Dynamic RNA-protein interactions govern the co-transcriptional packaging of RNA polymerase II (RNAPII)-derived transcripts. Yet, our current understanding of this process in vivo primarily stems from steady state analysis. To remedy this, we here conduct temporal-iCLIP (tiCLIP), combining RNAPII transcriptional synchronisation with UV cross-linking of RNA-protein complexes at serial timepoints. We apply tiCLIP to the RNA export adaptor, ALYREF; a component of the Nuclear Exosome Targeting (NEXT) complex, RBM7; and the nuclear cap binding complex (CBC). Regardless of function, all tested factors interact with nascent RNA as it exits RNAPII. Moreover, we demonstrate that the two transesterification steps of pre-mRNA splicing temporally separate ALYREF and RBM7 binding to splicing intermediates, and that exon-exon junction density drives RNA 5′end binding of ALYREF. Finally, we identify underappreciated steps in snoRNA 3′end processing performed by RBM7. Altogether, our data provide a temporal view of RNA-protein interactions during the early phases of transcription. Dynamic RNA-protein interactions govern the co-transcriptional packaging of RNA polymerase II derived transcripts. Here the authors use temporal-iCLIP which combines transcriptional synchronisation with UV cross-linking of RNA-protein complexes to reveal dynamic RNA-protein interactions during the early phases of transcription and beyond. This process initiates during transcription where early remodelling steps impact RNP formation5.The fate of an RNA polymerase II (RNAPII)-transcribed RNA is impacted by its dynamic association with RNA-binding proteins (RBPs). The early ribonucleoprotein (RNP) complex is acted upon by RNA processing-, transport- and decay-factors, which, dictated by transcript features, compete to shape RNP identity, while at the same time eliminate RNA processing by-products and mis- or excessively-produced material7G) cap6. CBP80 in turn initiates the binding of proteins capable of directing the fate of the elongating RNA9. After productive transcription is initiated, the CBC is joined by the Arsenite resistance protein 2 (ARS2) to form the CBC-ARS2 (CBCA) complex11, which then aids the recruitment of additional factors onto the nascent RNA  component ALYREF is being debated14. Adaptors of RNA decay factors are also loaded onto nascent transcripts to achieve RNA processing or complete degradation. One such adaptor is the trimeric Nuclear Exosome Targeting (NEXT) complex, which together with the ZC3H18 protein joins CBCA to form the CBC-NEXT (CBCN) complex7. Comprised of the MTR4, ZCCHC8 and RBM7 proteins, NEXT then recruits the ribonucleolytic RNA exosome to degrade a wide range of short non-adenylated transcripts and to process precursor forms of small RNAs, like snoRNAs, residing inside pre-mRNA introns19.An early and omnipresent member of RNAPII-derived RNPs is the Cap Binding Complex (CBC) composed of CBP20 and CBP80. CBP20 binds nascent RNA after the transcript 5′end has received its hallmark 7-methylguanylate  with the intron branchpoint (BP) and subsequently ligates the two exons while releasing the intron lariat. It is established that ALYREF settles upstream of exon-exon junctions22, however, it is unknown exactly when during splicing ALYREF engages with nascent RNA. Finally, and conspicuously, ALYREF can also be found at the 3′ends of multi-exonic transcripts23.While it is clear that the CBCA complex provides a basic platform for early factor recruitment, it remains elusive when and how these factors settle within their respective RNPs. Taking ALYREF as an example, it is unclear whether CBCA or the RNA splicing process serve to anchor ALYREF to the first exons of transcripts16. Most often embedded within introns of host transcripts, these RNAs require splicing and lariat debranching for subsequent biogenesis. Here, NEXT facilitates the resection of 3′intronic pre-snoRNA sequences and is suggested to be recruited onto these cap-distal regions via spliceosomal components sharing homology with RBM7 and ZCCHC824. As for ALYREF, the exact stage of the splicing process upon when RBM7/ZCCHC8 anchor on the RNA is unknown.Like ALYREF, the NEXT complex is involved in both cap-proximal and -distal RNA transactions. In addition to its CBCN-mediated degradation of promoter-proximal transcripts, NEXT facilitates the exosomal processing of pre-snoRNAs27. An established method of interrogation is UV cross-linking and immunoprecipitation (CLIP) coupled with high-throughput RNA sequencing28. UV irradiation covalently links RNA-RBP complexes at their sites of interaction, enabling recovery of spatial information of the given RBP throughout the transcriptome. A number of derivatives of the original protocol exist, which offer individual nucleotide resolution (iCLIP)29, enhanced crosslinking efficiency30 and reduced labour time33. Presently, however, these CLIP methods provide steady-state RNA-RBP binding information without temporal resolution. Instead, chromatin immunoprecipitation (ChIP) of RBPs and sequencing of associated nucleic acids have been used to describe co-transcriptional loading of RBPs throughout gene loci. Here, transcriptional synchronisation, using chemicals such as 5,6-Dichloro-1-beta-Ribo-furanosyl Benzimidazole (DRB), has been successfully employed to capture transcription kinetics in response to protein depletion or genetic insults37. Still, while ChIP localises a given RBP to chromatin, the technique is confounded by resolution limitations as the chemical cross-linkers employed do not provide direct RNA-RBP binding information.As illustrated by the examples above, what molecular features cater to more stable RNP formation and when this occurs during transcription are understudied processes. Proteins binding to RNA via its modification, structure or specific sequence are well known, but many RBPs display promiscuous RNA-binding. Therefore, low-affinity RNA-binding might serve to locate higher affinity sites where local concentrations of proteins and RNA influence binding kinetics. Direct in vivo investigations of such dynamic RNA-protein interactions have so far been limitedHere, we combine transcriptional synchronisation with iCLIP to capture RNA-RBP binding profiles during early and steady-state phases of transcription. Using this temporal-iCLIP (tiCLIP) approach, we find that the CBC, ALYREF and RBM7 all associate across nascent RNA before establishing their distinct binding profiles. Moreover, our data suggest that specific metabolic events, such as the transesterification steps of splicing, discriminate RBP recruitment.39, we performed at least duplicate tiCLIP experiments on cell lines harbouring ‘localisation and affinity purification’ (LAP)-tagged40 versions of ALYREF, CBP20 and RBM7 , and principal component analysis (PCA) revealed that the tiCLIP coverage for ALYREF over multi-exonic TUs was unique  were analysed Fig.  and strarse Fig. . Curiousrse Fig. , implyinrse Fig. . Additiorse Fig. . Corroboque Fig. , with a que Fig. .With the generated profiles, we conclude that tiCLIP effectively captures nascent RNA-RBP binding events with the capability of reflecting preferences, or lack thereof, of early vs. steady-state associations.44). These binding patterns were also visible in individual genes  to transcript end sites (TESs). This identified two principles, which were readily visible for CBP80 and RBM7, but not for ALYREF and CBP20. Firstly, over the time course, waves of RNA-RBP associations were produced that progressively invaded the interrogated TUs Fig. . SecondlLack of visible spatiotemporal RNA-RBP waves for CBP20 and ALYREF suggested that any nascent transcript binding might be masked by enrichment of CBP20 at RNA 5′ends and dominant exon binding preference of ALYREF, respectively. This is because reads mapping to exons may originate from stable RBP-RNA interactions preceding the DRB-block, and exonic regions make up a relatively small percentage of the TU; both of which prohibit the spatial resolution necessary to assess RNA-RBP binding. Hence, we generated spatiotemporal binding profiles using intronic reads only. Indeed, all interrogated proteins now bound RNA at similar rates - and PABPN1-anchoring of ALYREF, respectively23.Co-transcriptional sampling of nascent RNA is likely to be transient, while the high-density exonic binding, displayed by both ALYREF and the CBC, reflects their steady-state RNA associations. While it is presumed that CBC binding is restricted to 5′cap-proximal regions, ALYREF has been suggested to bind both at mRNA 5′- and 3′-ends as well as upstream of exon-exon junctionsted Fig. , CBP20- ted Fig. . In cleated Fig.  and indeted Fig. . From iCted Fig. . Moreovest exon 3′ends, indicative of RNAs having completed only the first transesterification reaction of splicing, leaving the exon 3′end unconnected to its downstream exon. Upon release of the DRB-block, this CBP80 3′CLIP density peaked at t15 before returning to steady state (DMSO) levels over subsequent timepoints 16. To obtain a temporal view of this, we quantified RBM7 tiCLIP read coverage over 101nt windows centred around the 3′ends of annotated snoRNAs and their downstream host intron 3′SSs. While the steady state RNA-binding profile of RBM7 displayed a clear enrichment in the 3′intronic flank, this was abrogated upon the 3 hr DRB-block  and unadenylated (pA-) transcripts from HeLa cells depleted of the core exosome component RRP4018 (see Methods), revealed an increased abundance of 3′ends immediately downstream of the 9nt and 25nt peaks identified by RBM7 3′CLIP analysis of H/ACA- and CD-box snoRNAs, respectively , phosphorylated at its Ser2 and Ser5 residues, was identified57. It is therefore possible that the CBC ‘hitches a ride’ with RNAPII to remain in close proximity to the exiting nascent RNA. In this configuration, CBP80 might be flexible to engage with downstream regions of nascent RNA to facilitate deposition of its cofactors, while remaining anchored to the 5′end of the transcript via CBP20.tiCLIP captured the dynamic RNA-binding profile of CBP20 and, via its co-precipitation, CBP80. Through its interaction with the mty Figs. . Interes23. However, our tiCLIP analysis contrasted this idea and leads us to conclude that the pre-mRNA splicing process is the main driver of ALYREF anchoring; also, at transcript 5′ends. This is based on three lines of evidence. First, low ALYREF coverage was detected over mono-exonic RNAs , supplemented with 10% foetal bovine serum and 1% penicillin/streptavidin, and incubated at 37 °C and with 5% CO2, unless stated otherwise.HeLa Kyoto cell lines containing stable integrations of LAP-tagged ALYREF, CBP80 or RBM7 were used for tiCLIP experiments and were sourced from Poser et al.Cells were grown to 70% confluency. To initiate the DRB block, the existing media was replaced with preconditioned media containing DRB (100μM) and incubated for 3 hrs. To release the DRB block, the DRB-containing media was aspirated and cells were washed twice with PBS before adding fresh media. Next, the cells were returned to the incubator for the appropriate time frame and then UV cross-linked. For timepoint 0, samples were UV cross-linked (see below) without replacing the media. For the non-synchronised control (DMSO), cells were incubated with DMSO-containing media for 3 hrs prior to UV cross-linking.Ice-cold PBS was used for washes and cell collection. 