-7 cells for the intracellular G2/M-specific antigen cyclin B1 (CCNB1), and show strong enrichment for G2/M-phase cells based on transcriptomic data. Switching to glyoxal fixation with RNA-compatible staining methods requires only minor adjustments of most existing staining and sorting protocols, and should facilitate routine transcriptomic analysis of sorted cells. High-throughput RNA sequencing (RNA-seq) is increasingly used as a first-pass method to characterise cell populations, providing detailed and robust data through standardised pipelines. In parallel, purification of cell populations by flow cytometric cell sorting has become routine, with the capacity to isolate cells based on simultaneous quantification of many antigens. It would seem natural to unite these techniques, such that carefully purified cell populations are characterised by RNA-seq as standard, however this has proved surprisingly problematic primarily due to difficulties in RNA recovery from fixed and stained cells.Ideally, cells destined for RNA-seq analysis should be fixed before removal from growth media or tissue context as tissue dissociation and cell staining methods induce stress responses that affect the transcriptome –3. FormaAlthough RNA recovered from formaldehyde-fixed cells is often of low quality, formaldehyde is a standard reagent in RNA electrophoresis and does not cause RNA degradation of itself . InsteadOf course, other fixatives are known that do not crosslink RNA to protein or introduce stable adducts. We and others routinely employ ethanol fixation on yeast prior to RNA isolation and find the fixation speed and preservation of RNA to be excellent , 25, whiNon-RNA-protein crosslinking fixatives therefore have the potential to be extremely useful in transcriptomic studies of flow sorted cells. However, little is known about the survival of RNA in alcohol fixed cells during downstream staining and sorting procedures, and to our knowledge the survival of RNA in glyoxal fixed cells has not been investigated. Here we show that glyoxal fixation allows extraction of high-quality RNA using standard protocols from stained and sorted cell samples, and that this RNA can be processed into high quality RNA-seq libraries.To determine the effectiveness of ethanol fixation in preserving intact and accessible RNA in mammalian cells, we extracted RNA from COLO205 cells that were either unfixed, fixed with 70% ethanol or fixed with 4% formaldehyde. RNA extraction was performed using TRI reagent, a monophasic GTC-phenol RNA extraction solution, and 20% of the RNA obtained was analysed on denaturing RNA mini-gels. Plentiful high quality RNA was obtained from the unfixed cells, whereas we did not recover detectable RNA from formaldehyde-fixed cells which was as expected given that proteinase K digestion was not performed  as a trial intracellular antigen for validation of RNA-seq analysis in glyoxal fixed and stained cells. CCNB1 accumulates during G2 phase of the cell cycle and is degraded at the end of M-phase, so the quality of staining and sorting based on CCNB1 can be verified by comparison to DAPI staining for DNA content. Furthermore, gene expression changes across the cell cycle are well characterised and so the successful purification of G2/M-phase cells should be detectable based on differential expression of cell cycle genes in RNA-seq data.To determine whether our staining protocol reproducibly maintained RNA quality, we compared cells immediately dissolved in TRI Reagent at harvest to cells that underwent staining. Antibody staining was performed according to the consensus protocol given in the Materials and Methods section entailing glyoxal fixation, methanol permeabilisation and two-step immunostaining with rabbit anti-CCNB1 followed by Alexa Fluor 488-labelled donkey anti-rabbit. Two replicate experiments were performed for COLO205 cells and two for MCF-7 cells, RNA was extracted with TRI reagent and RNA quality assessed by Bioanalyzer. Representative Bioanalyzer plots are shown  and deta2-transformed read-counts for all annotated genes in the directly harvested compared to the fixed and stained samples correlated well (2 = 0.989 for MCF-7 and R2 = 0.991 for COLO205), showing no major systematic change in mRNA distribution. Gene expression analysis between the sets of directly harvested versus fixed and stained cells by DEseq2 (p<0.05) identified 2 and 61 differentially expressed genes in MCF7 and COLO205 datasets respectively out of 60099 genes annotated in Ensembl GRCh38 + RNA Purification Module according to manufacturers’ instructions. RNA inputs and final library yields are given in l GRCh38 , indicat5 cells of each fraction were harvested, RNA was extracted using TRI Reagent, analysed on a Bioanalyzer, and mRNA-seq libraries prepared and sequenced as above. The correlation between the two biological replicates of CCNB1-positive cells was excellent (2 = 0.992), whilst the CCNB1-negative fractions were slightly more divergent indicating differences between the cell cultures, although the correlation was still very strong (2 = 0.988). DEseq2 analysis (p<0.05) identified 161 significantly different expressed genes between CCNB1-positive and CCNB1-negative samples  along with ~300 other significant enrichments (q<0.05) almost all of which are directly connected to cell cycle progression  and can be efficiently reverse transcribed.Glyoxal fixative is easy to prepare and use, and should be straightforward to substitute for formaldehyde in most protocols. However we find that staining protocols do need to be modified to maintain RNA quality: firstly, inclusion of RNasin or an equivalent placental RNase inhibitor proved to be important. Secondly, we performed antibody incubations on ice to further reduce RNase activity, though we suspect this is not absolutely required. Thirdly, some staining reagents are clearly incompatible with RNA isolation—harsh detergents such as Triton X-100 allow RNA to escape the cell (an unavoidable consequence of the RNA not being crosslinked), and serum can contain RNase although RNA compatible blocking agents are commercially available . We founWhether glyoxal is a better or worse fixative than formaldehyde for microscopy studies of protein antigens remains a matter of dispute, and likely varies depending on precise cell type and target , 36. In https://www.babraham.ac.uk/our-research/epigenetics/jon-houseley/protocolsStep-by-step up-to-date protocols describing these methods are available from the JH group website at 2.COLO205 and MCF7 cell lines were provided by the laboratory of Dr. Simon J Cook at the Babraham Institute. All cell culture reagents were purchased from ThermoFisher Scientific, UK. Cells were cultured in RPMI-1640 (#21875) (COLO205) or Dulbecco’s modified Eagle’s medium (DMEM) (#41966) (MCF-7) supplemented with 10% (v/v) foetal bovine serum (#10270), penicillin (100 U mL−1), streptomycin (100 mg mL−1) (#15140) and 2 mM glutamine (#25030) in a humidified incubator at 37°C and 5% (v/v) COhttps://github.com/s-andrews/celllinesleuth)Cell line identity was validated based on RNA-seq data generated in this work using Cell Line Sleuth, developed by Simon Andrews of the Babraham Institute Bioinformatics Facility  and mixed until a homogeneous lysate was obtained. For phase separation, 0.2 mL chloroform was added to the lysate, mixed thoroughly and allowed to stand for 10 minutes prior to centrifugation at 12,000 × g for 15 min at 4°C. The upper aqueous phase containing RNA was transferred to a fresh tube, and 0.5 mL 2-propanol per mL original lysate was then added, samples mixed and allowed to stand for 10 minutes. Samples were centrifuged , and RNA pellets washed with 1 mL 75% (v/v) ethanol . Pellets were air-dried for 5–10 minutes and resuspended in RNase-free water.Sorted cells were pelleted by centrifugation at 2000 x 6 cells were lysed in a 350 μl cell lysis