Vibrio cholerae is required for its persistence and evolution via horizontal gene transfer in the marine environment. Genes involved in the uptake and catabolism of the chitin disaccharide chitobiose are encoded by the chb operon. The orphan sensor kinase ChiS is critical for regulation of this locus, however, the mechanisms downstream of ChiS activation that result in expression of the chb operon are poorly understood. Using an unbiased transposon mutant screen, we uncover that the nucleoid occlusion protein SlmA is a regulator of the chb operon. SlmA has not previously been implicated in gene regulation. Also, SlmA is a member of the TetR family of proteins, which are generally transcriptional repressors. In vitro, we find that SlmA binds directly to the chb operon promoter, and in vivo, we show that this interaction is required for transcriptional activation of this locus and for chitobiose utilization. Using point mutations that disrupt distinct functions of SlmA, we find that DNA-binding, but not nucleoid occlusion, is critical for transcriptional activation. This study identifies a novel role for SlmA as a transcriptional regulator in V. cholerae in addition to its established role as a cell division licensing factor.Chitin utilization by the cholera pathogen  Vibrio cholerae is a natural resident of the aquatic environment and causes disease when ingested in the form of contaminated food or drinking water. In the aquatic environment, the shells of marine zooplankton, which are primarily composed of chitin, serve as an important food source for this pathogen. The genes required for the utilization of chitin are tightly regulated in V. cholerae, however, the exact mechanism underlying this regulation is currently unclear. Here, we uncover that a protein involved in regulating cell division is also important for regulating the genes involved in chitin utilization. This is a newly identified property for this cell division protein and the significance of a common regulator for these two disparate activities remains to be understood.The cholera pathogen  V. cholerae, the bacterium responsible for the diarrheal illness cholera, is naturally found in the marine environment. In this niche, this pathogen forms biofilms on the shells of microscopic crustaceans. The shells of these organisms are primarily composed of chitin, an insoluble polymer of β-1,4 linked N-acetylglucosamine (GlcNAc). Chitin is the second most abundant biopolymer on the planet, and Vibrio species play an important role in recycling chitin in the aquatic environment [Vibrionaceae [ionaceae .V. cholerae in the aquatic environment, biofilm formation on chitin is also important for promoting transmission of this pathogen to its human host. Indeed, cholera outbreaks are seasonal in endemic areas and closely associated with blooms in chitinous zooplankton [V. cholerae [Vibrio-chitin interactions are important for the persistence, transmission, and evolution of this important human pathogen in its environmental reservoir.In addition to promoting the survival of plankton . Furtherplankton , 4. Chitcholerae , 6. Thuschb operon, which encodes the genes required for uptake and utilization of the chitin disaccharide chitobiose [chb operon and characterize one newly identified transcriptional activator of this locus.The expression of genes required for chitin degradation, uptake, and utilization are regulated by the orphan sensor kinase ChiS , 8. ChiSchb operon, we first generated a Pchb transcriptional reporter. As expected, this reporter was activated in a cbp mutant, and this activation was dependent on ChiS . We exploited this fact and used our transcriptional reporter to perform two independent transposon mutant screens to identify potential activators and repressors of this locus. To identify repressors, we used a Pchb-lacZ reporter (which forms white colonies), and screened for mutant blue colonies. Conversely, we screened for activators by using a Pchb-lacZ Δcbp strain (which forms blue colonies) and isolated mutant white colonies. A positive control for the repressor screen was cbp, while a positive control for the activator screen was chiS. As expected, both of these genes were identified in their respective screens, which helped to validate this approach. No additional hits were identified in the repressor screen, but one hit that we identified in the activator screen was VC0214, the gene encoding the nucleoid occlusion protein SlmA . Consistent with a role in transcriptional activation of this locus, Pchb is no longer induced in a ΔslmA Δcbp mutant strain (cbp transcript levels in WT and ΔslmA cells when induced with (GlcNAc)6, the chitin hexasaccharide. While