20 sec before the end of the timepoint, the cell media was rapidly aspirated and followed by two washes with PBS to remove residual media. Next, the minimum volume of PBS required to cover the cells was added to the cell culture plate. The cell culture plate was then placed on a bed of ice and UV cross-linked for 30 sec using 254 nm (UVC) irradiation (Stratagene). UV cross-linked samples were lifted from plates using cell scrapers before collecting in PBS. These samples were then pelleted, snap frozen and stored at −80 °C.70. Key oligonucleotides and reagents can be found in Supplementary Table g for 20 min at 4 °C. Next, 100 µL of GFP-Trap (Chromotek) bead slurry was prewashed with LB and added to 1.9 mL of cleared lysate and rotated for 1 hr at 4 °C at 16 rpm. Bead slurries were then washed three times with LB, two times with high salt wash (HSW), two times with LiCl Buffer (LCW), one time with no-salt buffer (NSB), and one time with dephosphorylation buffer (DPB). Next, on bead dephosphorylation of RNAs from recovered ribonucleoproteins was performed using FastAP (ThermoFisher), supplemented with Ribolock (ThermoFisher) for 20 min at 37 °C at 1200 rpm. The dephosphorylation mix was aspirated while bead slurries were magnetically precipitated and then washed twice with phosphate wash buffer (PWB) and two times with 1× ligation buffer (LigBx1). Next, on-bead ligation was performed using pre-adenylated barcoded L3. Beads were incubated with 2 µL of 10× ligation buffer , 5 µL PEG-400, 0.5 µL RNase Inhibitor (Promega), 1 µL T4 RNA ligase 2 truncated KQ (NEB) and 2 µL of the appropriate pre-adenylated barcoded L3 linker (10 µM) and incubated for 16 hrs at 16 °C with shaking at 1250 rpm for 15 sec every 1.5 min. Beads were precipitated on a magnet and linker ligation mixes were aspirated. Beads were washed one time with HSW, two times with LB and one time with polynucleotide kinase wash (PNKW). RNAs recovered from RNPs were radiolabelled with gamma-ATP using polynucleotide kinase (NEB) for 45 mins at 37 °C. Phosphorylation was terminated by adding 100 µL of phosphatase buffer mix and radioactive PNK mix was aspirated whilst beads were precipitated on magnets. Radiolabelled RNPs were washed three times with NSW and resuspended in 20 µL of 1.5× NuPAGE loading buffer and incubated for 10 mins at 70 °C using 1100 rpm. Following this, 1 µL DTT (1 M) was added, and microfuge tubes were incubated for a further 5 mins at 95 °C with no shaking. The radiolabelled RNPs were then separated using SDS-PAGE and transferred to a nitrocellulose membrane and detected by exposing to x-ray films. Autoradiograms were used to guide isolation of radiolabelled RNPs from the nitrocellulose membrane. Sections of membranes were cut at least 15 kDa above the expected migration of the un-crosslinked protein as recovery of RNPs above this range ensured that RNA:L3 barcoded adapters were at least 50nt in length71, which was compatible with sequencing and downstream analysis. Excised membrane pieces were incubated with 400 µL of PK-buffer and 10 µL of proteinase K (Fischer Scientific) for 2 hrs at 55 °C to release cross-linked RNAs. RNAs were then extracted using phenol-chloroform and precipitated using EtOH and sodium acetate (pH 5.5). cDNAs were synthesised using Superscript IV reverse transcriptase (ThermoFisher) using a RtCLIP primer barcoded primer. Thereafter, RNA was removed from reverse transcriptase reactions by alkaline hydrolysis, and cDNA was precipitated using EtOH and sodium acetate. Next, cDNAs were separated on a 1× TBE Urea 6% polyacrylamide gel (Novex) and 3 bands were excised from the gel: high (135nt to 215nt), middle (100nt to 135nt) and low (85 nt to 100 nt), which represent cDNA lengths of 75nt to 150nt, 35nt to 75nt, 35nt to 20nt, respectively. Gel pieces were placed in a 1.5 mL microfuge tube and crushed using a 1 mL syringe plunger. 400 µL of TE buffer was added to the crushed gel pieces and the sample was snap frozen before incubating at 37 °C for 2 hrs with 1200 rpm shaking. Gel pieces were removed from the eluate by centrifugation through columns (Costar) plugged with 2 glass filters. The eluate was precipitated using EtOH and sodium acetate. Next, the PCR template was prepared by circularising the cDNA using CircLigse II (Epicentre) for 1 hr at 60 °C. This intramolecular ligation reaction ligated the 5′adapter contained in the RtCLIP reverse transcriptase primer to the 3′end of the cDNA. Next, cDNA circles were incubated with 1 µL of cut oligo [10 µM] by heating to 95 °C for 1 min, and then decreasing the temperature every 20 sec by 1 °C until 25 °C was reached. Next, 1 µL of BamHI (Fast Fermentas) was added and samples were incubated for 30 min at 37 °C, followed by 5 min 80 °C. Linearised cDNAs were precipitated using EtOH and sodium acetate. Finally, PCR libraries were generated by using Solexa_P5 and Solexa_P3 primers required for Illumina sequencing and amplified by Accuprime Supermix enzyme (Invitrogen). High, middle and low cDNA samples were precipitated using AmpureXP beads to remove primers and mixed 5:5:1 ratio. All tiCLIP libraries were pooled before sequencing using 75 bp paired-end sequencing.The steps for tiCLIP library construction were inspired byFor RNase titration assays displayed in Supplementary Fig. 72. Oligos were 5′ adenylated using 5′ DNA adenylation kit (NEB), purified using ssDNA/RNA Clean & Concentrator columns (Zymo Research), and diluted to 10 µM before storing at −20 °C till required.Previously tested in-line barcodes  in PBST (0.05% Tween) for 1 hr and incubated with primary antibodies ; Santa Cruz)was diluted 1:500 in PBS and incubated for 1 h at room temperature, washed three times with PBS, and incubated with secondary antibody (Alexa 488 conjugated Goat Anti-mouse IgG (H + L); ThermoFisher), which was diluted 1:1000 in PBS. Cells were imaged on a Zeiss AxioObserver microscope with a Hxp120V light source, an Axiocam 702 mono camera and a Plan-Neofluar 40 × 0.75 NA objective.73 and TrimGalore (10.5281/zenodo.5127899). The tiCLIP sample information and their associated L3 adapter and RtCLIP primer barcode sequences can be found in Supplementary Data 74, using default parameters with the following options: --rna-strandness FR –fr –phred33 –no-softclip. Unmapped and non-primary reads were discarded using SAMtools75. Mapped PCR duplicates were removed using UMI barcodes and the first mapping coordinate76. After assessing the raw mapped reads, CBP20-3 was omitted from the analyses due to a consistently low number of mapped reads across all timepoint and samples . This calculation was made given that the intervening distance between CBP80 and CBP20-LAP migrations was 65 kDa , which assumes all libraries have equal RNA inputs. For each tiCLIP library, the counts of reads mapping to 20 rRNA annotations  were summed and divided by 30,000 (arbitrary number) to create an rRNA factor  were generated by calculating the relative distance from the TSSs when only considering exonic segments. TUs were normalised for length by dividing each mature TU into 100 equally sized bins. Cross-link positions within each bin were aggregated and divided by the sum of all bins for that given TU. Only TUs >200nt in length with >20 mapped CLIP reads were considered. Displayed is the average profile of all TUs. K-means clustering analysis was performed on all of the TU profiles using 3 centres for each biological replicate of ALYREF-DMSO independently. The results from each biological replicate were intersected to produce bona fide TU profile groups.+/pA− data generated from HeLa cell total RNA after depletion of the exosome core component RRP40 or treated with control siRNA were downloaded from GEO Series accession number GSE13761218. The specific bigwig files were sourced from the following accession numbers GSM4083151, GSM4083150, GSM4083149, GSM4083132, GSM4083131 and GSM4083130. Briefly, bigwig files were converted to bedGraph files and then intersected with snoRNA annotation bed files, containing coordinates for the snoRNA 100nt up- and downstream regions, using bedtools intersect and map subcommands77. The resultant count files were converted to coverage profiles using custom R code employing tidyr and ggplot2 packages.RNA 3′-end-seq pAp < 0.05.Where statistical tests were used, their identity is indicated in the appropriate figure legend. Values were considered significant when Further information on research design is available in the Supplementary InformationPeer Review FileDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Reporting Summary


However, the direct detection of viruses has so far posed a challenge as the viral genome is often encapsidated by a proteinaceous layer surrounded by a lipid envelope. This necessitates an additional and undesired RNA extraction step prior to RT-qPCR amplification. To circumvent this limitation, we have developed a direct RT-qPCR method for the detection of RNA viruses. In our method, we provide a proof-of-concept using phage phi6, a safe-to-use proxy for pathogenic enveloped RNA viruses that is commonly utilized in e.g. aerosolization studies. First, the phage phi6 envelope is removed by 1% chloroform treatment and the virus is then directly quantified by RT-qPCR. To identify false negative results, firefly luciferase is included as a synthetic external control. Thanks to the duplex format, our direct RT-qPCR method reduces the reagents needed and provides an easy to implement and broadly applicable, fast, and cost-effective tool for the quantitative analysis of enveloped RNA viruses.  Specifications tableThe COVID-19 pandemic has called to attention the need for rapid yet sensitive and cost-effective methods for direct detection and quantification of airborne viruses. The most commonly used methods rely on isolating and purifying the viral RNA genome followed by reverse transcription (RT) and quantitative polymerase chain reaction (qPCR). However, the applicability of these methods has been hampered by the high demand for viral RNA purification reagents. A handful of direct (i.e. RNA extraction-free) RT-qPCR methods have been recently developed for SARS-CoV-2 virus detection [Since there are inherent risks in working with pathogenic viruses, bacteriophages are often utilized as safe-to-use alternative models Required reagents and equipment•DNA template plasmids for preparation of RNA standard and control RNA, Addgene plasmids #101156 and #182535.•Primers and hydrolysis qPCR probes see , Metabio•P2 RNA standard and Firefly Luciferase RNA , ThermoFisher Scientific, product no. EP0111.•DNase I (1 U/µL), RNase-free, ThermoFisher Scientific, product no. EN0521.•RiboLock RNase inhibitor (40 U/µL), ThermoFisher Scientific, product no. EO0381.•Agarose, Molecular Biology grade, Fisher BioReagents, product no. BP160–100, CAS: 9012–36–6.•1xTAE buffer .•Razors or gel cutters.•NTP Set (100 mM each), ThermoFisher Scientific, product no. R0481. Note that 25 mM dilution needs to be prepared.•Phenol, saturated pH 4.3, Fisher Chemical, product no. BP1751I_400, CAS: 108–95–2.•Ethanol (99.6%), grade Aa.•E.Z.N.A.® Plasmid DNA Mini Kit I, (V-spin), Omega Bio-tek, product no. D6943–01.•E.Z.N.A.® Gel Extraction Kit (V-spin), Omega Bio-tek, product no. D2500–01.•Qubit® RNA BR Assay Kit, ThermoFisher Scientific, product no. Q10210.•Chloroform, pure, Acros Organics, product no. 423550010, CAS: 67–66–3.•UltraPlex® 1-Step ToughMix® Low ROX™ (4X), QuantaBio; product no. 95168–500.•Adjustable pipettes (1–1000 µL)•Tips with aerosol barrier (1–1000 µL), low retention.•1.5 mL tubes, low retention.•Vortex.•Benchtop microcentrifuge.•Laminar cabinet for assembling qRT-PCR reactions .•Qubit® Fluorometer, ThermoFisher Scientific.•QuantStudio™ 3 Real-Time PCR System, ThermoFisher Scientific (or another qPCR system).Procedurein vitro transcription (IVT) using T7 polymerase. The template plasmids can be easily propagated in Escherichia coli and linearized with a single restriction enzyme (EcoRI). For RNA preparation, all buffers, reagents, and disposable material need to be RNase-free grade and good laboratory practices for work with RNA should be followed.The single-stranded RNA standards used for generating the calibration curve can be prepared by Template preparation for in vitro RNA synthesis1.Propagate template vectors and isolate the plasmid. For plasmid isolation, we use E.Z.N.A.