buffer and the lysate was mixed with equal amounts of 70% ethanol. The resulting mixture was applied to the RNeasy silica membrane and centrifuged (> 8000 x g for 15 seconds) to allow binding of RNA to the membrane. The membrane was washed twice by centrifugation (> 8000 x g for 15 seconds) with wash buffers to eliminate contaminants. RNA was eluted in 30μl nuclease-free water.RNA extraction from cells using Qiagen’s RNeasy extraction kit  was carried out according to the manufacturer’s instructions. Briefly, 1.5 x 10Total RNA (100 ng) from COLO205 cells was subjected to one-step RT-qPCR analysis using the Luna Universal One-Step RT-qPCR Kit  according to the manufacturer’s instructions. For each reaction, amplification was carried out in 20 μl reaction mixture containing 5 μl RNA sample (20 ng/μl), 10 μl Luna Universal One-Step Reaction Mix (1x), 1 μl Luna WarmStart RT Enzyme Mix (1x), 0.8 μl each of forward and reverse primers (0.4 μM) and 2.4 μl nuclease-free water. The PCR cycling conditions were reverse transcription at 55°C for 10 minutes, initial denaturation at 95°C for 1 minute, followed by 40 cycles of denaturation at 95°C for 10 seconds and extension at 60°C for 30 seconds on the Bio-Rad CFX-96 qPCR machine . Data were analysed with CFX-manager software v.3.1. Details of primer sequences are provided in Filter tips and RNase-free tubes were used throughout this project. Certified RNase-free water (Sigma W3513) was used for elution and storage of RNA. Bulk solutions  were assembled from standard laboratory chemicals and deionised or milliQ water without further purification. DEPC treatment was not used at any point. Gel electrophoresis was performed in a dedicated mini-gel tank although we consider this optional.This is a very simple and reliable RNA gel electrophoresis protocol, a minor adjustment of the method originally described here :Assemble 10x BPTE buffer: 30 g PIPES free acid (Melford P40140), 60 g Bis-Tris (Melford B75000), 20 ml 0.5 M EDTA pH8, deionised water to 1L, stir or shake to dissolve. Store at room temperature >1 year, dilute to 1x with deionised water before use.Assemble RNA denaturation mix: 6 ml DMSO (Sigma 34869), 2 ml 40% glyoxal solution , 1.2 ml 10x BPTE, 0.6 ml 80% glycerol. Split in 1 ml aliquots, keep a working aliquot at -20°C that can be used many times, store remaining aliquots at -70°C.Obtain ethidium bromide solution , ensure this is less than 2 years old. Ideally keep frozen aliquots for RNA gels. Problems with migration, particularly the molecular weight marker, are most frequently due to aged ethidium solutions in our experience.Add ethidium bromide solution at a final concentration of 50 μg/ml to sufficient RNA denaturation mix for 15 μl per sample including DNA molecular weight marker, vortex well (solution should be pink).Add up to 3 μl RNA to 15 μl RNA denaturation mix containing ethidium bromide and incubate 1–2 hours at 55°C. Treat the molecular weight marker in the same manner.Cast a 1.2% agarose gel in 1x BPTE, and assemble apparatus with 1x BPTE as the running buffer.Remove samples from heating block and load onto gel within 15 minutes.Run gel at 90 V on a standard mini-gel system for 30 minutes to 1 hour before imaging.Glyoxal was purchased as a 40% aqueous solution from Sigma (50649), stored at 4°C, checked for precipitation before use and if necessary warmed to 55°C until precipitate re-dissolved. This solution was not deionised at any point. A 3% glyoxal solution at pH 4–5 was used in all experiments as described . The solTrypsinise cells and collect the cell pellet in a 1.5 ml tube.Wash cells once with 1 ml PBS and remove the supernatant.Gently re-suspend cells in 100 μl of 3% Glyoxal fixation solution with 1:25 RNasin Plus (Promega N261B) and incubate for 15 minutes on ice.g for 3 minutes at 4°C. Discard the supernatant.Add 1 ml of 1xPBS with 1:100 RNasin Plus and centrifuge at 2000 x Gently re-suspend cells in 100 μl of ice-cold methanol (100%) with 1:25 RNasin Plus slowly (drop by drop while gently vortexing cells) to cells and incubate on ice for 30 minutes.g for 3 minutes at 4°C. Discard the supernatant.Add 1 ml of 1% BSA in PBS with 1:100 RNasin Plus and centrifuge at 2000 x Gently re-suspend cells in 100 μl of diluted antibody  in 1% BSA in PBS with 1:25 RNasin Plus and incubate for 1 hour on ice.g for 3 minutes at 4°C. Discard the supernatant.Add 1ml of 1% BSA in PBS with 1:100 RNasin Plus and centrifuge at 2000 x Re-suspend cells in 100 μl of diluted secondary antibody  in 1% BSA in PBS with 1:25 RNasin Plus. Incubate on ice for 30 minutes in the dark.g for 3 minutes at 4°C. Discard the supernatant.Add 1 ml of 1% BSA in PBS with 1:100 RNasin Plus and centrifuge at 2000 x Re-suspend the cell pellet in 200 μl of 1% BSA in PBS with 1:100 RNasin Plus.Prior to sorting, all samples were filtered to eliminate cell aggregates by passage through a sterile, 30 μm CELLTrics filter  into 5 mL polypropylene round-bottom tubes . Samples were then labelled with 4,6-diamidino-2-phenylindole (DAPI)  at a final concentration of 1 μg/mL in PBS. Cells were sorted using a 100 μm nozzle (at 20PSI) on a BD FACSARIA III cell sorter  with cooling of the sample and collection chambers enabled. Cells were collected in 1.5 mL RNase-free tubes pre-coated with 1% BSA in PBS supplemented with 1:25 RNasin Plus RNase inhibitor. The Alexa488 was excited by a 488nm laser and 530/30 bandpass filter used for collection of fluorescence. The DAPI was excited by the 355nm laser and its emission collected using a 450/50 filter.RNA prepared from flow sorted cells was quality controlled on a Bioanalyzer 2100 (Agilent) using RNA Pico 6000 chips (Agilent 5067–1513). RNA-seq libraries were prepared using the NEBNext Ultra II Directional RNA Library Prep Kit for Illumina (NEB E7760S) with the NEBNext Poly(A) mRNA Magnetic Isolation Module (NEB E7490S), following the protocol provided by the manufacturer with the exception that two successive 0.9x AMPure purifications were performed on the final amplified libraries. All libraries were amplified using 12 PCR cycles. Library quality was assessed on a Bioanalyzer 2100 using High Sensitivity DNA chips (Agilent 5067–4626) and quantification performed with a KAPA Library Quantification Kit . Libraries were sequenced by the Babraham Institute Next Generation Sequencing Facility on an Illumina HiSeq2500 in Rapid Run 50bp Single End mode.https://www.bioinformatics.babraham.ac.uk/projects/seqmonk/) and antisense reads mapping to each annotated gene were quantified and normalised to the total read count in each library. Scatterplots and hierarchical clustering were generated using SeqMonk, and analysis of differential gene expression performed using DEseq2 via SeqMonk Reviewers' comments:Reviewer's Responses to QuestionsComments to the Author1. Is the manuscript technically sound, and do the data support the conclusions?The manuscript must describe a technically sound piece of scientific research with data that supports the conclusions. Experiments must have been conducted rigorously, with appropriate controls, replication, and sample sizes. The conclusions must be drawn appropriately based on the data presented. Reviewer #1: YesReviewer #2: Yes**********2. Has the statistical analysis been performed appropriately and rigorously? Reviewer #1: YesReviewer #2: Yes**********3. Have the authors made all data underlying the findings in their manuscript fully available?PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.The Reviewer #1: YesReviewer #2: Yes**********4. Is the manuscript presented in an intelligible fashion and written in standard English?PLOS ONE does not copyedit accepted manuscripts, so the language in submitted articles must be clear, correct, and unambiguous. Any typographical or grammatical errors should be corrected at revision, so please note any specific errors here.Reviewer #1: YesReviewer #2: Yes**********5. Review Comments to the AuthorPlease use the space provided to explain your answers to the questions above. You may also include additional comments for the author, including concerns about dual publication, research ethics, or publication ethics. Reviewer #1: The authors present here a convincing protocol for transcriptome analysis from cells, performed after fixation and antigen staining.The main point of the manuscript is to allow analyses that could not be performed using conventional formaldehyde fixation. This point is thoroughly presented by the present manuscript, but one aspect, the poor performance observed in formaldehyde-fixed samples, should be enhanced further. I suggest that the authors add the "formaldehyde" condition to their RT-PCR analysis in Figure 2E. This should make this point more convincing, and will therefore improve the manuscript.Reviewer #2: This manuscript reports a detailed analysis for the use of the fixative Glyoxal for RNA studies. The authors provide first a thorough assessment of the impact of Glyoxal on RNA integrity alongside other fixative agents . They then use their protocol to performed RNA sequencing on FACS sorted cells stained for a cell-cycle marker . This demonstrates that glyoxal introduced only minimal, if any, biases to the transcriptome of treated cells while leading to good quality data.This is a nice and thorough report that will be of very useful to the community. I particularly thank the authors for providing a detailed step by step protocol on their website. I only have a handful of minor comments which are listed below.1) P. 4: “and library yields obtained were no more variable than we would normally expect between samples”. What is the expected and observed variability?2) P. 4: “identified 2 and 61 differentially expressed genes”. If not already done please report the genes in a supplementary table. If table exists please cite it.3) P. 5: “An equivalent experiment performed with COLO205 cells will be described elsewhere.”. I would recommend to either show the data or delete this statement.4) Glyoxal is reacting with RNA bases. Could the authors please comment on whether this creates mutations that may affect down-stream read-mapping or SNP calling?**********what does this mean?). If published, this will include your full peer review and any attached files.6. PLOS authors have the option to publish the peer review history of their article  digital diagnostic tool,  10 Dec 2020Response to Reviewer #1 The authors present here a convincing protocol for transcriptome analysis from cells, performed after fixation and antigen staining. The main point of the manuscript is to allow analyses that could not be performed using conventional formaldehyde fixation. This point is thoroughly presented by the present manuscript, but one aspect, the poor performance observed in formaldehyde-fixed samples, should be enhanced further. I suggest that the authors add the "formaldehyde" condition to their RT-PCR analysis in Figure 2E. This should make this point more convincing, and will therefore improve the manuscript.We thank the reviewer for this suggestion and we have now added data in Figure 2E in order to demonstrate the poor performance of formaldehyde-fixed RNA in PCR amplification reactions. We had included this control in the original experiment but did not present the results, hence the data for the other two conditions remains the same and we have simply added the formaldehyde condition. Because the formaldehyde-fixed samples yielded almost no RNA as measured by a nanodrop, a volume of RNA sample equivalent to the control unfixed sample (from the same number of cells) was used for each RT-PCR reaction. The remarkable reduction in signals from formaldehyde-fixed samples compared to untreated and glyoxal-fixed samples is evident from the data. We have added text to include this information and corrected the figure legend for clarity.Response to Reviewer #2 1) P. 4: “and library yields obtained were no more variable than we would normally expect between samples”. What is the expected and observed variability?We find that yields with these RNAseq library preparation kits involving poly(A)+ enrichment are rather variable (over a 10-fold range or so), probably because they involve so many processing and purification steps. Furthermore yields vary substantially with cell type, user and batch of reagents. We would therefore be hesitant to define an expected range. However, we take the reviewer’s point that our phrasing suggests we have such a range, so we have re-phrased to simply quote the observed variability:“library yields per cell type varied by <8-fold (3 amplification cycles)”2) P. 4: “identified 2 and 61 differentially expressed genes”. If not already done please report the genes in a supplementary table. If table exists please cite it.We have added this information in supplementary Table S1 and cited in the text. 3) P. 5: “An equivalent experiment performed with COLO205 cells will be described elsewhere.” I would recommend to either show the data or delete this statement.We thank the reviewer for this recommendation. We included this statement as a manuscript describing this data is in preparation and we hoped to be able to cite the preprint. However, the preprint will not now be submitted in time and therefore we have deleted this statement 4) Glyoxal is reacting with RNA bases. Could the authors please comment on whether this creates mutations that may affect down-stream read-mapping or SNP calling?This is a good point. As far as is known, the guanine-glyoxal adduct should revert to guanine; certainly the long history of using glyoxal as a denaturant in northern blotting means there cannot be a major level of mutation as that would impair hybridisation. We have looked at the mapping efficiencies in our various libraries; they are slightly reduced in the glyoxal fixed samples, but we think this is more likely due to the slight decrease in RNA quality caused by staining . Nonetheless, we have only validated this method for calling RNA levels and would be wary of applying it for SNP calling without further validation. We have therefore added the following to the manuscript:“Residual adducts would be expected to decrease the fidelity of reverse transcription, and resulting mutations to impair mapping. Indeed mapping efficiencies were slightly decreased in libraries prepared from fixed and stained samples, but the effect was small (unique mapped reads constituted 81±2% reads in libraries from unprocessed samples versus 75±5% in libraries made from fixed and stained samples). We suggest that this reduction stems from the slight decrease in RNA quality in stained samples, but further validation would be required before confidently calling SNPs from glyoxal-fixed RNA.”AttachmentRebuttal letter PLOS ONE.docxSubmitted filename: Click here for additional data file. 5 Jan 2021Glyoxal fixation facilitates transcriptome analysis after antigen staining and cell sorting by flow cytometryPONE-D-20-30764R1Dear Jon,We’re pleased to inform you that your manuscript has been judged scientifically suitable for publication and will be formally accepted for publication once it meets all outstanding technical requirements.Within one week, you’ll receive an e-mail detailing the required amendments. When these have been addressed, you’ll receive a formal acceptance letter and your manuscript will be scheduled for publication.http://www.editorialmanager.com/pone/, click the 'Update My Information' link at the top of the page, and double check that your user information is up-to-date. If you have any billing related questions, please contact our Author Billing department directly at authorbilling@plos.org.An invoice for payment will follow shortly after the formal acceptance. To ensure an efficient process, please log into Editorial Manager at