this locus is strongly induced in the WT, there is little to no activation in the slmA mutant, which is consistent with what we observed when using a cbp mutant to artificially activate this locus . Using 5’ RACE, we mapped the transcription start sites (TSS) for the chb transcript and found that there are two distinct TSSs for this locus. One TSS representing a longer transcript that is basally expressed in the absence of chitin induction (uninduced transcript), and a second TSS for a shorter transcript that is expressed when this locus is induced by chitin oligosaccharides or artificially induced by deletion of cbp (induced transcript) . Using primers that are specific to the longer uninduced transcript, we find that levels of this transcript are induced only 2 to 4-fold when cells are grown in the presence or absence of (GlcNAc)6 in both WT and ΔslmA cells, indicating that SlmA is specifically playing a role in the activation of this locus at the downstream TSS .To study ChiS regulation of the chb operon encodes an ABC transporter that mediates uptake of chitobiose (permease encoded by VC0618-0619). V. cholerae, however, can also degrade chitobiose in the periplasm into the chitin monomer GlcNAc [chb operon and whose regulation is independent of ChiS  [Fig 1C). However, upon deletion of both the chitobiose and GlcNAc transporters, cells are unable to grow on chitobiose as the sole carbon source . To determine if SlmA-dependent gene regulation of Pchb was physiologically relevant, we assessed whether SlmA was required for growth on chitobiose ((GlcNAc)2) as a sole carbon source. When we performed this analysis, we found that SlmA was required for growth on chitobiose in a ΔVC0995 mutant background, which is consistent with SlmA playing a physiologically important role in regulating the chitobiose transporter in the chb operon. As expected, all strains grew equally well when glucose or tryptone was provided as the sole carbon source whereas any strains with a ΔVC0995 mutation were unable to grow on GlcNAc . A slmA mutant, however, does not phenocopy a chiS mutant. In fact, a ΔchiS strain is unable to grow on (GlcNAc)2 as the sole carbon source, which is not surprising since this regulator is required for the expression of a number of chitin catabolic genes in addition to the chb operon [S3 Fig).The r GlcNAc , 9, 10, E. coli [E. coli and V. cholerae . To determine if gene regulation by SlmA is a conserved property, we replaced the native copy of slmA in V. cholerae with the gene for slmA from E. coli. We found that Ec SlmA was able to activate transcription of Pchb, albeit, less effectively than Vc SlmA, suggesting that this activity is not unique to the Vc homolog, but is a conserved property of this protein .SlmA function has primarily been characterized in E. coli –15. ThisslmA was responsible for the phenotypes observed on the Pchb operon, we complemented slmA on a plasmid. As expected, we found that ectopic expression of SlmA was sufficient to recover activation of Pchb in a ΔslmA Δcbp mutant . As seen previously [chb .To further confirm that deletion of eviously , expresschb based on the SBS consensus sequence of E. coli SlmA  [in vitro whole genome binding analysis that identified putative SBSs in V. cholerae [SlmA is a TetR family protein and binds to a specific and well-defined sequence . Through . This wacholerae .chb that contains the putative SBS. Indeed, we found that SlmA could directly bind to Pchb . SlmA was previously shown to bind to an SBS as a dimer-of-dimers [chb promoter probe . As a negative control, we tested binding of SlmA to PnanH, an unrelated promoter that lacks a predicted SBS. As expected, we found that SlmA could not bind to this probe . To determine if the putative SBS in Pchb is responsible for the shift observed in our EMSAs, we mutated 6 highly conserved residues in the SBS. We found that SlmA no longer shifted the Pchb probe when these residues were mutated, suggesting that this sequence represents a bona fide SBS in the chb promoter .To determine if SlmA could bind to this site, we performed electrophoretic mobility shift assays (EMSAs) using a probe from Pf-dimers , which ichb promoter, we next wanted to determine if this site was important for the transcriptional activation of this locus. To that end, we assessed activation of Pchb in a mutant strain where we mutated the SBS. When the SBS was mutated, we no longer observed activation of Pchb . Promoter truncations of Pchb also confirmed that the SBS in this promoter was required for activation of this locus . Furthermore, we replaced the native SBS in the Pchb promoter with the consensus SBS for E. coli SlmA. This