® Plasmid DNA Mini Kit I (Omega Bio-tek), but any other plasmid purification kit can be used.2.EcoRI cleavage. An example for the reaction set-up is given in Linearize the plasmid by 3.Separate the linearized plasmid on a 0.8% agarose gel in 1xTAE buffer, extract the plasmid from gel and purify it using a gel extraction kit . Alternatively, the linearized plasmid can be purified directly, without agarose gel electrophoresis, using any PCR purification kit that is suitable for up to 10 kb DNA fragments. Store the purified DNA at −20 °C.In vitro RNA synthesis1.To synthesize single-stranded RNA, assemble the IVT reactions as described in 2.Incubate the reactions at 37 °C for 2 h.3.Remove the DNA template by adding 2 µL of DNase I, mix well and incubate at 37 °C for another 15 min. Proceed immediately with RNA purification.RNA purification1.Add 250 µL of nuclease-free water to the IVT reaction.2.Add 300 µL of acidic phenol and 60 µL of chloroform.Note: The purification reaction can be upscaled. The volume of added phenol should be equal to the total reaction volume in step 1, add chloroform at 1/5 of the phenol volume.1.Vortex vigorously at full speed for 15 s.2.g for 15 min at room temperature.Separate the phases by centrifugation at 10 000 3.Collect the aqueous phase into a new tube and add 300 µL of isopropanol. Precipitate the RNA at RT for 30 min.4.g for 30 min.Pellet the RNA by centrifugation at 16 000 5.Wash the RNA by 1 mL of 80% ethanol. Vortex the sample vigorously.6.g for 10 min.Pellet the washed RNA by centrifugation at 16 000 7.Carefully remove the supernatant and air dry the RNA pellet.8.Dissolve the pellet in nuclease-free water and determine the RNA concentration with Qubit BR RNA Assay kit. Optional: Check the quality of the purified RNA by agarose gel electrophoresis. Store the purified RNA at −80 °C.The typical yield from this reaction is between 40 and 70 µg of purified RNA.1.Thaw the samples while mixing .2.Transfer 100 µL of the sample into a new Eppendorf tube.3.Add 1 µL of chloroform to each sample.4.Vortex vigorously for 15 s. Next, incubate the samples for 5 min at room temperature.5.g.Centrifuge the samples for 1 min at 10 000 6.Place the samples on ice and proceed with quantification by RT-qPCR.Note: Samples containing virus should be stored at −80 °C to preserve the RNA quality and obtain unbiased data.The RT-qPCR enzyme mix used in this assay is designed for use with hybridization probes such as TaqMan®. The enzyme mix utilizes ROX passive reference and therefore, the instrument compatibility needs to be checked prior to designing the assay. The RT reaction can be carried out in a wide range of temperatures (42 °C to 60 °C), however 50 °C proved to be the most optimal (data not shown).The assay design includes the Firefly Luciferase (FFluc) internal control template to detect possible inhibitory contaminants from the sample. The primer and probe designs were done using IDT PrimerQuest™ and primers and probes were checked for cross-reactivity with the IDT OligoAnalyzer™ tool. The primers and probes for the duplex reaction were designed according to the recommendations for multiplex qPCR design with minimal cross-reactivity between primers and probes C. The se1 – 109 genome copies) (1.Prepare the reaction mastermix as described in 2.Place the qPCR plate on ice and pipette 5 µL of the reaction mastermix into each well used of the qPCR plate.3.2O (ddH2O) instead of the template. All samples should be analyzed in technical triplicates.Add 5 µL of template/well and seal the plate. As a negative control, add 5 µL of double-distilled HNote: At the same time, construct the calibration curve using the P2 RNA standard (see section below).4.g to collect the reaction mix to the bottom of the wells.Centrifuge the plate for 1 min at 1000 5.Run the RT-qPCR reaction in a cycler using the program described in Furthermore, a wide range of primer/probe concentrations were tested in the RT-qPCR set-up to find the most optimal efficiency of the reaction. The final reaction mix constitutes the most efficient amplification (primer efficiency ∼95%) with the widest detection range (10 copies) D.1.PrepaMethod validation1, 103 and 105 PFU/reaction. As expected, the amount of genome copies detected was approx. 10-fold higher than the infective counts. This difference can be attributed to the presence of non-infective virus particles. Despite the tightly regulated packaging mechanism of phage phi6, which ensures the addition of one copy of each genome fragment into the viral capsid To validate our method, we quantified three different concentrations of 1x purified virus diluted in Tris–HCl, pH 7.2 buffer  using the standard curve generated as described above. Alternatively, the analysis can be performed using the formula below, where Cq is the quantitation cycle and Yintercept as well as the slope are derived from the calibration curve D. An exaHere we describe a simple RNA-extraction free method for direct RT-qPCR quantification of phage phi6. Additionally, we provide a detailed protocol for in-house synthesis of RNA standards, making the method cost-effective and easy to implement. Similarly to other direct RT-qPCR methods, the limitations of this method are primarily dictated by the sample volume The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.


Acropora tenuis, to establish symbioses with Symbiodiniaceae species in four genera that have varying thermal thresholds . Over a 2‐week period in January 2018, a series of both no‐choice and four‐way choice experiments were conducted at three temperatures . Symbiont acquisition success and cell proliferation were measured in individual larvae. Larvae successfully acquired and maintained symbionts of all four genera in no‐choice experiments, and >80% of larvae were infected with at least three genera when offered a four‐way choice. Unexpectedly, Gerakladium symbionts increased in dominance over time, and at high temperatures outcompeted Durusdinium, which is regarded as thermally tolerant. Although Fugacium displayed the highest thermal tolerance in culture and reached similar cell densities to the other three symbionts at 31°C, it remained a background symbiont in choice experiments, suggesting host preference for other symbiont species. Larval survivorship at 1 week was highest in larvae associated with Gerakladium and Fugacium symbionts at 27 and 30°C, however at 31°C, mortality was similar for all treatments. We hypothesize that symbionts that are currently rare in corals  may become more common and widespread in early life stages under climate warming. Uptake of such symbionts may function as a survival strategy in the wild, and has implications for reef restoration practices that use sexually produced coral stock.Reef‐building corals form nutritional symbioses with endosymbiotic dinoflagellates (Symbiodiniaceae), a relationship that facilitates the ecological success of coral reefs. These symbionts are mostly acquired anew each generation from the environment during early life stages . Symbiodiniaceae species exhibit trait variation that directly impacts the health and performance of the coral host under ocean warming. Here, we test the capacity for larvae of a horizontally transmitting coral,  Acropora tenuis larvae successfully acquired rare thermally tolerant algal symbionts, and symbiont identity influenced survival under elevated temperatures. At elevated temperatures, rare heterologous Fugacium kawagutii in the no‐choice treatment had the highest survivorship. In the four‐way choice treatment (symbiont competition), Gerakladium sp. was the most prevalent and dominant, exhibiting the highest relative abundances of homologous or heterologous symbionts.Acquiring algal symbionts is critical for the survival of coral during early life stages.  We exposed larvae of A. tenuis, a broadcast spawner and horizontal transmitter, to four species of Symbiodiniaceae in no‐choice and four‐way choice experiments over a 14 day period at 27, 30, and 31°C. Acropora tenuis is a common reef‐building coral on the Great Barrier Reef, whose adult colonies mostly associate with Symbiodiniaceae in the genus Cladocopium , and/or Durusdinium   associate with a greater proportion of thermally tolerant 22.1A. tenuis  despite their extended maintenance in tanks. While coral larvae can begin to settle days to weeks post‐spawning, the upper length of duration they are able to remain in the water column is not well known. A study of another acroporid, Acropora latistella, and four other species on the Great Barrier Reef found that after an initial drop in survivorship during the first month, there was low mortality until 100 days, with some surviving past 200 days  observed a median survivorship of 57 days, at which point larvae were observed to successfully settle. Additionally, Nishikawa et al.  being an important selection criterion. Symbiodiniaceae used in this experiment have been maintained in culture  at the AIMS Symbiont Culture Facility: Cultures were kept at 27°C in a Steridium E500 growth chamber on a 14:10 h light/dark cycle at 65 ± 10 μmol photons m−2 s−1 (Sylvania FHO24W/T5/865 fluorescent tubes). The Cladocopium C1acro symbiont culture has been previously used successfully in infection studies of larval A. tenuis  in 0.2 µm FSW with one jar incubated at each temperature. After acclimation, larvae in each temperature were transferred into 16 transparent 500 ml plastic jars, which were preconditioned in reverse osmosis water for 72 h prior to the experiment, for inoculation treatments (n = 225–250 larvae per jar). Treatments included no‐choice exposures to each of the four species individually and a four‐way choice exposure . Each of these five treatments were replicated in three jars at each of the three temperatures, along with one jar per temperature of aposymbiotic controls (N = 48 jars). Symbiodiniaceae cell densities in each culture at each temperature treatment were counted on a hemocytometer and the algae were added to larval jars to achieve a final density of 3 × 104 cells/ml. To achieve high initial exposure densities, larvae were inoculated in 100 ml of 0.2 µm FSW for 72 h, thereafter the volume was increased to 400 ml with 50% water changes via pipette every 2–3 days for the duration of the experiment.Larvae in this experiment were exposed to Symbiodiniaceae cultures under three temperature treatments Figure   with ProteinaseK (20 mg/ml), bead beaten , and incubated (55°C 2 h); 5 m KOAc was added and samples were incubated on ice (30 min) and subsequently centrifuged at 25,000 g‐force for 15 min. Incubation was followed by an RNAse A treatment, ethanol cleanup, and resuspension in DNAse‐free water. We used previously developed primers . Fluorescence amplitude minima thresholds for droplets assigned as positive amplification for each primer set were set conservatively within manufacturer guidelines; relative fluorescence intensity thresholds were as follows: Cladocopium , Durusdinium , Fugacium , and Gerakladium . To evaluate false positives, we ran 52 no template control PCR assays (water blanks), ~14 per primer pair. Of these reactions two had one positive droplet and the remaining 50 had zero positive droplets, ascribing an assay false positive rate of 4% at a droplet positivity proportion of 0.01%. We additionally ran 104 control assays on DNA from nominally aposymbiotic larva (19–30 assays per primer pair), which had a droplet positivity proportion <0.03%. Lastly, we evaluated each primer set for specificity by running 6–17 pairwise challenge assays for each combination of primer pair and symbiont (no‐choice larva) for a total of 137 specificity tests: 8 of 14 assays of Gerakladium primers × Fugacium larva had a droplet positivity proportion of <0.04%, and all other assays maintained a proportion positive <0.02% in less than 1/3 of the tests.Digital droplet polymerase chain reaction (ddPCR) was used to quantify the number of symbiont cells per species per larva. ddPCR partitions reaction mixtures into thousands of nanoliter‐scale water–oil emulsion droplets. PCR amplification occurs within each of the individual droplets and a fluorescence threshold is used to evaluate target amplification in each droplet. Analyzing the fraction of positive and negative droplets on a Poisson distribution is used to estimate the number of gene copies in the original reaction. ddPCR reactions were run on the BioRad QX200 Droplet Digital PCR System using EvaGreen Supermix and following the manufacturer's protocol. Briefly, for each 25 µl reaction, 4 µl DNA template was added to 12.5 µl EvaGreen Supermix, 0.6 µl of each Forward and Reverse Primers (10 µm), and 7.3 µl molecular grade water. For larvae from the four‐way choice experiment, each primer set was run independently (not multiplexed). Twenty microliter of the PCR reaction mix was loaded into a sample well (DG8 Cartridge) with 70 µl of QX200 Droplet Generation Oil for EvaGreen and droplets generated on the QX200 Droplet Generator. PCRs were run on the C1000 Touch Thermal Cycler |4°C 5 min, 90°C 5 min) and subsequently read on the QX200 Droplet Reader. Each plate contained negative controls (2.5R2 > 0.98) and used to convert the number of copies per DNA extraction into the number of Symbiodiniaceae cells per larva.To calculate the number of actin or ITS2 gene copies per cell for each species of Symbiodiniaceae, cells from three replicate 300 µl samples of each culture were first counted by hemocytometer, and then DNA was extracted from the remaining 260 µl. Each extraction was serially diluted , and three replicates of each dilution were analyzed on the ddPCR machine. Copy number for each species was calculated by comparing the number of positive copies (ddPCR) to the number of counted cells  in no‐choice and four‐way choice treatments; in the four‐way choice experiments acquisition success was calculated for each symbiont species. We used binomial mixed effect models to test the effects of choice treatment (no‐choice and four‐way choice treatment), temperature, time, and their interactions on acquisition success. Then, for each symbiont species, we compared acquisition success when offered alone (the no‐choice treatments) and when offered in the presence of other species (four‐way choice).n < 5, were not included in this, or any subsequent, analysis. These groups included: four‐way choice treatment at 31°C on day 14 and no‐choice Durusdinium at 31°C on day 14 across all analyses, as well as cell density in the no‐choice Fugacium and Cladocopium at 31°C on day 14 (further described