onepress@plos.org.If your institution or institutions have a press office, please notify them about your upcoming paper to help maximize its impact. If they’ll be preparing press materials, please inform our press team as soon as possible -- no later than 48 hours after receiving the formal acceptance. Your manuscript will remain under strict press embargo until 2 pm Eastern Time on the date of publication. For more information, please contact Kind regards,Juan Mata, Ph.D.Academic EditorPLOS ONEAdditional Editor Comments :Reviewers' comments: 11 Jan 2021PONE-D-20-30764R1 Glyoxal fixation facilitates transcriptome analysis after antigen staining and cell sorting by flow cytometry Dear Dr. Houseley:I'm pleased to inform you that your manuscript has been deemed suitable for publication in PLOS ONE. Congratulations! Your manuscript is now with our production department. onepress@plos.org.If your institution or institutions have a press office, please let them know about your upcoming paper now to help maximize its impact. If they'll be preparing press materials, please inform our press team within the next 48 hours. Your manuscript will remain under strict press embargo until 2 pm Eastern Time on the date of publication. For more information please contact plosone@plos.org. If we can help with anything else, please email us at Thank you for submitting your work to PLOS ONE and supporting open access. Kind regards, PLOS ONE Editorial Office Staffon behalf ofDr. Juan Mata Academic EditorPLOS ONE


S. cerevisiae PPR protein essential for mitochondrial gene expression and involved in the stability of 15S ribosomal RNA. We demonstrate that in vitro Dmr1p specifically binds a motif composed of multiple AUA repeats occurring twice in the 15S rRNA sequence as the minimal 14 nt (AUA)4AU or longer (AUA)7 variant. Short RNA fragments containing this motif are protected by Dmr1p from exoribonucleolytic activity in vitro. Presence of the identified motif in mtDNA of different yeast species correlates with the compatibility between their Dmr1p orthologs and S. cerevisiae mtDNA. RNA recognition by Dmr1p is likely based on a rudimentary form of a PPR code specifying U at every third position, and depends on other factors, like RNA structure.PPR proteins are a diverse family of RNA binding factors found in all Eukaryotic lineages. They perform multiple functions in the expression of organellar genes, mostly on the post-transcriptional level. PPR proteins are also significant determinants of evolutionary nucleo-organellar compatibility. Plant PPR proteins recognize their RNA substrates using a simple modular code. No target sequences recognized by animal or yeast PPR proteins were identified prior to the present study, making it impossible to assess whether this plant PPR code is conserved in other organisms. Dmr1p  is a  The current widely accepted hypothesis explaining the evolutionary origin of the complex internal structure of Eukaryotic cells points to an endosymbiotic origin of mitochondria and chloroplasts, that still possess vestigial genomes of eubacterial origin . OrganelPentatricopeptide (PPR) proteins are a prominent family of mostly organellar RNA-binding proteins, common to all eukaryotes, and involved in various aspects of RNA metabolism . The PPRSequence-specific binding of RNAs is postulated for all the PPR proteins, but exact target sequences and mechanisms of substrate recognition were identified only for several plant members of the family . BioinfoTrypanosoma brucei, two PPR proteins were found to bind poly(A) and poly(G) stretches in mitochondrial RNAs  gene is the mitochondrial small subunit rRNA (15S rRNA)  respiratory deficiency (COB and COX1 transcripts (hence the name CCM1)  . Deletio15S rRNA , followe15S rRNA . Replacificiency . Paralleme CCM1) , 2012. Sme CCM1) . The obs15S rRNA , furtherIn this study we performed detailed in vitro studies of the interaction between Dmr1p and 15S rRNA, identifying a distinct motif of at least 14 nucleotides, composed of repeats of the AUA triplet, occurring in two regions of the 15S rRNA sequence, as the proposed recognition site.Previous work indicated that Dmr1p binds the 15S rRNA, consistent with its role in ensuring this molecule's stability inferred from genetic studies . This bi6 protein . Hybridization signal from each probe was quantified and the ratio of bound/unbound RNA complementary to each of the nucleotides calculated A,B. The 4AU motif, whereas the fragment recognized by probe #20 contains a longer 21-nt stretch of (AUA)7. Even though the entire 15S rRNA sequence is AU-rich, there are no other occurrences of more than four AUA repeats in other regions of the molecule. The fragment complementary to probe #7, that also showed some weak binding, does not contain an uninterrupted (AUA)n tract. There is, however, a somewhat similar sequence (AUA)2UGGGUAAUA, and this fragment was included in further experiments.Analysis of RNA sequences complementary to the probes #15 and 20#, that exhibited strong binding to Dmr1p in vitro, revealed that they share identical repetitive stretches comprising several repeats of the AUA trinucleotide C. The fr4AU motif occurs only twice in the mature 15S rRNA sequence, at positions 457 , and 761 (#15). Additionally, it can be found in the 5′ extension of the 15S rRNA primary transcript which is removed by processing .The (AUA)ocessing , at posiocessing . With th6 tag. The MBP-His6 protein was cross-linked to fragmented radiolabeled 15S rRNA, and the ratio of bound/unbound RNA fragments was calculated using the same method as described above for the Dmr1-MBP-His6 protein. Some enrichment of the AU-rich fragments was observed, but the bound/unbound ratio was always much lower than for the Dmr1-MBP-His6 protein .Copurification of bacterial RNA-binding factors, most notably Hfq, with heterologous proteins containing the hexahistidine tag and purified by metal-affinity chromatography is a known source of false positive RNA binding activities . As an a6 tag was removed from the heterologously expressed Dmr1-MBP-His6 protein using TEV protease, followed by size-exclusion chromatography to obtain highly purified Dmr1p. Radiolabeled RNA substrates were obtained by T7 in vitro transcription on synthetic oligonucleotide templates, and purified by polyacrylamide gel electrophoresis.In order to confirm that the identified AUA trinucleotide repeats constitute the specific Dmr1p binding site, we performed a series of electrophoretic mobility shift assays (EMSA). For these experiments the MBP-HisThe first series of experiments, using long RNA substrates corresponding to the probes used in the coprecipitation experiment, confirmed the interaction of Dmr1p with fragments complementary to probes #15 and #20 and a very weak interaction with the fragment complementary to probe #7. The fragments corresponding to probes #6, #11, #17, #19, and #24 showed very weak or no appreciable interaction with Dmr1p A. At the7 stretch (found in fragment #20) strongly binds Dmr1p in vitro, and the shorter (AUA)4AU motif is still sufficient to ensure reliable binding. The substrate containing the (AUA)2UGGGUAAUA sequence found in fragment #7 showed only minimal binding, and the negative control did not exhibit any mobility shift indicative of protein binding. In the assay using shorter, 14 nt substrates, the (AUA)4AU sequence (common for fragments #15 and #20) showed clear binding 5AUAU was not bound at all. In the experiment using 24 nt probes, and to a lesser extent also the 14 nt probes, even unbound RNAs migrate in the nondenaturing gel as multiple bands, indicating that they easily form different secondary structures which may be relevant for their interactions with Dmr1p.As these preliminary results indicated visible background binding to AU-rich sequences in addition to clear interaction with the regions identified in the coprecipitation