alternative SBS was able to support activation of the locus . Cumulatively, these results suggest that SlmA recruitment to the Pchb promoter via an SBS is required for transcriptional activation. Our previous results indicated that Ec SlmA supported activation of Pchb less effectively than Vc SlmA . To determine if this was due to a reduced affinity of Ec SlmA for the SBS in Pchb, we tested activation of Pchb containing the consensus Ec SBS in a strain expressing Ec SlmA at the native locus. This strain, however, had similar levels of Pchb induction compared to a strain with Ec SlmA and the native SBS at Pchb . Thus, reduced activation of Ec SlmA is likely due to a reduced ability to promote transcriptional activation and not due to reduced affinity for the SBS.Since we found that SlmA could directly bind to an SBS in the chb operon for optimal growth on chitobiose . Deletion of SlmA, however, may have pleiotropic effects. To determine if SlmA binding at the SBS in Pchb was responsible for this effect, we mutated the SBS in the native chb promoter and tested growth on chitobiose. We found that mutating the SBS in Pchb prevents growth on chitobiose in the ΔVC0995 background, which is consistent with this SBS being critical for regulating the chitobiose transporter in the chb operon . These results indicate that SlmA-dependent regulation of Pchb is both direct and physiologically relevant.Above, we showed that SlmA was required for regulation of the chitobiose ABC transporter encoded by the chb. So, we mutated residues in SlmA involved in DNA binding , FtsZ interaction , and dimerization (R173) . We then tested these mutants for their ability to activate expression of Pchb. We also assessed whether these SlmA variants could bind to DNA using a previously described synthetic SBS-gfp reporter where SlmA binding at a high affinity SBS represses expression of GFP [E. coli instead of the native SBS found in Pchb because SlmA binds to this SBS with a higher affinity, allowing for a more sensitive evaluation of DNA binding . We also tested the nucleoid occlusion activity of these SlmA variants by overexpressing each and assessing the morphology of cells. Overexpression of WT SlmA results in a dramatic filamentous phenotype due to excess nucleoid occlusion activity, whereas alleles deficient in nucleoid occlusion do not cause this phenotype [S9 Fig). We also triple-FLAG tagged all SlmA alleles tested at the native locus to assess their expression levels by western blot analysis. This analysis demonstrated that all mutant alleles were expressed at least at WT levels . Importantly, SlmA-triple FLAG constructs were still functional for Pchb activation .The ability of SlmA to mediate nucleoid occlusion is dependent upon its ability to dimerize, bind DNA, and interact with FtsZ. The residues involved in these interactions have been previously characterized –14. We nn of GFP . We usedhenotype .chb, although one mutant (R71D) displayed a reduced degree of activation . As expected, these SlmA variants were still able to bind DNA  and had lost nucleoid occlusion activity . These results suggest that the nucleoid occlusion activity of SlmA is not required for its ability to act as a transcriptional activator of the chb operon.First, we found that the FtsZ binding mutants (F63A and R71D) were still able to facilitate transcriptional activation of PE. coli [chb. The T31A mutant was still able to bind DNA as indicated by the synthetic SBS-GFP reporter . The E43A mutant, however, was not able to bind DNA in this assay . Our previous results with the mutated SBS would suggest that SlmA binding to Pchb is required for activation, therefore, we hypothesized that the E43A mutant may still have the ability to bind an SBS, however, this may be below the limit of detection of our DNA binding reporter. To test this, we purified SlmA E43A and assessed its ability to bind Pchb in an EMSA. Indeed, we found that SlmA E43A was still able to bind to this promoter, although with a greatly reduced affinity compared to WT SlmA or SlmA T31A . Thus, these results indicate that while these mutations may eliminate DNA binding of Ec SlmA [Vc SlmA. Overexpression of these DNA binding mutants indicated that they had lost the ability to mediate nucleoid occlusion .Next, we tested DNA binding mutants T31A and E43A that were previously identified in E. coli , and, su Ec SlmA , they machb. As expected, this variant had also lost the ability to bind DNA  and mediate nucleoid occlusion .Dimerization of SlmA is required for DNA binding as well as nucleoid occlusion . Dimerizchb. To that end, we performed error-prone PCR of SlmA and screened for colonies that had lost the ability