below). Due to low larval survivorship on day 14 and at 31°C, models were additionally run on the following comparisons (1) 27 and 30°C, and (2) days 3 and 7, which can be found in the supplemental information.Treatment groups with 2.810 transformed to meet normality assumptions. First, we used a linear mixed effect model to test the effects of treatment (no‐choice and four‐way choice treatment), temperature, time, and their interactions on cell densities. Then, for each species, we compared proliferation (increase over time) when alone (the no‐choice treatment) to proliferation when in competition (four‐way choice).We examined symbiont cell density (cells per larva) for each symbiont species in larvae exposed to no‐choice and four‐way choice experiments that were successfully infected; larvae that were not infected were removed. Cell densities were log2.9The four‐way choice treatment allowed us to examine competitive dynamics between Symbiodiniaceae species. Because ddPCR reactions for each species primer pair were run individually (not multiplexed), there were occasions when reactions for all species in a larva were not successful . Therefore, for all relative abundance models, only larvae where ddPCR reactions were successful for all four species were included. We conducted Kruskal–Wallis non‐parametric tests at each time point to examine the effects of temperature on the number of species that infected larvae. Next, we used linear mixed effects models to test the effects of Symbiodiniaceae species, temperature, time, and their interactions on relative abundance, which was arcsine square root transformed to meet normality assumptions of analyses.2.10We used multivariate analysis to analyze the effect of temperature treatment and time on symbiont community structure in the four‐way choice treatment. Here, we visualized the variation in symbiont community relative abundance (arcsine square root transformed) among larvae using nonmetric multidimensional scaling (nMDS) based on a Bray–Curtis dissimilarity matrix and evaluation of main effects using a two‐way PERMANOVA (999 permutations).33.1p < .01; Table p = .72) or time (p = .33) or their interactions. Acquisition success in no‐choice treatments was not uniformly higher in homologous Cladocopium than heterologous  symbionts and was high in the four‐way choice experiment across temperature and time (>92%). Across all temperature treatments, larvae exposed to either Cladocopium or Fugacium symbionts only had the lowest mean acquisition success on day 3 . However, by day 14, >75% of the larvae at 27 and 30°C were infected in all treatments except the no‐choice Fugacium, which was significantly lower with a mean of 47% (27°C) and 67% (30°C).Acquisition success  over the course of the experiment differed between symbiont treatments  but proliferated to the highest mean cell densities of all choice treatments on day 14 (2060 cells per larva). We did not find evidence of a negative effect of 30°C on mean cell density or proliferation in no‐choice Cladocopium . No‐choice Durusdinium had high cell densities over time and was not negatively impacted by 30°C , however, there were significantly reduced cell densities on day 7 at 31°C . Although there was no difference in cell densities on day 3 in no‐choice Durusdinium between temperature treatments , there was significant increase in symbiont cells at 27°C . No‐choice Fugacium had lower cell densities than all other choice groups on day 14 at both 27 and 30°C. However, elevated temperature did not negatively affect no‐choice Fugacium cell densities at any time point , temperature, time, and their interactions on symbiont cell densities in larvae that had acquired symbionts  compared to the four‐way choice. There were also higher cell densities in the no‐choice Durusdinium as compared to the four‐way choice on day 7 at 27°C. There was a significant interaction of choice and time , however, there was no difference by day 7 , and no‐choice groups for these species were higher than the four‐way treatment at 27°C by the end of the experiment .To understand how symbiont choice influenced cell proliferation, we compared cell densities over the course of the experiment for each symbiont species between no‐choice and four‐way choice treatments across the range of temperatures Table . There w3.3p < .01; Table p < .01), and reduced the percent of larvae infected with four species at all time points  or four species (40%) Figure , and thes Figure .N = 208; Table Gerakladium—except for two individuals at 27°C and one individual at 30°C on day 3 , 30°C (53%), and 31°C (78%) on day 3. Despite high infectivity, Fugacium was only dominant in a single larva at 30°C and three larvae at 31°C. Elevated temperatures negatively affected the presence and dominance of Cladocopium. At 27°C, Cladocopium was present in 75% of the three‐species larvae on day 3, despite only being the dominant species in 6%. The infectivity of Cladocopium increased over time at this temperature and it was present in 100% of the larvae on day 14 and was the dominant species in 48%. At both elevated temperatures, Cladocopium was present in fewer larvae as compared to those at 27°C.Infectivity and dominance in a larva varied by species. Every symbiotic larva (98.5%) in the four‐way choice experiment , which was further modulated by temperature (p < .01) and time . Cladocopium, initially at 13%, was the only species to significantly proliferate over time , increasing to 36% by day 14, at which point Durusdinium, Gerakladium, and Cladocopium each represented approximately one third of the relative abundances . On day 3, the relative abundance of Fugacium (14%) was similar to Cladocopium, however, over time it was outcompeted  by the other three species, and was the only symbiont to significantly decrease in relative abundance over time .Larval symbiont community structure was influenced by both temperature  and day 7 (59%); however, there was no difference between the two by day 14 . At 31°C, Gerakladium was again the most abundant symbiont on day 3 (66%) and day 7 (62%), and there was a greater difference in mean relative abundances between Gerakladium and Durusdinium at 31°C compared to 30°C. Relative abundance of Durusdinium on day 3 did not differ between temperature treatments . Cladocopium was less competitive as temperatures increased. Cladocopium and Fugacium represented low mean relative abundance on day 3 (both 6%); however, unlike Cladocopium at 27°C, neither increased over time at 30°C  or 31°C .At elevated temperatures, 3.4p < .01) with the lowest survivorship in larvae exposed to the high temperature treatments  and 30°C (31%), followed by no‐choice Gerakladium . Larvae in the no‐choice Fugacium and Gerakladium treatments were the only groups to not experience significantly increased mortality on day 7 at 30°C compared to 27°C . Survivorship across all choice treatments dropped significantly between days 7 and 14 , and larvae in all choice treatments had higher survivorship at 27°C than at either of the two elevated temperatures.Symbiont choice treatments responded differently to elevated temperature  and lower survival as compared to Fugacium  at 30°C on day 7. At 30°C, however, larvae had higher survivorship in the four‐way choice treatment than those in no‐choice Durusdinium treatment , and lower survivorship than the no‐choice Fugacium treatment , although these differences were no longer significant by day 14 .Survivorship was not significantly different between the four‐way choice treatment and any of the no‐choice acquisition treatments  and a homologous thermally sensitive symbiont (Cladocopium species) in a no‐choice setting contrasted with acquisition dynamics under four‐way choice in A. tenuis larvae. We found that Gerakladium, which is normally a background symbiont, achieved similar cell densities to Cladocopium and Durusdinium at 27°C, and had even higher acquisition and abundance at elevated temperatures (30 and 31°C). These findings show that A. tenuis larvae can establish symbiosis with Gerakladium when symbionts are available at high densities and that Gerakladium can infect larvae when Cladocopium and Durusdinium symbionts are present. We found that the number of algal symbiont species simultaneously acquired by A. tenuis larvae decreased with temperature, which may be due to the negative effect of elevated temperatures on some of the symbiont species, or possibly, an increased selectivity of A. tenuis larvae for certain symbiont species at elevated temperatures. Our findings further demonstrate that symbiont identity matters for larval survival under high temperature   and the negative consequences of random uptake, resulting in suboptimal taxa  that may outcompete higher performers. In contrast, our results in larvae show higher survivorship in the four‐way choice treatment at 14 days, where larvae had high relative abundances of Fugacium and Gerakladium. This difference could be related to symbiont competition resulting in a trade‐off in juveniles that may not occur in the comparatively shorter larval stage. Although survival benefits were small, these differences were significant and as climate change stressors increase, even marginal gains in survival are important to consider. Here, survivorship after 1 week is influenced the most, and as temperatures and time increase, this effect becomes smaller. However, even the small but significant differences, for example in no‐choice Fugacium on day 7 at 30°C, could make the difference for more larvae surviving long enough to settle, which would need to be tested, along with if those symbionts are beneficial at the juvenile stage or not. With the growing trend of annual bleaching on the reef, and the huge impact those events are having on recruitment  is not a known dominant symbiont in any reef‐building coral, and therefore, less is known about its functional traits in hospite . While Gerakladium is found predominantly in symbiosis with sponges  than typically found as a background symbiont , with increasing infectivity and competitive capacity at warmer temperatures. The low cell densities could be the result of A. tenuis not actively attracting Fugacium, which could be due to a mismatch in symbiont attraction mechanisms. For example, a recent study by Takeuchi et al.  are, on average, smaller than Durusdinium, Cladocopium, and Fugacium species measured in LaJeunesse et al.  also impacts proliferation within the host. Growth rate for the cultures used here was previously measured in Chakravarti et al.  in the symbiont, which leak into the host tissues and causes significant damage to the host, leading to bleaching  and Durusdinium produced significantly more ROS than Fugacium (F2) at ambient (26°C) in an ex situ experiment  of Cladocopium C1acro . This supports our hypothesis that shifts toward rare or background symbionts may occur in coral larvae under future climate change conditions and warrants further study.Lastly, natural larval acquisition experiments under ocean warming are occurring on reefs around the world due to global climate change, yet we have not observed large symbiont shifts in adult or juvenile corals on reefs. The number of studies using genetic methods that reveal rare symbionts  is still relatively low and recent studies have found that more coral species host diverse assemblages that include rare or background symbionts  and in larvae offered a four‐way symbiont choice. Our observations at the larval stage point to a new direction in evaluating approaches to symbiont optimization strategies. Acropora larvae typically become competent to settle after ~5 days (Graham et al., Gerakladium, could persist into the adult stage. Third, it is important to consider nutritional properties of the symbiont and whether there is a trade‐off between thermal tolerance and capacity to meet the host's energetic needs. Together, these considerations will improve our understanding of the role of currently unusual symbionts in driving host thermal tolerance and fitness. Because both of the strains of Fugacium and Gerakladium tested here were isolated from cnidarians from the GBR and are present on the GBR (Fujise et al., Symbiosis manipulation has been suggested as a possible mechanism of human assisted evolution to aid in coral restoration practices under increasingly warmer conditions (van Oppen et al., The authors declare no conflict of interest.SM, LC, MvO, and RG conceived of and designed the study. SM and LC conducted the experiments. SM, AH, CN, and RC analyzed the data. SM and AH prepared the original draft of the manuscript with writing, reviewing, and editing from MvO, LC, CN, and RC.Figures S1–S6Click here for additional data file.Tables S1–S13Click here for additional data file.