experiment, we designed shorter (24 nt and 14 nt) RNA substrates comprising the putative Dmr1-binding motif of fragment #15 and #20, as well as the weakly bound fragment #7. Unrelated, but similarly AU-rich fragments were used as negative controls. The results obtained with longer, 24 nt RNA transcripts B clearly binding C, while The EMSA assay results have thus confirmed that the sequence containing four AUA repeats  is specifically recognized by the Dmr1 protein, and that the addition of more repeats (to seven) seems to increase the strength of interaction. Binding to a sequence containing fewer AUA repeats cannot be excluded, but is markedly weaker, whereas other AU-rich sequences do not interact with Dmr1p sufficiently to cause a detectable electrophoretic mobility shift.DMR1 gene results in fragmentation of the 15S rRNA . The substrate containing the longest (AUA)7 binding site (UU(AUA)7U) was completely protected from PNPase degradation by bound Dmr1p, whereas fragments containing the core 14 nt (AUA)4AU sequence, and the possibly weakly interacting (AUA)2UGGGUAAUA, as well as the negative control with the distinct, but similarly AU-rich sequence were not protected and efficiently degraded by PNPase sequence A, thus e repeats A. We hav results C indicat4AU sequence can protect it from degradation from the 5′ end, where only two additional nucleotides can be found upstream of the putative binding site. The transcript was treated with 5′ polyphosphatase to generate an RNA fragment monophosphorylated at the 5′ end. Incubation of this substrate with a commercially available 5′–3′ exoribonuclease (Terminator) indicates that partial but evident protection can be achieved achieved D.Even though some unspecific background RNase activity  in the Dmr1 protein preparation is apparent, both experiments confirm that binding of Dmr1 to the recognized motif is specific and strong enough to protect from exoribonucleolytic activity from either end.4AU binding sequence with additional 12 nt downstream and 9 nt upstream was incubated with Dmr1p and a recombinant yeast processive hydrolytic 3′–5′ exoribonuclease Dis3p (Rrp44p)  . In the  product , and furS. cerevisiae and S. bayanus, which likely arose upon the divergence between S. bayanus and the common ancestor of S. cerevisiae, S. paradoxus, S. mikatae, and S. kudriavzevii  is preserved, whereas in S. castellii, Candida glabrata, and all the species that diverged prior to the Whole Genome Duplication (WGD) this motif cannot be found anywhere in the 15S rRNA sequence. Remarkably, Dmr1p from S. mikatae and S. kudriavzevii, as well as from S. paradoxus, can function in the context of S. cerevisiae mtDNA 4AU motifs in the 15S rRNA sequence failed to support any perceptible respiratory growth, signifying a complete lack of compatibility with S. cerevisiae mtDNA.In order to provide experimental confirmation of these findings, we replaced the perature B. The or reports . Partialer group . DMR1 orS. cerevisiae 15S rRNA sequence in at least two distinct regions, are conserved in the Saccharomyces species that show at least partial compatibility of Dmr1p orthologs with S. cerevisiae, and are absent from the species where the Dmr1p ortholog is completely incompatible with S. cerevisiae mtDNA. The regions containing the (AUA)4AU motif are the only ones that fulfil these criteria .To further strengthen these results, we performed a reverse in silico analysis, searching for sequence motifs that are at least 14 nt long, occur in the 4AU motifs in their 15S rRNA completely fail to function in the context of S. cerevisiae mtDNA, while those originating from species where at least one such motif can be found will at least partially complement the phenotype of a S. cerevisiae Δdmr1 strain (S. paradoxus Dmr1p (one motif) is more compatible with S. cerevisiae 15S rRNA than S. bayanus Dmr1p , suggesting that other factors can influence the strength of Dmr1p–15S rRNA interaction.Overall, Dmr1p orthologs from species that do not have any occurrences of the identified (AUA)1 strain . This fuYeast and animal PPR motifs are more divergent than the ones found in plants , and theStudies correlating the sequences of individual PPR motifs with their target RNA sequences in plants revealed the existence of a simple combinatorial code wherein two amino acids in two adjacent α-helical segments of a PPR motif determine the identity of a single nucleotide in the recognized sequence . AttemptCOB mRNA was implicated in the interaction with the Cbp1 protein through suppressor analysis  and poly(A), respectively 7. Both regions correspond to regions near the surface of mitoribosome 2A, five (AUA)2A, and eight (AUA)2), yet fragments containing them failed to yield significant enrichment in the RNA pull-down experiment, suggesting that the 14-nt (AUA)4AU motif is close to being the minimal target sequence required for Dmr1p binding.The 15S rRNA was previously identified as the primary, and likely sole target of yeast Dmr1p through a combination of genetic, molecular and evolutionary studies . By coprribosome . Prefere4AU motif or the longer (AUA)7 sequence were protected from exoribonucleolytic activity by incubation with Dmr1p. The shorter (AUA)4AU motif did not provide protection from degradation by the 3′–5′ exoribonuclease PNPase when the substrate contained eight additional nucleotides downstream, suggesting that the strong processive activity of PNPase is sufficient to displace bound Dmr1p from this RNA. Binding of Dmr1p to a similar substrate was, however, sufficient to arrest the degradation by another 3′–5′ exoribonuclease—Dss1p, at a site consistent with the occupation of the (AUA)4AU motif by the bound Dmr1p. Protecting the 15S rRNA from degradation by ribonucleases was proposed to be the primary function of Dmr1 protein, based on the distinct degradation pattern observed in the deletant deletant . Cautiondeletant , normallactivity . The proDMR1 results in the fragmentation of 15S rRNA  with concomitant disturbance of translation and RNA processing. Based on the observed presence of Dmr1p within the large ribosome-bound complex  corresponds to a single nucleotide in the RNA . As the As the sequence recognized by Dmr1 consists of trinucleotide AUA repeats, we turned to structural predictions obtained via the I-TASSER server, looking for regularities in the motif sequences. We compared sequence profiles in every third motif in three possible phases . Two of 1 N6 pattern. However, the longer of the recognized motifs reaches the length of exactly 21 nt. The general rule of one PPR motif recognizing a single RNA nucleotide appears thus to be valid for the interaction of Dmr1p with its target sequence, yet, not all of the predicted pairs of α-helices conform to the current HMM profiles of yeast PPR motifs  temperature, with concomitant decrease in the mature 15S rRNA level 4AU motif in the 15S rRNA sequence. In all the species where a complete  or partial (S. bayanus) compatibility is preserved, at least one such motif can be found, while it is absent from the 15S rRNA sequences in the more distant species, where no compatibility is retained. Remarkably, in silico analysis shows that (AUA)4AU is the only 14 nt motif in the 15S rRNA sequences of the analyzed yeast species that shows such phylogenetic distribution, further strengthening the evidence for its key role in the recognition of substrate by the Dmr1 protein. The number of the (AUA)4AU motifs is not, however, strictly correlated with the degree of compatibility, as only one such motif can be found in S. paradoxus, and in S. mikatae that show complete compatibility, as well as in S. bayanus, where the compatibility is only partial.One of the most fascinating aspects of PPR protein biology is their role in the evolution of nucleo-organellar compatibility. In plants, one of their postulated roles is maintaining the function of