to activate Pchb-lacZ. We then counter-screened colonies for the ability to bind DNA using our synthetic SBS-GFP reporter. We screened >15,000 colonies and identified ~500 colonies that were deficient for activation of Pchb-lacZ. In our counter-screen, however, we found that none of these SlmA mutants maintained the ability to bind DNA, suggesting that DNA binding and transcriptional activation of Pchb are tightly linked. One variant identified in this screen was SlmA E43K, which is the same residue previously identified as playing a role in DNA binding [chbin vitro. Indeed, by EMSA we found that E43K could not bind Pchb . Importantly, this mutation does not simply result in an unstable protein because the E43K mutant still makes WT levels of SlmA protein . This mutant also lost the ability to mediate nucleoid occlusion when overexpressed . Cumulatively, these results indicate that high affinity binding of SlmA to DNA may be required for proper nucleoid occlusion, while weak binding is sufficient to mediate transcriptional activation of Pchb.Next, we sought to identify residues that are required for SlmA-mediated activation of P binding , howeverchb requires activation of ChiS, either via deletion of cbp or induction with chitin oligosaccharides. To determine if SlmA is the sole activator that acts downstream of ChiS we decided to overexpress SlmA to see if it was sufficient to activate Pchb in the absence of either inducer. Overexpression of WT SlmA results in a dramatic filamentous phenotype . Interestingly, overexpression of WT SlmA from a chromosomally integrated Ptac-SlmA construct still induced Pchb expression in a Δcbp-dependent manner, even at levels that induced morphological defects (10 μM), although activation was lost when cultures were induced with 100 μM IPTG . Above, we show that the nucleoid occlusion mutants T31A, F63A, and R71D are still able to activate Pchb , while their overexpression does not result in filamentation . Indeed, overexpression of SlmA F63A from a chromosomally integrated Ptac-SlmA F63A construct still supported activation in a Δcbp-dependent manner, however, induction with 100 μM IPTG still prevented activation similar to what was observed with overexpression of WT SlmA . Also, this was independent of any obvious morphological defect, suggesting that overexpression of SlmA at this level prevents Pchb activation independent of its role in nucleoid occlusion .Thus far, our data suggest that activation of Pchb activation, we overexpressed SlmA T31A, F63A, and R71D mutants with 10 μM IPTG and assessed whether we could activate Pchb under non-inducing conditions (i.e. in a cbp+ strain grown in the absence of chitin oligosaccharides). Overexpression of these constructs, however, did not result in activation under non-inducing conditions . Cumulatively, these results indicate that the repression of ChiS by CBP plays a dominant role over the activity of SlmA as a transcriptional activator of Pchb. This is consistent with a model whereby another, as yet unidentified, protein acts downstream of ChiS and is required for coactivation of this locus along with SlmA.Next, to determine if SlmA was sufficient to mediate Pchb activation, we decided to test whether this phenotype was the result of excess SlmA protein, which titrated away a putative co-activator from the Pchb operon. To that end, we overexpressed ChiS, which might enhance the expression / activity of this putative downstream coactivator. Interestingly, ChiS overexpression at 10–1000 μM also prevented Pchb activation, albeit, not to the degree of SlmA F63A overexpression . ChiS overexpression, however, did not rescue the loss of Pchb activation when SlmA F63A was overexpressed . The lack of increased Pchb expression over the native levels in the presence of higher levels of ChiS and/or SlmA indicate that neither protein is limiting and that native levels of these proteins are sufficient for maximal activation of Pchb.Since overexpression of SlmA F63A with 100 μM IPTG inhibited PThe transcriptional regulator cAMP receptor protein (CRP) binds to a specific DNA sequence in the presence of cAMP, allowing for activation or repression of target genes. One turn of B DNA is approximately 10 bp, and moving the CRP binding site in increments of 10 bp proximally or distally from the promoter maintains transcriptional activation of genes regulated by CRP by maintaining the helical phase of the CRP binding site in relation to the promoter , 19. Conchb by a similar mechanism. We found that moving the SBS as little at 3 bp or as much as 12 bp resulted in a significant decrease in Pchb activation . Constructs in which the SBS was moved to maintain