The small multidrug resistance (SMR) family is composed of widespread microbial membrane proteins that fulfill different transport functions. Four functional SMR subtypes have been identified, which variously transport the small, charged metabolite guanidinium, bulky hydrophobic drugs and antiseptics, polyamines, and glycolipids across the membrane bilayer. The transporters possess a minimalist architecture, with ∼100-residue subunits that require assembly into homodimers or heterodimers for transport. In part because of their simple construction, the SMRs are a tractable system for biochemical and biophysical analysis. Studies of SMR transporters over the last 25 years have yielded deep insights for diverse fields, including membrane protein topology and evolution, mechanisms of membrane transport, and bacterial multidrug resistance. Here, we review recent advances in understanding the structures and functions of SMR transporters. New molecular structures of SMRs representing two of the four functional subtypes reveal the conserved structural features that have permitted the emergence of disparate substrate transport functions in the SMR family and illuminate structural similarities with a distantly related membrane transporter family, SLC35/DMT. From atomistic descriptions of membrane transport mechanism to global spread of multidrug resistance over the last century, small multidrug resistance (SMR) proteins have provided broad insights along multiple research fronts since the family’s discovery in the mid 1990s . With juA). The resting membrane potential and pH gradient of most bacteria implies that they typically function in the active efflux of substrates. Four major functional subtypes have been described within the SMR family, and according to our bioinformatic analysis described later in this review, at least 97% of bacterial SMR genes correspond to one of these four subtypes. The first, and likely primal, SMR subtype transports guanidinium ion, a small cationic byproduct of nitrogen metabolism, and is referred to as Gdx (guanidinium export) , are promiscuous exporters of hydrophobic cationic compounds, including quaternary ammonium antiseptics like benzalkonium and cetyltrimethylammonium , and polyaromatic cationic biocides like methyl viologen , acriflavine, and ethidium  has been implicated in the transport of small polyamine metabolites like spermidine and putrescine (arnE/arnF) acts as a glycolipid flippase  . These aethidium , 6, 7. Tethidium , and thitrescine , and theflippase , and finB, left), where they can oligomerize with antiparallel subunits . Sometimes called “paired SMRs” or PSMRs , in which a single protein possesses structurally homologous domains arranged antiparallel with respect to each other .Although dual topology and fixed antiparallel topology are only rarely observed among membrane proteins , 22, 23,Fig. 1ch other , 25, 26.ch other , 28, no ch other . Alternach other   genomes . This seequences , 33, 34.equences , 36, we −20. Consensus sequences for each SMR subtype are shown in A and B, and sequence information and annotations for individual SMR proteins from this set of genomes is available for download from the Deep Blue Data repository hosted by the University of Michigan with unique identifier doi.org/10.7302/0ynd-b343.SMR genes were identified from GEBA genomes with HMMER3.3.2  using a A). This count of genomes with multiple SMR genes reflects both paired SMR genes that encode heterodimers, as well as genomes with more than one SMR functional subtype. The majority of SMR genes have no other SMR gene within 100 base pairs, suggesting they are expressed independently. As expected for dual topology proteins (B). SMR genes are also found as adjacent gene pairs. In our dataset, >95% of adjacent gene pairs encode subunits with opposite Arg/Lys biases. The Arg/Lys distributions are centered around +4 and −4 for inward- and outward-facing protomers, respectively, which assemble to form heterodimeric transporters (B). Approximately 10% of Qac transporters and ∼20% of Gdx transporters are encoded by such paired genes, as are all polyamine transporters (C). In the GEBA genome set, all the SMR lipid transporters are encoded by singleton genes, although functional pairs have been identified in some bacteria . Qac genes are also frequent, found in ∼25% of bacterial genomes overall, including ∼50% of Proteobacteria, ∼30% of Actinobacteria, and ∼25% of Firmicutes. Many species possess both Gdx and Qac transporters. The lipid and polyamine transporters are less common and found mainly in Proteobacteria, where they are found in <10% of species (F). Thus, the SMR transporters are widespread among bacteria and dual topology Gdx and Qac transporters are the predominant SMR variants.Most bacterial phyla possess genes encoding SMRs. The most prevalent SMRs are Gdx, which are found in about ∼50% of all bacterial genomes, including 80% of Actinobacteria, half of Proteobacteria and rmicutes , D and E species F. Thus, Suppressor of GroEL mutations) (+) was established  . Howevertations) , 35, 39.tations) , 40, 41,tations) , 43, 44.ablished . These p+ has been recognized as a byproduct of bacterial metabolism since the late 1800s, when high concentrations of Gdm+ were found in spoiled meats . Dimerization is mediated primarily by TM helix 4. The extramembrane loops also form extensive hydrogen bonded cross-subunit interactions to seal the closed side of the transporter. Each subunit of the dimer is composed of two discrete lobes delineated by a conserved Gly-Ile-Gly motif that acts as a fulcrum in TM helix 3 (TM3). The subunits differ according to a ∼35° rotation between N- and C-terminal lobes that stems from a difference in the angle of the Gly-Ile-Gly kink (B). The outward facing to inward facing conformational transition involves a structural swap between the two subunits, each changing the degree of rotation between the N- and C-terminal lobes. As a result, the inward- and outward-open conformations are 2-fold symmetric to each other, and the structural swap opens an identical, symmetry-related aqueous cavity on the opposite side of the membrane (A). This elegant mechanism for the alternating access of the substrate-binding site—a prototype for the “rocker-switch” mechanism used by many other transporters—was first predicted and demonstrated for EmrE . Mutation of any one of these residues to remove H-bonding capacity substantially impairs transport function  . Thests (D) , 62, sugA and TM2B, which form one side of the binding pocket and splay apart on the open side of the transporter. The gap between these helices is lined by hydrophobic sidechains and could, in principle, permit substrate access between the aqueous substrate-binding site and the membrane interior (E). In EmrE, spectroscopic experiments lead to the suggestion that the hydrophobic residues lining this portal act as a gate that permits access for the lipophilic substrates to the binding site  and pharmaceuticals like metformin, which is excreted into wastewater, where it is slow to degrade and accumulates to levels of environmental concern . Early, low resolution electron microscopy (EM) of 2D crystals demonstrated the unusual antiparallel architecture and established an elementary understanding of the helical connectivity and protein fold  are low quality and incomplete (only Cα atoms are modeled) and experimental maps are unavailable in the Protein Data Bank. These prior structures are broadly considered inadequate for molecular inference .Both the recent structures and the long history of functional data show that EmrE has many major mechanistic features in common with Gdx-Clo, including the central glutamates in the binding pocket that contribute to alternate binding of drug and protons , 83, 84,structed B.C). As a result, the central glutamates and conserved binding site Trp (W63 in EmrE) are comparatively unconstrained and able to adopt different rotamers in the presence of different substrates. EmrE’s central glutamates move closer together or farther apart to accommodate flat planar substrates or bulky quaternary compounds (D). At the same time, the binding site Trp rotates over ∼80° to stack against aromatic rings of different substrates bound in different poses (D). Unlike Gdx-Clo, in which the residues that contribute to the H-bond stack stabilizing the central glutamates cannot be altered  and single proton–bound transporter have been reported , 90, 95, for Gdx . These s for Gdx , 97.in vivo and in vitro. In one proposal, the central glutamates are electrostatically independent so that the proton cannot “hop” from the glutamate with the lower pKa to that with the higher pKa, preventing proton release from the singly protonated state after the conformational swap  are found in larger biosynthetic operons that contribute to the chemical modification of lipid A in the outer membrane with 4-amino-4-deoxy-l-arabinose (L-Ara4N)  . This syL-Ara4N) . In S. erabinose , a lipidrabinose . Deletiois route . There acolipids , 122.via facilitated diffusion might be sufficient. However, this has not been established experimentally. A membrane portal similar to that observed in structures of EmrE and Gdx-Clo would be an obviously useful feature for lipid transport, permitting the substrate’s prenyl tail access to the membrane while the polar headgroup is ensconced within its protein-binding site, a familiar feature of other lipid flippases . DMT and SMR transporters have previously been proposed to be evolutionarily related .Although other dual topology transporter families have representatives with inverted repeat topology , 28, theFig. 6 related , 130. Homembrane . The secvia convergent evolution cannot be ruled out; structural correspondence between proteins with similar functions is usually considered evidence for evolutionary relatedness (B). This model posits an ancestral, dual topology transporter with the SMR fold and a TM helix insertion between TM helices 1 and 2. The introduction of this new TM helix would enforce a reorientation of TM helices 2, 3, and 4 relative to TM helix 1, disrupting the packing between TM helix 1 and the other TM helices in that monomer . The 3D SMR fold could be preserved, however, by a domain swap during dimer assembly  such that the now-inverted TM1 from the first monomer trades positions with TM1∗ in the opposite subunit and vice versa . TM helix domain swaps have been observed in other membrane proteins . Duplication/fusion of 4-TM dual topology transporters are not unprecedented (Although the possibility that the similarities between the SMR and SLC35/DMT folds arose atedness , 132. Moatedness B. This mproteins , 134 andproteins , and an proteins . Finallycedented , but thicedented , 29. Altcedented , 136.+, drugs, polyamines, and glycolipids. Moreover, structural homology with SLC35/DMT transporters suggests that, contrary to the proposal that the SMRs are unusual in having not evolved fused, inverted repeat architecture (The SMRs have provided a tremendously productive system for studying membrane protein evolution, transport mechanism, and microbial multidrug resistance. After 25 years of such studies, the SMR transporters finally joined the high resolution structural era in 2021. These recent crystal structures representing two of the four known functional subtypes have provided a platform for analyzing prior functional studies and understanding the structural features that contribute to substrate binding and transport for each SMR subtype: Gdmitecture , it is pitecture , 138 or itecture .In addition, the molecular framework described here opens the door for future integrative functional, structural, and computational studies to understand how the SMR scaffold has been tailored to transport diverse substrates as the family has evolved. Such lines of inquiry are urgently important as bacteria continue to evolve around us. The Qac and Gdx subtypes, in particular, have found new roles in human-impacted environments, conferring bacterial resistance to household antiseptics , “dead-eSequences and annotation for SMR proteins identified in the bacterial GEBA genome set are available for download from the Deep Blue Data repository hosted by the University of Michigan with unique identifier doi.org/10.7302/0ynd-b343.The authors declare that they have no competing interests with the contents of this article.