organellar genes by compensating for mutational pressure, known as “genome debugging” . In yeasA level  determines affinity either for guanine or uridine, in cooperation with amino acid in position 6 of a preceding PPR motif. On the other hand asparagine 6 (N6) indicates an affinity for pyrimidine (uridine or cytosine) . In plants aspartate in this position (Dytosine) . In our 6 fusion protein was preformed essentially as described previously  sequence of DMR1 ORF  was obtained by custom gene synthesis (Generay Biotech Co.) and cloned in the pET28a vector (Novagen). The pET28a::DMR1 vector was transformed into BL21(DE3) CodonPlus-RIL E. coli (Agilent). An amount of 1 mL of the overnight culture was inoculated into 500 mL of AIM , and the cells were grown at 23°C for 36 h. Metal affinity and size exclusion protein purification procedures were performed as described previously , and Phusion DNA polymerase (Thermo Fisher), and cloned into pTZ19R (Thermo Fisher). The pTZ19::15S vector was linearized with SmaI (Thermo Fisher) and transcribed using the T7 transcription kit (Thermo Fisher), according to the manufacturer's instructions, with α-32P-UTP  added to the reaction to produce radiolabeled substrate, followed by digestion with RNase-free DNase I (Roche). Templates for shorter RNAs were generated by annealing appropriate complementary synthetic DNA oligonucleotides , containing the T7 promoter sequence, and transcribed in vitro as described above. The RNA transcripts were then purified by electrophoresis in denaturing acrylamide gels  and isolation of the appropriate bands as described previously  with double amounts of all the reagents. The product was precipitated with EtOH and resuspended in 200 µL RNase free water. The specific activity of this preparation was about 6000 cps/µL. Radioactive RNA was fragmented with NEBNext Magnesium RNA Fragmentation Module (#E6150S) for 7 min. The reaction was stopped by addition of Tris-HCl to 300 mM. Fragmentation of RNA was confirmed by agarose gel electrophoresis.One third of the fragmented RNA (80 µL) was incubated with 60 µg of the purified protein and 200 ng of unlabeled 15S rRNA competitor in 150 µL total volume of binding buffer  for 20 min. on ice. Cross-links were induced by UV irradiation using the Hoefer UVC500 device (2′20″ at 3.5 cm from the lamp in an Eppendorf tube). The cross-linked sample was then mixed with 500 µL of buffer A  and 80 µL of HisPur cobalt resin (Thermo Fisher) equilibrated with buffer A, and incubated on a rotary shaker at 6°C for 1 h. The supernatant was then collected and saved. The resin was washed three times with 400 µL of buffer A (decrease of radioactivity in each subsequent wash was monitored using a Geiger counter), and all the washes were pooled with the original supernatant to yield the unbound fraction. An amount of 400 µL of elution buffer (buffer A + 250 mM imidazole) was then added to the resin and incubated on a rotary shaker for 15 min. The eluate constituted the bound fraction (its radioactivity was confirmed to be at least five times that of the last wash). An amount of 10 µL (>6 U) of proteinase K (Thermo Fisher) and SDS to 0.5% were added to both the bound and unbound fraction preparations, and incubated at 37°C for 30 min. Bound and unbound fraction RNA was then purified by phenol:chlorophorm extraction, followed by EtOH precipitation.Supplemental Table S1), with each probe sequence overlapping the next by 20 nt. An unrelated oligonucleotide, complementary to the araA gene from Aspergillus nidulans was used as the negative control (N). The array was prepared on Nytran N nylon membrane (Whatman) using the Bio-Dot SF (BioRad) device. 5× SSPE buffer was used to preincubate the membrane and prewash the wells. An amount of 5 µg of each oligonucleotide in 200 µL of 5× SSPE was passed through each well using a vacuum, followed by washing with 500 µL of 5× SSPE. The membrane was then dried, and the oligonucleotides were UV-fixed to the membrane using the Hoefer UVC500 device at 1200 µJ/cm2. The membrane was prehybridized for 6–8 h at 42°C in PerfectHyb Plus (Sigma Aldrich). Radioactive RNA samples (bound or unbound) were added to the membrane in 1 mL of PerfectHyb Plus and hybridized overnight at 40°C. The membranes were then washed twice in 2× SSC, 0.1% SDS at room temperature, followed by 0.2× SSC, 0.1% SDS at room temperature and 0.2× SSC, 0.1% SDS at 37°C. Autoradiography of the membrane was performed for at least 48 h on a phosphor screen (Fuji). Typhoon FLA 9000 laser imaging system  device was used to quantitate the signal from each slot, and the ratio of bound/unbound fraction signal for each probe was then calculated. The experiment was performed in triplicate.The slot-blot macro array was prepared using 27 synthetic oligonucleotides, 80 nt each, covering the entire sequence of mature 15S rRNA  in 14 µL of reaction buffer  with 1 U/μL of RiboLock RNase inhibitor (Thermo Fisher) for 20 min. on ice. 0.5 U of Terminator 5′-Phosphate-Dependent Exonuclease (Epicentre) was then added, and the reaction incubated for 30 min at 30°C.Gel-purified radiolabeled RNA fragments obtained using in vitro transcription were treated with RNA 5′ Polyphosphatase (Epicentre) according to the manufacturer's protocol, purified by phenol:chlorophorm extraction and EtOH precipitation, and resuspended in RNase free water. Radiolabeled RNA (30–40 cps) was incubated with 1.2 µg of Dmr1-His6 fusion protein purified by metal affinity and size exclusion or with 1.2 µg of BSA (negative control) in 14 µL of reaction buffer  with 1 U/μL of RiboLock RNase inhibitor (Thermo Fisher) for 20 min on ice. An amount of 0.2 µg (>0.1 U) U of recombinant Synechocystis sp. PNPase (Sigma) was then added, and the reaction incubated for 30 min at 25°C.Gel-purified radiolabeled RNA fragments obtained using in vitro transcription (30–40 cps) were incubated with 1.2 µg of Dmr1-His6 fusion protein purified by metal affinity and size exclusion or with 0.8 µg of BSA (negative control) in 14 µL of reaction buffer  with 1 U/μL of RiboLock RNase inhibitor (Thermo Fisher) for 20 min on ice. 0.3 µg of purified Dis3p (a gift from Dr. Rafał Tomecki) was then added, and the reaction incubated for 30 min at 30°C.Gel-purified radiolabeled RNA fragments obtained using in vitro transcription (30–40 cps) were incubated with 0.8 µg of Dmr1-HisIn each assay the reaction was stopped by addition of the same volume of the electrophoresis loading buffer , and the products were separated in a 20% denaturing polyacrylamide gel.S. cerevisiae DMR1 gene in the YCplac111  vector was described previously . The nonstandard CTG (serine) codon in the D. hansenii gene was changed to the standard TCG (serine) using site-directed mutagenesis following the Quikchange Site Directed Mutagenesis (Agilent) PCR protocol. All the constructs were verified by Sanger sequencing performed in the Laboratory of DNA Sequencing PAS.Standard yeast genetic media and methods were as described previously . Yeasts eviously . The DMRsion PCR   vector  was described previously  and YPG (glycerol) plates and incubated at 30°C for 3 d. Empty YCplac111 vector and the vector carrying the S. cerevisiae DMR1 sequence were used as negative and positive controls, respectively.Construction of the HS-DPPR strain with the eviously . Vectorselection . For tesS. cerevisiae: KP263414.1, S. paradoxus: AF114922.1, S. bayanus: AF114901.1, C. glabrata: AJ511533.1, K. lactis : AY654900.1, Y. lipolytica: AJ307410.1, D. hansenii: DQ508940.1, S. mikatae: KX657747.1, S. kudriavzevii: KX657746.1, S. castellii: AF437291.1, L. kluyveri: HE664112.1, K. waltii: AF442341.1, A. gossypii: AE016821.1, C. albicans: AF285261.1. The sequences and the Python scripts used to analyze them are available at https://github.com/golikp/dmr1_motif. The cladogram in The I-TASSER server  was usedSupplemental material is available for this article.