the helical phase (i.e. in increments of 10 bp) were also unable to promote robust activation, indicating that SlmA may not activate transcription by direct contact with RNA polymerase in a mechanism similar to CRP . This is in line with the distance of the SBS from the TSS, 167 bp, which makes it unlikely that SlmA bound to its SBS directly interacts with RNAP to activate transcription. This supports the hypothesis that a coactivator may promote SlmA-dependent activation of Pchb.We tested whether there was helical phase-dependence for SlmA-mediated activation of Pchb in the region between the SBS and promoter element for the chitin-induced TSS. We hypothesized that if a coactivator was required for expression, there may be sequence specificity within this region, while if no coactivator was required, maintaining the spacing between the SBS and TSS may be sufficient to support activation. We tested this by replacing the region between the SBS and the -35 signal with the same number of nucleotides from an intragenic region of the lacZ gene . The activation of this reporter was abolished after swapping the native sequence with lacZ sequence , indicating that sequence specificity within this region may be required for activation. Furthermore, truncation studies indicated that elements necessary for activation include the SBS, TSS and intervening regions . Also, a bioinformatically determined hairpin exhibits some inhibitory activity . Together, these data are consistent with a model whereby the region between the SBS and TSS contains a coactivator-binding site required for Pchb induction. There are, however, a number of alternative models that can account for the results obtained, which are addressed in the Discussion.Next, we assessed whether there was sequence specificity for activation of Pchb. To test this, we inactivated every non-essential sigma factor  in V. cholerae and assessed transcriptional activation of Pchb. We observed activation in all mutant strains, suggesting that the essential housekeeping sigma factor RpoD likely plays a dominant role in Pchb expression . The degree of Pchb activation, however, was reduced in the rpoE mutant, suggesting that this sigma factor either plays a minor role in activation of this locus or that RpoE is required for proper expression or folding of the GFP reporter. Additionally, we observed an increase in expression in the rpoS mutant, suggesting that this sigma factor may play a role in the expression of some upstream repressor of this locus.Finally, we were interested in determining which sigma factor was required for activation of PV. cholerae. We show that this regulation is direct because SlmA binds to Pchb, the promoter that regulates these genes. Furthermore, we demonstrate that this direct regulation is required for growth of V. cholerae on chitobiose.Here, we show that the cell division licensing factor SlmA is required for activation of chitobiose utilization genes in chb, the exact mechanism of activation is still unclear. We have shown that the location of the SBS within Pchb is important for activation. Additionally, we show that the sequence between the SBS and the -35 sequence is required for activation, consistent with this region harboring a coactivator binding site. Because the SBS is located 167 base pairs away from the TSS, it is unlikely that SlmA directly interacts with RNA Polymerase to mediate activation. We also explored the possibility that SlmA binding may occlude a repressor binding site thereby allowing for activation of Pchb indirectly. We formally tested this hypothesis by performing a Tn mutant screen for repressors in a ΔslmA Δcbp Pchb-lacZ reporter strain, however, this screen did not identify any putative repressors . As a result, we propose a mechanism in which SlmA requires a presently unknown ChiS-dependent coactivator for activation of Pchb . A candidate for this unknown factor was not identified in the Tn-mutant screen for activators that identified SlmA. This screen, however, may not have been saturating or, alternatively, this factor may be essential for viability or masked by genetic redundancy.While we show that SlmA is a direct activator of Pchb [Fig 5). Alternatively, it is possible that the coactivator loops, bends, or structurally alters DNA in the chb promoter to enhance SlmA-dependent recruitment of RNA polymerase. Identifying and characterizing a putative coactivator for Pchb that acts downstream of ChiS will be the focus of future work.Because ChiS is an orphan sensor kinase, the most logical coactivator would be a cognate response regulator. However, it has been shown that the conserved histidine and aspartate residues of ChiS