X-chromosome inactivation is a paradigm of epigenetic transcriptional regulation. Female human embryonic stem cells (hESCs) often undergo erosion of X-inactivation upon prolonged culture. Here, we investigate the sources of X-inactivation instability by deriving new primed pluripotent hESC lines. We find that culture media composition dramatically influenced the expression of XIST lncRNA, a key regulator of X-inactivation. hESCs cultured in a defined xenofree medium stably maintained XIST RNA expression and coating, whereas hESCs cultured in the widely used mTeSR1 medium lost XIST RNA expression. We pinpointed lithium chloride in mTeSR1 as a cause of XIST RNA loss. The addition of lithium chloride or inhibitors of GSK-3 proteins that are targeted by lithium to the defined hESC culture medium impeded XIST RNA expression. GSK-3 inhibition in differentiating female mouse embryonic stem cells and epiblast stem cells also resulted in a loss of XIST RNA expression. Together, these data may reconcile observed variations in X-inactivation in hESCs and inform the faithful culture of pluripotent stem cells. Cloutier et al. discover that human embryonic stem cells (hESCs) cultured with media containing inhibitors of GSK3 proteins undergo erosion of X-chromosome inactivation, which equalizes X-linked gene expression between females and males. The findings inform the faithful culture of hESCs. The promise of hPSCs relies on their ability to faithfully maintain their epigenetic and transcriptional profiles in culture.Human pluripotent stem cells (hPSCs) offer the possibility to model early human development in vitro and are substrates for regenerative medicine6. Once inactivated, with a few key exceptions, replicated copies of the silenced X chromosome remain stably inactive in descendant cells7. X-inactivation is an experimentally tractable system to interrogate epigenetic transcriptional regulation because two equivalent X chromosomes become transcriptionally divergent and these divergent transcriptional states are subsequently stably transmitted across mitotic cell division.X-chromosome inactivation is a paradigm of epigenetic transcriptional regulation that equalizes X-linked gene expression between female and male mammals10. At the peri-implantation stage, the inactivated paternal X chromosome is reactivated in the pluripotent epiblast progenitor cells12. Conventionally cultured mESCs capture this transient population of pluripotent cells and harbor two active X chromosomes13. Upon differentiation, pluripotent mouse embryonic epiblast cells as well as mESCs inactivate either the maternal or the paternal X chromosome in individual cells in a process termed random X-inactivation18.X-inactivation has been studied extensively in mouse embryos and mouse embryonic stem cells (mESCs). In the female preimplantation mouse embryo, all cells undergo imprinted inactivation of the paternal X chromosome22. Instead, both X chromosomes in female preimplantation human embryos appear to initiate some degree of silencing22. The X chromosome-encoded long noncoding XIST RNA, whose expression is a hallmark of the inactive X chromosome and is required for stable X-inactivation in mice23, is expressed from and coats in cis both X chromosomes in most cells of female human blastocyst-stage embryos21. XIST RNA coating in turn recruits a diverse array of proteins to the X chromosome that silence gene expression28. Despite XIST RNA coating of the X chromosomes, however, X-linked genes are not fully silenced in preimplantation human embryos21.In contrast to the mouse, human female preimplantation embryos do not undergo imprinted inactivation of the paternal X chromosome13, female human ESC (hESC) lines display variable patterns of X-inactivation49. This variability appears to reflect both differences in X-inactivation patterns in early mouse vs. human female embryos and potentially how hESCs are derived and cultured.Unlike female mESCs, which harbor two activeXs50. In these naïve hESC lines, a proportion of cells display XIST RNA coating of both X chromosomes but do not appear to transcriptionally inactivate the XIST RNA-coated Xs, similar to cells in early female human embryos. Most female hESCs cultured in naïve conditions, however, harbor one XIST RNA-coated X chromosome that is transcriptionally active50. The heterogeneity of XIST RNA expression in naïve female hESCs appears to be due to the coexistence in culture of at least two populations of pluripotent cells47. Blocking autocrine basic fiborblast growth factor (bFGF) signaling reduces this heterogeneity and is reported to yield nearly all hESCs with two XIST RNA-coated X chromosomes47, recapitulating the pattern observed in epiblast cells of preimplantation female human embryos21. Differentiation of these naïve hESCs into the ‘primed’ pluripotent hESCs results in a majority of the cells undergoing random X-inactivation and exhibiting XIST RNA coating of a single X chromosome that is transcriptionally inactive47.The pattern of X-inactivation in early preimplantation human embryos is partially recapitulated in vitro through the derivation of ‘naïve’ pluripotent female hESCs56. Female mEpiSCs contain one inactivated, XIST RNA-coated X chromosome14. Primed female hESCs, by contrast, exhibit at least three patterns of X-inactivation. Primed female hESCs can harbor no inactive X chromosome, one inactive-X, or a leaky inactive-X45. Upon prolonged culture, many primed female hESC lines lose XIST RNA coating, which is accompanied by the expression of a subset of previously silenced genes from the inactive X chromosome58. This loss of XIST RNA coating and re-expression of silenced X-linked genes has been termed X-inactivation erosion58. Loss of XIST RNA coating and the leaky expression of inactive X-linked genes also characterize cultured human induced pluripotent stem cells (hiPSCs) and are thought to be irreversible59. Increased X-linked gene expression due to X-inactivation erosion or failure can be deleterious to development and differentiation60. The instability of the epigenetically inactivated X chromosome lends caution to the use of pluripotent female human cells in disease modeling and regenerative medicine.Compared to naïve hESCs, primed pluripotent hESCs capture a later stage of embryonic development and may be analogous to mouse epiblast stem cells (mEpiSCs)The underlying reasons for XIST RNA loss and X-inactivation instability in cultured hESCs are unclear. To gain insights into changes in X-inactivation in hESCs, we derived new hESC lines and analyzed expression of XIST RNA and other X-linked genes in these hESCs under diverse culture conditions and across many passages. Our results implicate the presence of lithium chloride or other GSK-3 inhibitors in the culture media as a cause of XIST RNA expression loss in female hESCs.61. To minimize bias due to clonal expansion of the mechanically passaged hESCs, we quantified and stratified the percentage of nuclei with XIST RNA coats on a per colony basis with a minimum of 100 nuclei counted per colony.To test the kinetics of X-inactivation in hESCs, we derived and characterized a series of new female hESC lines under primed pluripotency conditions on human fibroblast feeder cells (HFFs) Fig. ; Methods66. In the UM33-4 hESCs, we quantified the percentage of nuclei with XIST RNA coats per colony at each passage (P) from P1 to P29  and at P27 XIST RNA coating disappeared altogether in all nuclei in all colonies. During subsequent passaging, hESC line UM33-4 stably maintained the complete absence of nuclei with XIST RNA coats. Of note, nuclei lacking XIST RNA coats were devoid of any XIST RNA FISH signals, suggesting that a lack of XIST RNA coating reflected an absence of XIST RNA expression since the RNA FISH assay detects stabilized as well as nascent RNAs61.After derivation on HFFs, we passaged the first hESC line analyzed, UM33-4, onto a Matrigel-coated surface and grew the cells at atmospheric oxygen concentration (20%) in mTeSR1 medium. Matrigel is a widely used extracellular matrix substrate that bypasses the need for feeder cells in culturing hESCs and mTeSR media are commonly used to culture hESCs on Matrigel2 concentrations. Atmospheric O2 has been reported to cause loss of XIST RNA expression in cultured female hESCs in some studies67 but not others70. We derived two new female hESC lines on HFFs, UM63-1 and UM77-2, and passaged them from P5 to P14 onto Matrigel-coated surfaces under atmospheric and physiological O2 concentrations in parallel. We quantified the proportion of XIST RNA-coated nuclei per colony in the UM63-1 and UM77-2 hESCs from P5 to P14 since UM33-4 hESCs in Fig. 2 culture conditions, both the UM63-1 and UM77-2 hESC lines displayed decreasing percentages of nuclei with XIST RNA coats per colony during passaging, recapitulating the pattern observed in the UM33-4 hESC line .To investigate the underlying causes of XIST RNA coating loss in hESCs, we next cultured hESCs in atmospheric (20%) and physiological (5%) OATRX and USP9X, by RNA FISH. Under both O2 concentrations, nuclei that had lost XIST RNAcoating maintained silencing of one ATRX allele but displayed an increased frequency of biallelic expression of USP9X across passaging .We next sought to examine other variables that could explain the loss of XIST RNA coating in cultured hESCs. After their derivation and initial culture on HFFs, the hESC lines in Figs. 66, whereas our hESC lines grown on HFFs were cultured with a XenoFree (XF) medium (see Methods for detailed culture media composition). We therefore investigated whether culture media could underlie the differences in XIST RNA coating observed in hESCs cultured on Matrigel vs. HFFs. We cultured hESC lines UM63-1 and UM77-2 on HFFs in mTeSR1 medium and in XF medium in parallel. Unexpectedly, we found that culturing hESCs on HFFs with XF medium stably maintained nuclei with XIST RNA coating, whereas hESCs cultured on HFFs with mTeSR1 medium displayed a significant decrease in the proportion of nuclei with XIST RNA coating per colony during equivalent passaging   , suggesting that the loss of XIST RNA expression is irreversible. From these findings, we conclude that culturing hESCs in XF medium stably maintains XIST RNA coating and that culturing hESCs in mTeSR1 medium causes an irreversible loss of XIST RNA expression.When hESCs were cultured on HFFs first in mTeSR1 medium and then switched to XF medium, the frequency of XIST RNA-coated nuclei per colony stabilized , XF medium does not. Lithium is known to intersect with a number of intracellular signaling pathways71, including, importantly, with the Wnt pathway by inhibiting GSK-3 proteins74. We thus tested if addition of LiCl to XF medium (XF with LiCl) at the concentration present in mTeSR1 (0.98 mM) would result in loss of XIST RNA coating. We derived an independent hESC line, UM90-14, and cultured the cells in parallel in XF, mTeSR1, and XF