N6-methyladenosine (m6A), 2-O-dimethyladenosine (m6Am), N1-methyladenosine (m1A), 5-methylcytosine (m5C) and 7-methyl guanosine (m7G). Dynamic and reversible methylation on RNA is mediated by RNA modifying proteins called “writers” (methyltransferases) and “erasers” (demethylases). “Readers” (modified RNA binding proteins) recognize and bind to RNA methylation sites, which influence the splicing, stability or translation of modified RNAs. Herein, we summarize the biological functions and mechanisms of these well-known RNA methylations, especially focusing on the roles of m6A in gastrointestinal cancer development.RNA methylation has emerged as a fundamental process in epigenetic regulation. Accumulating evidences indicate that RNA methylation is essential for many biological functions, and its dysregulation is associated with human cancer progression, particularly in gastrointestinal cancers. RNA methylation has a variety of biological properties, including  Gastrointestinal (GI) cancers refer to malignant conditions of the GI tract and accessory organs of digestion including liver and pancreas. Globally, GI cancers account for roughly half of all cancer-related deaths , 2. GI cEven though modification events in RNA molecules were discovered in the 1950s, our understanding of RNA modification is limited –6. RecenAmong RNA modifications, RNA methylation is the most well-characterized type. Highly dynamic and reversible methylation on RNA is mediated by a number of proteins, which are called RNA-modifying proteins (RMPs) , 9. RMPsIn this review, we focus on metabolic regulatory mechanisms and biological functions of RNA methylation. Moreover, we summarize the role of dysregulated m6A modifying proteins in cancer initiation and progression, and discuss the potential of targeting m6A modifying proteins with aberrant expression for therapy.N6-methyladenine, N2-methylguanine and 1-methylguanine occurred in the ribonucleic acid hydrolysates in yeast [N6-methyladenosine (m6A), 2-O-dimethyladenosine (m6Am), N1-methyladenosine (m1A), 5-methylcytosine (m5C) and 7-methyl guanosine (m7G) -binding protein, is identified as a catalytic subunit in this complex, while METTL14 serves as an essential component to facilitate RNA binding [N(6)-hydroxymethyladenosine (hm6A) as an intermediate modification, and further oxidize hm6A to N(6)-formyladenosine [m6A is dynamically regulated by specific methyltransferases and demethylases. The most well-characterized methyltransferases is the core methyltransferase complex, which consists of methyltransferase-like 3 (METTL3) and methyltransferase-like 14 (METTL14) Fig. a. METTL3 binding . Recent  binding . Fat masdenosine , 28. Alkdenosine . Severaldenosine , 30, 31.O-methyladenosine (Am)  Fig. b. Then, m1A is a reversible methylation with the addition of a methyl group and a positive charge at the N1 position of adenosine, which blocks the Watson–Crick interface, alters RNA secondary structures and protein-RNA interactions  Fig. 2cc2c. The 1A methyltransferase responsible for cytoplasmic tRNAs has two subunits, TRMT6 and TRMT61A in eukaryote [N1-methyltransferase catalytic activity, and TRMT6 is important for tRNA binding [The best-characterized mukaryote   (CAA) precursors [Asp, tRNAGly and tRNAVal [− and 2-oxoglutarate-dependent dioxygenases to demethylate m5C sites in mammalian RNAs [m5C occurs in both DNA and RNA by introducing a methyl group in the 5th carbon atom of cytosine  Fig. 2dd2d. The ecursors , 59. The tRNAVal . The tenian RNAs . These Tian RNAs , 62.Most eukaryotic mRNAs have a methyl group and a positive charge at the N7 position of the terminal guanosine at its 5′ cap Fig. e. m7G isRecently, the rapid development of transcriptome-wide profiling of diverse RNA methylations, along with the identification of the corresponding writers, erasers and readers, enable researchers to reveal the regulatory roles of RNA methylation in RNA metabolism and a wide range of cellular and biological processes.RNA methylations appear to function through nearly all steps of RNA metabolism, including RNA capping, splicing, stability, translation, and nuclear exportation Fig. . The pro+ RNA retention in the nucleus [Nuclear export is an essential step for mRNA to ensure its translation in the cytoplasm. Knockdown of the m6A easer ALKBH5 induces poly (A) nucleus . The m5C nucleus . These siMet, suggesting that m1A modification is important for tRNA stability [The dynamic synthesis and degradation of mRNA maintains an acute amount of mRNA in different contexts. For example, METTL3 knockdown decreases the m6A modification of cytokine signaling 2 (SOCS2) mRNA, leading to its degradation . In additability . These dtability . NSUN2-mtability . Furthertability , 48.Mettl3 conditional knockout mice display the enhanced apoptosis of newborn cerebellar granule cells in the external granular layer, and lower levels of m6A modification in brain mRNA with aberrant RNA splicing and stability [Mettl3 in the mouse hippocampus exhibit no significant morphological change but reduced memory consolidation ability [deleted of Mettl3 or Mettl4 manifests a prolonged cell cycle [Considering the diverse regulation of RNA methylation on RNA fate, it would not be surprising that RNA methylation regulates many cellular and developmental processes. m6A modification has been documented to be important for nervous system development. tability . Postnat ability . Embryonll cycle . Additioll cycle .Ythdc2 knockout mice fail to undergo pachytene, leading to defects in spermatocyte development [m6A modification is also important for germline development in mice. The ablation of mouse METTL3 inhibits spermatogonial differentiation and blocks the initiation of meiosis in germ cells . The defelopment .Ime4 abolishes the m6A modification of the sex determination factor sex-lethal (Sxl), leading to X inactivation failure and improper sex determination [RNA methylation also influences other fundamental physiological processes in development. Deletion of drosophila MTEEL3 homolog mination , 77. Zebmination . Ythdf2-mination .As mentioned above, RNA methylation has been implicated in diverse biological processes. There is no doubt that its dysregulation is highly associated with tumor initiation and progression. Accumulating evidence has shown that dysregulation of RNA methylation in cancer development, and the most well-studied one is m6A. m6A involves in cancer progression often through the actions of writers and easers that alter the abundance of m6A in the mRNA of oncogenes or tumor-suppressive genes and then the readers that recognize the modified sites to further upregulate oncogene or downregulate tumor-suppressive gene expression.SRY-box transcription factor 2 (SOX2), which enhances the stability and protein expression of SOX2 mRNA [yes-associated protein mRNA [matrix metallopeptidase 2 [Many studies have shown that m6A modification plays a crucial role in various cancers. For example, the master methylase of m6A METTL3 has been reported to involve in many steps of RNA processing and the oncogenic pathway in different cancer types. In glioblastoma, METTL3 facilitates the maintenance and radiation resistance of glioma stem-like cells via increasing the m6A modification of OX2 mRNA . In non-ein mRNA . METTL3 tidase 2 . MeanwhiGastric cancer develops from the lining of the stomach. More than 950,000 patients are diagnosed as gastric cancer, and about 720,000 patients died from gastric cancer every year worldwide . It is uMETTL3 mRNA in the metastatic colorectal cancer tissue is positively associated with poor prognosis of patients [18F-FDG uptake is observed in colorectal cancer patients [XIST and attenuates its expression. YTHDF2 could recognized the m6A-methylated XIST to mediate its degradation [Colorectal cancer develops from one or a few small polyps inside the colon, and gradually develops into mature colorectal cancers. More than 1.8 million new colorectal cancer cases and 881,000 deaths are estimated to occur in 2018 [patients . Knockdopatients  also reppatients  found thpatients . Furtherpatients . Functiopatients . Mechanipatients . Knockdoradation . Additioradation . Furtherradation .METTL3, METTL14 expression in HCC tissues shows a mild increase, and promotes HCC cell proliferation, colony formation and migration in vitro [METTL14 enhances tumor metastasis. METTL14 was further showed to bind to the microprocessor protein DGCR8 mRNA and increase microRNA-126 in an m6A-dependent manner. WTAP also enhances the capability of HCC cell proliferation and tumor progression via the HuR-ETS1-p21/p27 axis [PKM2 mRNA and enhancing its translation efficiency. In contrast, FTO is found to be downregulated in intrahepatic