that are critical for autophosphorylation and phosphorelay activity, respectively, are not important for activation of Pchb . Thus, wchb abolishes activation . If DNA spreading were critical we would predict that this perturbation should have a limited effect on transcriptional activation. Further, we have shown that overexpression of SlmA can actually decrease Pchb transcription, which again would not be predicted by a DNA spreading model . Alternatively, it is possible that increased SlmA spreading (when SlmA is overexpressed) may prevent RNA polymerase from binding, resulting in decreased expression of Pchb.Another possible mechanism of activation is that SlmA directly interacts with RNA polymerase through its ability to polymerize and spread on DNA . We havechb locus. The short transcript is strongly induced in the presence of chitin oligosaccharides, while the long transcript is basally produced in the absence of chitin and is only modestly induced in its presence (2–4 fold). This indicates that there may be an additional level of regulation involving the 5’ untranslated region (UTR) of the chb transcript. A putative hairpin in Pchb also seemed to have some inhibitor effect on Pchb expression . It is possible that the long transcript and hairpin play an important role in the basal expression of the chb operon, which is required for a rapid response to chitin oligosaccharides.We have shown that there are two transcriptional start sites for the Vibrio harveyi  [chb may be a mechanism to integrate the cell division status of the cell with activation of chitin utilization.TetR proteins are generally transcriptional repressors, however, there are examples where these proteins can mediate transcriptional activation. One well-studied example is the LuxR protein in nificus) , 22. Slmnificus) . Their rnificus) . One sucnificus) . Thus, Sin vitro whole genome binding analysis identified 79 putative SBSs in V. cholerae [chb), while only ~12% of the genome constitutes intergenic sequence. By contrast, in E. coli, ~8.3% (2/24) of SBSs are in intergenic sites [Bacillus subtilis, ~12% (9/74) of binding sites are intergenic [matS sites) ~24% (6/25) are in intergenic regions [matS sites in E. coli and SBSs in V. cholerae) is because these sites are more flexible in regards to accommodating mutations compared to coding regions of the genome. Alternatively, SBSs may be enriched in intergenic sites in V. cholerae possibly to regulate the expression of additional genes. This will be the focus of future work. Preliminary RNA-seq analysis of a slmA mutant grown in rich medium, however, did not uncover any additional loci regulated by SlmA in V. cholerae. An effect of SlmA on gene regulation at Pchb, however, was only uncovered by deletion of cbp or growth on chitin (both of which induce this locus). Thus, it may be that additional inducing cues are required to uncover a role for SlmA-dependent regulation at additional genetic loci. Nucleoid occlusion factors are present across diverse bacterial genera and it is tempting to speculate that these proteins may also participate in regulating gene expression in addition to their established roles in cell division licensing.To our knowledge, this is the first example of gene regulation by a nucleoid occlusion protein. A recent cholerae . We detetergenic  and ~11% regions . Thus, iV. cholerae strains were routinely grown in LB medium and on LB agar supplemented when necessary with Carbenicillin (20 or 50 μg/mL), Kanamycin (50 μg/mL), Spectinomycin (200 μg/mL), Trimethoprim (10 μg/mL), and/or Chloramphenicol (2 μg/mL). For growth on a defined carbon source, strains were grown in M9 minimal medium containing the indicated carbon source.R mini-Tn10 transposon exactly as previously described [chb-lacZ or Pchb-lacZ Δcbp reporter strain. Transposition was induced by plating cultures on Carb containing media at 42°C. To screen colonies, plates also contained 40 μg/mL X-gal and 5 mM IPTG. IPTG was added to competitively inhibit the basal activity of the Pchb-lacZ reporter. This made it easier to distinguish colonies where the locus was uninduced (e.g. Pchb-lacZ) from when the locus was induced (e.g. Pchb-lacZ Δcbp).Transposon mutant libraries were generated with a Carbescribed . BrieflyS1 Table and S2 Table, respectively.The parent strain used in this study was E7946 . Mutant 600 of 1.0. Fluorescence was determined using a BioTek H1M plate reader with excitation set to 500 nm and emission at 540 nm.GFP was measured exactly as previously described . BrieflyS2 Table.The 5’ end of transcripts were mapped using the SMARTer 5’/3’ RACE kit according to manufacturer’s instructions (Takara). The primers used