with LiCl media on HFFs under physiological oxygen concentration (5% O2). As in Fig. p = 0.3)   45 but not other studies75. We assayed XIST RNA coating in hESCs differentiated into embryoid bodies (EBs) with a commercial medium, AggreWellTM, that is based on the mTeSR1 medium and contains LiCl64 and with XF medium lacking the pluripotency promoting factor bFGF. bFGF is widely used as a supplement to promote growth of stem cells in vitro, including in the mTeSR1 and XF media used in this study76. XIST RNA coating decreased in a significant proportion of nuclei in EBs differentiated with the AggreWellTM medium  but not with the XF medium lacking bFGF . We next assayed XIST RNA coating in hESCs differentiated into EBs in XF medium lacking bFGF but supplemented with LiCl at the concentration present in mTeSR1 (0.98 mM) and again observed loss of XIST RNA coating in a significant proportion of nuclei   in XF medium supplemented with each of the three GSK-3 inhibitors individually. The 90-14 hESC line at P6 exhibited a slightly reduced frequency of nuclei with XIST RNA coating compared to earlier experiments , similar to hESCs cultured in XF with LiCl and mTeSR1 media 88 (Methods). We then cultured the mEpiLCs in the presence of the GSK-3 inhibitor CHIR99021 (CHIR) for 48 h and assessed Xist RNA coating. The mEpiLCs cultured without CHIR stably maintained Xist RNA coating, whereas mEpiLCs cultured with CHIR lost Xist RNA coating in a significant number of nuclei per colony   ESCs. Female mESCs harbor two active X chromosomes and undergo stochastic inactivation of one of the two Xs upon differentiation54. Like primed hESCs, mEpiSCs harbor a randomly inactivated X chromosome14. We found that a significant number of mEpiSCs also lost Xist RNA coating upon culture in media supplemented with CHIR  . mEpiSCs are primed pluripotent stem cells that are analogous to primed pluripotent hESCs90. The activation of Wnt signaling is conventionally thought to only activate transcription91, but recent studies suggest that Wnt signaling may also repress the expression of some target genes90. We therefore examined human and mouse genomic sequence 5' of the XIST/Xist transcription start sites (TSSs) and found three conserved putative TCF binding motifs within this region  and β-catenin to regulatory sequences of the target genesion Fig. . We also59. Consistent with previous findings, our data rule out atmospheric O2 concentration as a cause of XIST RNA loss in hESCs70. Instead, we found that a primary source of XIST RNA loss in hESCs is the composition of the hESC culture medium. The loss of XIST RNA expression in the hESCs is irreversible, in agreement with prior findings39. Of note, a recent report suggests that DNA methylation may contribute to the irreversibility of XIST RNA loss in cultured female hESCs that have undergone X-inactivation erosion59.PSCs are a model system to investigate epigenetic mechanisms that underlie cell fate transitions. X-inactivation is an experimentally tractable system to dissect epigenetic transcriptional regulation in PSCs. Much prior work has demonstrated that prolonged culture of female hPSCs results in loss of XIST RNA coating and erosion of X-inactivation29. Our data, though, argue against this possibility because non-XIST RNA-coated hESCs did not increase in proportion when hESCs cultured in mTeSR1 medium were switched to culture with XF medium, which stably maintained but did not lead to increased frequency of nuclei with XIST RNA coating. Instead, our data suggest that LiCl in hESC culture medium can actively cause loss of XIST RNA expression. The loss of XIST RNA coating, however, did not appear to result in chromosome-wide reactivation of all silenced X-linked genes. XIST RNA loss in the hESCs, though, has been reported to result in the reactivation of a subset of silenced X-linked genes93, consistent with our findings. Through increased expression of X-linked genes, the erosion of X-inactivation can negatively impact hESC physiology, for example through compromised differentiation45.Female hESCs lacking XIST RNA coating have been suggested to expand in culture due to a proliferation advantage compared to female hESCs with XIST RNA coating94. XACT expression also precedes loss of XIST RNA coating from the inactive-X during prolonged hESC culture95. An analysis of published female hESC RNA-Seq data, however, suggests that XACT expression is likely to be transient in cultured hESCs97.One factor suggested to cause loss of XIST RNA expression in hESCs is the X-linked XACT RNA. XACT RNA is expressed from and coats the active X chromosome at the onset of X-inactivation72. The inclusion of GSK-3 inhibitors in hESC culture media recapitulated the loss of XIST RNA coating observed with the addition of LiCl to the culture medium, suggesting that inhibition of GSK-3 proteins by lithium may be responsible for loss of XIST RNA coating in hESCs cultured in mTeSR1 medium.That culture media components like LiCl can alter X-inactivation counters the conventional notion that X-inactivation is a cell autonomous process that is immune to extracellular influences. Lithium interferes with a number of cell signaling pathways, most prominently through the inhibition of GSK-3 proteins84. GSK-3 phosphorylates TCF/LEF coactivator β-catenin, which prevents β-catenin from entering nucleus98. GSK3 inhibition permits β-catenin to enter the nucleus, where it associates with TCF/LEF transcription factors to activate gene expression99. But, recent work has found that Wnt signaling can also repress gene expression in some contexts90. TCF binding at target sites together with other cofactors that silence gene expression may repress transcription of Wnt targets89. The putative TCF binding sites and conserved surrounding sequences upstream of XIST suggests that TCF together with other cofactors that may directly repress XIST in PSCs. It’s also possible that GSK-3β and other components of the Wnt signaling pathway indirectly repress XIST by inducing the expression of genes that in turn silence XIST or by crosstalk with other signaling pathways that silence XIST101. Future work will dissect the direct vs. indirect regulation of XIST/Xist RNA expression by GSK-3 and Wnt signaling in PSCs.A primary mode of action of GSK-3 is the negative regulation of the canonical Wnt signaling pathway104. GSK-3 inhibition and Wnt activation are both suggested to promote naïve hESC pluripotency105. In naïve hESCs, XIST RNA can be expressed from and coat both X-chromosomes48. However, the XIST RNA coats in naïve hESCs appear more diffuse47, suggesting decreased XIST expression compared to that in primed pluripotent hESCs. This lower XIST RNA expression in naïve hESCs may potentially be due to the inclusion of GSK-3 inhibitors in naïve hESC culture media.Naïve hESCs are cultured with a cocktail of inhibitors that target components of a number of cellular signaling pathways, including Wnt signaling via the inhibition of GSK-322. Furthermore, whereas human XIST and mouse Xist transcripts share some sequence homology, the genomic sequences surrounding human XIST and mouse Xist are considerably divergent106. The identification of conserved TCF binding motifs and surrounding sequences upstream of the human XIST and mouse Xist TSSs suggest a possible conserved role for Wnt signaling in XIST/Xist regulation and dosage compensation in humans and mice.Although human and mouse embryos both undergo X-chromosome inactivation, the dynamics of X-inactivation differ during human and mouse embryonic developmentOur findings suggest a careful assessment and selection of culture media in order to faithfully maintain the epigenetic state of hESCs. Given that lithium chloride and other GSK-3 inhibitors can cause loss of XIST RNA expression, the use of media without LiCl and other GSK-3 inhibitors may be more appropriate in the maintenance and differentiation of primed pluripotent hESCs. The inclusion of GSK-3 inhibitors in naïve hESC culture media and their impact on the cellular epigenome may also require careful investigation.34.Although our study has shown that the inclusion of LiCl and GSK-3 inhibitors in culture media impedes expression of XIST RNA in PSCs, it does not exclude other causes of loss of XIST RNA expression in the PSCs. In addition to GSK-3 inhibition, sources of cellular stress, such as freeze-thaw cycles of hESCs, may also alter XIST RNA expressionWritten informed consent for human embryo donation was obtained from both gamete providers as outlined by NIH guidelines, and hESC line derivation was performed under University of Michigan’s Institutional Review Board approved study, “Derivation of human Embryonic Stem Cells” . This study was also approved by the Human Pluripotent Stem Cell Research Oversight (HPSCRO) Committee (HPSCRO record #1035).XIST BAC (RP13-183A17), ATRX BAC (RP11-42M11), and XIST fosmid (G135P63425C4) were obtained from the BAC PAC Resources Center (https://bacpacresources.org). USP9X BAC was obtained from Invitrogen . Normal Goat Serum (#31872) was obtained from Invitrogen. Venor™ GeM Mycoplasma Detection Kit was obtained from Sigma-Aldrich (#MP0025); RNeasy Kit was obtained from Qiagen (#74104); High Capacity cDNA Reverse Transcription Kit was obtained from Applied Biosystems (#4368814); DAPI was obtained from Invitrogen (#D21490); Cy3-dCTP and Cy5-dCTP were obtained from GE Healthcare ; Fluorescein-12-UTP was obtained from Roche (#11427857910); NaCl was obtained from ThermoFisher Scientific (#BP358); Sucrose was obtained from ThermoFisher Scientific (#BP220); MgCl2 was obtained from ThermoFisher Scientific (#AA12315A7); PIPES was obtained from Sigma-Aldrich (#P6757); Paraformaldehyde was obtained from Electron Microscopy Sciences (#15710); Vectashield was obtained from Vector Labs (#H-1000); Deionized formamide was obtained from ISC Bioexpress (VWR:#0606); SSC was obtained from Ambion (#AM9765); BSA was obtained from New England Biolabs (NEB:#B9001S); Yeast tRNA was obtained from Invitrogen (#15401–029); Sheared boiled salmon sperm was obtained from Invitrogen (#15632–011); Dextran sulfate was obtained from Millipore (#S4030); Blue Hoechst dye was obtained from ThermoFisher Scientific (#33258); Matrigel was obtained from Corning (#354277); mTeSR1 medium was obtained from StemCell Technologies (#85850); KnockoutTM DMEM was obtained from Gibco (#10829); Xenofree KnockoutTM Serum Replacement was obtained from Gibco (#12618012); GlutamaxTM was obtained from Gibco (#35050–061); β-mercaptoethanol was obtained from Sigma-Aldrich ; nonessential Amino Acid 100× was obtained from Gibco (#11140-05); Collagenase Type IV (#17104-019) was obtained from Gibco; Basic Fibroblast Growth Factor (bFGF) for culture of hESCs was obtained from MilliporeSigma (#GF003AF-100UG); DMEM F12 was obtained from Gibco (#11330); LY2090314 was obtained from Sigma-Aldrich (SML1438); BIO was obtained from Sigma-Aldrich (#B1686); Alsterpaullone was obtained from Sigma-Aldrich (#126870); AggreWellTM medium was obtained from StemCell Technologies (#05893); bFGF (#233-FB) for differentiation of mESCs was obtained from R&D Systems; L7 passage solution was obtained from Lonza, (#FP-5013); G2TM Plus medium (#10132), OvoilTM (#10029), and G-MOPSTM plus medium (#10130) were obtained from Vitrolife; Neurobasal medium (#21103049) was obtained from ThermoFisher Scientific; L-glutamine (#25030) was obtained from Gibco; Penicillin–streptomycin (#15070–063) was obtained from Gibco; B27 supplement (#17504) was obtained from Invitrogen; N2 supplement (#17502048) was obtained from Invitrogen; CHIR99021 (#04–0004) was obtained from Stemgent; MEK inhibitor PD0325901 (#04–0006) was obtained from Stemgent; LIF was obtained (#ESG1106) was obtained from Millipore; Activin A (#338-AC) was obtained from R&D Systems; Fibronectin (#F1141) was obtained from Sigma; bFGF (#233-FB) was obtained from R&D Systems; SsoAdvance SYBR Green Supermix was obtained from Bio-Rad (#1725272); 3 M Sodium Acetate was obtained from Teknova (#S0298); Triton X-100 was obtained from Fisher Scientific (#EP151); TRIzol was obtained from Life Technologies (#15596-018); 60 mm tissue culture dishes were obtained from BD Falcon (#353652). 