cholangiocarcinoma (ICC). Loss of FTO in ICC is associated with tumor aggressiveness and poor prognosis. Suppression of FTO reduces the apoptosis of ICC cells and increases resistance to cisplatin treatment [Liver cancer is predicted to be the fourth leading cause of cancer death worldwide, with about 841,000 new cases and 782,000 deaths annually . METTL3 in vitro . In contin vitro  reportedp27 axis . Moverovp27 axis  showed tp27 axis . FTO facreatment . The expreatment . YTHDF2 KCNK15-AS1 [Pancreatic cancer is the seventh leading cause of cancer death, with over 1,000,000 new cases and 65,000 deaths estimated globally . METTL3 NK15-AS1 . In addiNK15-AS1 . SilenciO-methyltransferase inhibitor, as an inhibitor of FTO. Furthermore, citrate and IOX3 may be potential inhibitors of another m6A demethylase, ALKBH5, which also maintains tumorigenicity of several types of cancer cells [Given that m6A modifying proteins play important roles in so many malignant behaviors of cancer cell, they could be used as potential therapeutic targets for cancer treatment. Some FTO inhibitors have been developed by different research groups to increase the m6A abundance , 110. MAer cells , 116. Iner cells , ovarianer cells . Thus, rDuring the past 5 years, researchers have found that RNA modification are implicated in mRNA splicing, stability, translation, nuclear export and a wide range of biological processes. RNA modification and RNA-modifying proteins are often found to be aberrant expressed in tumor tissues. The changes of RNA modifications and their RNA-modifying proteins in cancer affects almost every aspect of RNA fate and is context-dependent. These RNA-modifying proteins linked to cancer are important regulators of cellular events by regulating RNA metabolism and expression of certain gene required for cancer cell proliferation, transformation, invasion, and other malignant behaviors. Studies on cancer epitranscriptomics focused on the complex context and heterogeneity of tumor could help us to better understand the exact roles of RNA modifications and their enzymes, which is benefit for exploring RNA modifying proteins as potential therapeutic targets.Even though emerging data show the potential roles of RNA methylation in gastrointestinal cancer development, some important questions still remain to be addressed. First, how does the same RNA modifying protein selectively catalyze the different target transcripts in different cancer types. This may be attributed to the different upstream regulator of RNA modifying protein upon the specific cellular context in different cancers. Second, the inconsistent findings of the same RNA modifying protein were reported in the same cancer. One possible interpretation is that our current hypothesis is only based on individual cohorts with a relatively limited number of cancer patients with different genetic background and environmental exposure. Third, although a number of new types of RNA methylation in human have been identified until now, their functions in GI cancers need to be further investigated in the future.


Although follicular thyroid carcinoma (FTC) generally has a good prognosis, it occasionally metastasises, leading to poor prognosis. Unfortunately, minimally invasive FTC (mi-FTC) and encapsulated angioinvasive FTC (ea-FTC) cannot be distinguished cytopathologically from thyroid follicular adenoma (FTA), a benign tumour with a good prognosis. Therefore, a molecular diagnosis to distinguish mi- or ea-FTC from FTA is needed for clinical treatment. Several transcriptomics/proteomics studies have searched for FTC biomarkers. However, the results of these studies were not consistent, which could be partly explained by inaccurate diagnosis of the specimens analysed.We conducted a microarray-based genome-wide transcriptome analysis using formalin-fixed paraffin-embedded mi- or ea-FTC specimens from patients who developed distant metastasis up to 10 years postoperatively, which ensured the accuracy of diagnosis. The recent 4th edition [Follicular thyroid carcinoma (FTC) and papillary thyroid carcinoma are major histological types of thyroid carcinoma. Until recently, FTC was subdivided into minimally invasive FTC (mi-FTC) and widely invasive FTC, based on the World Health Organization (WHO) classification  are transcribed from the human genome . InteresThe specimens used in this study were equivalent to ea-FTC or mi-FTC in postoperative pathological examination. To increase the accuracy of analysis, we selected patients with a diagnosis of metastatic mi/ea-FTC that had been established using the following criteria: i) routine thyroglobulin testing and neck ultrasonography for ≥ 10 years, and ii) recognition of distant metastasis after the initial operation. RNA was extracted from formalin-fixed paraffin-embedded (FFPE) specimens of the tumour and adjacent non-tumour tissues from patients with mi/ea-FTC. The RNA samples were then subjected to genome-wide transcriptome analysis, which enabled profiling of transcripts including ncRNAs.Table ® totalRNA FFPE XS . RNA concentrations were measured spectrophotometrically . The size distribution of total RNA was analysed using an Agilent 2100 Bioanalyzer  (Data file 8).In this study, specimens from three patients who underwent surgery at Kuma Hospital  were selected for analysis: one had diagnostic features of mi-FTC and two had features of ea-FTC. Data file 7 summarises the clinical findings of these patients. The primary surgical specimens were evaluated histopathologically in accordance with WHO criteria. This study was conducted in accordance with the principles of the 1975 Declaration of Helsinki; informed consent was obtained from each patient. Archival mi/ea-FTC FFPE samples were processed into 20-μm sections; regions containing carcinoma or non-carcinoma tissues in each section were isolated separately. Total RNA was extracted from these samples using NucleoSpin® WT Pico Reagent Kit (Thermo Fisher Scientific) with slight modification. Briefly, 50 ng of total RNA from each sample were reverse transcribed and subjected to polymerase chain reaction to synthesise T7 promoter-tagged double stranded cDNA. The cDNA was then subjected to in vitro transcription to synthesise complementary RNA; 20 ng of complementary RNA was reverse-transcribed using random primers for sense strand cDNA synthesis. After removal of template RNA using RNase H, 5.5 or 3.2 μg sense strand cDNA were digested with uracil-DNA glycosidase into 40–70-nt fragments. Successful fragmentation was confirmed using an Agilent 2100 Bioanalyzer (Agilent) (Data file 9). The fragmented cDNA was then biotin-labelled with terminal deoxynucleotidyl transferase and subjected to hybridisation performed in accordance with the GeneChip® WT Pico Reagent Kit manual. The Clariom D array was then processed through the automatic washing step using the GeneChip® Hybridisation, Wash, and Stain Kit (Thermo Fisher Scientific) and Fluidics Station 450 (Thermo Fisher Scientific). Hybridised targets on the array were stained with streptavidin–phycoerythrin provided in the kit and detected using Scanner 3000 7G (Thermo Fisher Scientific).An Affymetrix Clariom D Assay (Thermo Fisher Scientific), a next-generation microarray for transcriptome profiling, was used for genome-wide transcriptome analysis to increase the possibility of biomarker discovery. Labelled targets were prepared from RNA samples using the GeneChipRaw data (CEL files) were produced for the six samples using Affymetrix GeneChip Command Console Software and processed using Affymetrix Expression Console Software. The CEL files are registered under Gene Expression Omnibus (GEO) accession no. 701012. A detection call algorithm was used to filter and remove missing expression values based on absent/present calls. Using this algorithm, present, marginal, or absent calls were obtained for each probe set in each array. A scaling factor was applied to the normalised data from the CEL files to bring the average intensity for all probes on the array to 500, generating CHP files using Microarray Suite 5 software. To compare gene expression, data assigned to absent calls were omitted.Data file 10 shows scatterplots of the correlation of signal values between carcinoma and non-carcinoma samples. Normalised signal values for individual genes are listed in Data file 11.Herein, we provide information regarding a mi/ea-FTC microarray dataset, which will help to identify prognostic biomarkers for predicting high risk of metastasis among patients diagnosed with mi/ea-FTC, following the initial operation. Furthermore, such biomarkers may be used to diagnose FTC in thyroid specimens obtained by fine-needle aspiration biopsy for liquid-based cytology . However