1X PBS (#10010023) was obtained from ThermoFisher Scientific.Antibodies were used at the following concentrations and obtained from the following sources: Mouse anti-TRA 1-60, pluripotency marker, 1:200 dilution, Millipore #MAB4360; Goat anti-OCT3/4, pluripotency marker, 1:300 dilution, Santa Cruz Biotechnology #sc-8628; Rabbit anti-SOX2, pluripotency marker, 1:800 dilution, EMD Millipore #AB5603; Rabbit anti-NANOG, pluripotency marker, 1:150 dilution, Abcam #ab21624; Mouse anti-SSEA4, pluripotency marker, 1:100 dilution, Millipore #MAB4304; Donkey anti-Rabbit, secondary antibody, NANOG 1:200 dilution, SOX2 1:800 dilution, Jackson ImmunoResearch #711-165-152; Donkey anti-Goat, secondary antibody; 1:800 dilution, Jackson ImmunoResearch #705-096-147; Donkey anti-Mouse, secondary antibody; TRA 1-60: 1:100 dilution; SSEA4: 1:800 dilution, Jackson ImmunoResearch #715-165-150. https://grants.nih.gov/stem_cells/registry/current.htm).The sources and identifying information for all hESC and feeder cell lines used in this study are included in Supplementary Table 2/5% O2/89% N2 (low O2) until blastocysts were fully expanded or had completed the hatching process from the zona pellucida. The inner cell mass (ICM) of each embryo was either removed immediately or the embryo was incubated for several hrs prior to removal of the ICM, depending on the degree of blastocoele expansion. Blastocyst morphology was scored according to the Gardner and Schoolcraft criteria107.Vitrified human embryos were warmed according to the donating fertility healthcare provider vitrification/warming protocol specific for the donated embryos. Warmed day (d) 5 embryos were cultured in equilibrated 50 ul drops of G2 Plus medium , overlaid with Ovoil  in a 37 ˚C humidified incubator at 6% COThe ICM of each expanded blastocyst was isolated via laser-assisted microdissection (Hamilton Thorne Biosciences) in 10 ul drops G-MOPS Plus medium , overlaid with Ovoil , and were immobilized with adequate tension on the exposed, laser-breached area of the TE. On the opposite side, adequate tension was applied to the zona pellucida with the smaller  holding pipet and the TE/ICM complex was extracted from the zona pellucida with gentle pulling motion of the holding pipets. Each ICM was then microdissected from the overlying TE cells with infrared laser pulses, ensuring that the distance of the laser pulses to the ICM was adequate .2, in a 35 mm tissue culture dish on a coverslip  pre-coated with 0.1% recombinant human gelatin (Fibrogen) , 0.1 mM β-mercaptoethanol , 10 mM nonessential Amino Acid 100x , 4 ng/ml animal-free basic human recombinant Fibroblast Growth Factor (bFGF) . The ICM and cell outgrowths were cultured under 5% CO2/5% O2/90% N2 (low O2) in a humidified incubator at 37 °C. Attachment of ICM was evaluated 48 h after plating onto HFFs and the hESCs were first passaged (P1) 4–6 days after initial evaluation , inactivated by g-irradiation at 3.7 × 104 cells/cm2/5% O2/90% N2 (low O2) at 37 °C. During this early hESC derivation passages, epiblast-like structures were identified, grown, split, and expanded until well-recognized hESC colonies were observed  at 5% CO88. mEpiLCs utilized for RNA FISH staining were cultured, permeabilized, and fixed on fibronectin-coated (15 μg/ml) (Sigma #F1141) coverslips.To differentiate mESCs into EpiLCs, mESCs were cultured in N2B27 medium supplemented with 10 ng/ml bFGF  and 20 ng/ml Activin A  in Fibronectin (15 μg/ml) (Sigma #F1141) coated tissue culture dishes for 48 h. The mEpiLCs were then cultured without or with 3 μM CHIR99021 (CHIR) (Stemgent #04–0004) for an additional 48 h in N2B27 medium. A concentration of 3 μM was selected for CHIR because this is the CHIR concentration used in naïve mESC culture2 well containing a MEF feeder layer and cultured for an additional 4–6 days in K15F5 medium. The culture was then passaged by a brief exposure (2–3 min) to 0.05% trypsin/EDTA with gentle pipetting to prevent complete single-cell dissociation of pluripotent clusters and plated into a 9.6-cm2 well containing MEF feeders in K15F5 medium. Morphologically distinct mEpiSC colonies became evident over the next 4–8 days and were subcloned from a mixed population of cells, including mESCs. mEpiSC colonies were manually dissociated into small clusters using a glass needle and plated into 1.9-cm2 wells containing MEF feeders in mEpiSC medium consisting of Knockout DMEM  supplemented with 20% Knockout Serum Replacement , 2 mM GlutamaxTM , 1× nonessential amino acids , 0.1 mM β-mercaptoethanol , and 10-ng/ml FGF2 .mEpiSCs were derived from individual E3.5 preimplantation mouse embryos, which were plated on quiescent mouse embryonic fibroblast (MEF) feeder cells in K15F5 medium containing Knockout DMEM  supplemented with 15% Knockout Serum Replacement , 5% ES-FBS , 2 mM L-glutamine , 1X nonessential amino acids , and 0.1 mM β-mercaptoethanol . After 5–6 days, blastocyst outgrowths were dissociated partially with 0.05% trypsin . The partial dissociates were plated individually into a 1.9-cmAfter derivation, mEpiSCs were cultured in mEpiSC medium and passaged every third day using 1.5 mg/ml collagenase type IV  with pipetting into small clumps. mEpiSC medium used to culture CHIR-treated mEpiSCs was supplemented with 3 μM CHIR99021 (CHIR) (Stemgent #04–0004). mEpiSCs generated for RNA FISH staining were cultured, permeabilized, and fixed on fibronectin-coated (15 μg/ml) (Sigma #F1141) coverslips.XIST BAC ; ATRX BAC ; USP9X BAC ; XIST fosmid ] were labeled with Fluorescein-12-dUTP (Invitrogen), Cy3-dCTP , or Cy5-dCTP . Labeled probes for multiple genes were precipitated in a 3 M sodium acetate  solution along with 300 μg of yeast tRNA , and 150 μg of sheared, boiled salmon sperm DNA . The solution was then centrifuged at 21,130 X g for 20 min at 4 °C. The resulting pellet was washed in 70% ethanol, then washed in 100% ethanol, dried, and re-suspended in deionized formamide . The re-suspended probe was denatured via incubation at 90 °C for 10 min followed by an immediate 5 min incubation on ice. A 2X hybridization solution consisting of 4X SSC, 20% Dextran sulfate , and 2.5 mg/ml purified BSA  was added to the denatured probe/formamide solution. Probes were stored at −20 °C until use.Human and mouse probes [2 , and 10 mM PIPES buffer , pH 6.8 for 30 s; ice-cold CSK buffer containing 0.4% Triton X-100  for 30 s; followed twice with ice-cold CSK buffer for 30 s each. After permeabilization, cells were fixed by incubation in 4% paraformaldehyde  for 10 min. Cells were then rinsed 3 times in 70% ethanol and stored in 70% ethanol at −20 °C prior to RNA FISH staining. Prior to RNA FISH probe hybridization, coverslips were dehydrated through 2 min incubations in 70%, 85%, 95%, and 100% ethanol solutions and subsequently air-dried for 15 min. The coverslips were then hybridized to the FISH probe overnight in a humid chamber at 37 °C. The samples were then washed 3 times for 7 min each at 37 °C with 2X SSC /50% deionized formamide , 2X SSC, and 1X SSC. A 1:250,000 dilution of DAPI  was added to the third 2X SSC wash. Coverslips were then mounted on slides in Vectashield  and sealed with nail polish.hESCs, differentiated mEpiLCs, and mEpiSCs grown on coverslips were permeabilized through sequential treatment with ice-cold cytoskeletal extraction (CSK) buffer containing 100 mM NaCl , 300 mM sucrose , 3 mM MgClCoverslips containing stained cells were imaged using a Nikon Eclipse TiE inverted microscope with a Photometrics CCD camera. The images were deconvolved and uniformly processed using NIS-Elements software (Version 4.60.00).ATRX and USP9X in individual nuclei was quantified in the following categories: nuclei with XIST RNA coats and monoallelic ATRX or USP9X expression; nuclei with XIST RNA coats and biallelic ATRX or USP9X expression; nuclei without XIST RNA coats and monoallelic ATRX or USP9X expression; and, nuclei without XIST RNA coats and biallelic ATRX or USP9X expression. hESC colonies with <100 cells or colonies with excessive cell overlaps were not quantified. All measurements were taken from distinct samples. Raw quantification data for all RNA FISH experiments in this study are included in Source Data file.Expression of X-linked genes was quantified at 10X to 100X resolution starting at the upper left corner of each image. Percent expression of monoallelic XIST RNA was calculated for each colony on the coverslip and recorded in one of the following expression categories for all experiments except Fig. Total RNA was isolated from TRIzol  according to the manufacturer’s instructions. cDNA libraries were generated and sequenced on the Illumina HiSeq2000 platform to generate 75 bp single-end reads.111 (Version 2.7.10a) and counted using FeatureCounts112 (Version 1.22.2). Differential expression analysis was conducted using DESeq2 (Version 1.34.0).Quality control analysis was conducted using FastQC (Version 0.11.9). Reads were aligned to the hg19 (human) reference genome using STARRequests for hESC lines should be directed to Gary D. Smith (smithgd@umich.edu). Requests for mESC and mEpiSC lines and other resources, reagents, or information should be directed to Sundeep Kalantry .p-values for individual linear models or p-values comparing the linear models are reported, as indicated. A threshold of p = 0.05 was used to test for statistical significance. Principal Component Analysis (PCA) was conducted using R.General linear model regression analysis was performed in R to determine statistically significant differences in XIST RNA coating through passaging Figs. –10. For Further information on research design is available in the Supplementary InformationPeer Review FileReporting Summary