are listed in 600 on a BioTek H1M plate reader.Cells were grown in M9 minimal medium in the presence of 0.2% of the indicated carbon source at 37°C. Growth was kinetically monitored by measuring ODS2 Table) and the reaction was monitored on a Step-One qPCR system.RNA was isolated from cells using an RNEasy minikit according to manufacturer’s instructions (Qiagen). RNA was reverse transcribed using AffinityScript QPCR cDNA Synthesis Kit (Agilent). Quantitative PCR was performed using iTaq Universal SYBR Green Supermix (Bio-Rad) with primers specific for the genes indicated  or mouse monoclonal α-RpoA (Biolegend) primary antibodies. Blots were then incubated with α-rabbit or α-mouse secondary antibodies conjugated to IRdye 800CW as appropriate and imaged using an Odyssey classic LI-COR imaging system.Cells were grown to mid-log in LB medium and then mounted on 1% agarose pads. Cells were imaged using an Olympus IX83 phase contrast microscope.S1 Figchb transcript. Data are shown as the mean ± SD and are from at least three independent biological replicates. Fig 1D shows the qRT-PCR data for the short “induced” transcript.Transcript levels were determined via qRT-PCR using primers specific for the long P(PDF)Click here for additional data file.S2 FigA) glucose, (B) tryptone, or (C) N-acetylglucosamine (GlcNAc) as a sole carbon source.Growth curves of the indicated strains in M9 minimal medium containing 0.5% ((PDF)Click here for additional data file.S3 FigA) chitiobiose, (B) N-acetylglucosamine, (C) glucose, or (D) tryptone as a sole carbon source.Growth curves of the indicated strains in M9 minimal medium containing 0.5% ((PDF)Click here for additional data file.S4 FigResidues highlighted in black are identical and those highlighted in gray are similar. Arrows indicate residues used in site-directed mutational analysis. T31 and E43 are involved in DNA-binding, F63 and R71 are involved in interaction with FtsZ, and R173 is involved in dimerization.(PDF)Click here for additional data file.S5 Figchb fused to gfp to serve as a transcriptional reporter. Also, strains are either intact for cbp (+) or harbor a cbp deletion (Δ). Data are shown as the mean ± SD and are from at least three independent biological replicates.Promoters that were predicted computationally (Softberrry BPROM), are indicated as P1, P2, and P3, as well as a putative hairpin. All strains harbor the indicated region of P(PDF)Click here for additional data file.S6 Figchb-gfp transcriptional reporter. “Ec SBS” indicates that the native SBS sequence in Pchb was swapped for the consensus SBS binding site for Ec SlmA. Data are from at least three independent biological replicates and shown as the mean ± SD. Please note that data from the first four bars is identical to that shown in Fig 1E, and are included here to allow for easy comparison to the additional samples included in this figure.GFP fluorescence was measured in the indicated strains, all of which contain a P(PDF)Click here for additional data file.S7 FigA) EMSAs performed where the indicated SBS sequence containing probes were incubated with (from left to right): 0 nM, 7.5 nM, 15 nM, 30 nM, 60 nM, 120 nM, and 240 nM of purified SlmA. Data are representative of at least two independent experiments. (B) Labeled Vc SBS from Pchb (0.2 nM) was incubated with increasing concentrations of unlabeled Vc SBS from Pchb or the consensus SBS from E. coli. Concentrations of unlabeled “cold competitor” probe from left to right are: 0 nM, 0.2 nM, 0.4 nM, 0.8 nM, 1.6 nM, 3.2 nM, 6.4 nM, 12.8 nM, 25.6 nM, 51.2 nM, and 200 nM.((PDF)Click here for additional data file.S8 FigA) Fluorescence measurement of strains containing a Pchb-mcherry reporter. Strains with slmA-3x FLAG were at the native slmA locus. Data indicates that the SlmA-3x FLAG construct is functional for Pchb activation. (B) SlmA site directed-mutants were engineered to contain a 3x FLAG tag at the native locus. Cell lysates of these strains were subjected to Western blot using a FLAG specific antibody and separately with an antibody to RpoA, which served as a loading control.((PDF)Click here for additional data file.S9 Figtac promoter were grown in the presence or absence of 100 μM IPTG and imaged by phase contrast microscopy. Data are representative of at least two independent experiments.Strains containing chromosomally integrated constructs for overexpression of the indicated SlmA mutants via an IPTG-inducible P(PDF)Click here for additional data file.S10 FigA-D) GFP fluorescence was assessed in strains that harbor a Pchb-gfp reporter as well as a chromosomally integrated construct for overexpression (Ptac-X) of ChiS and/or SlmA as indicated. All strains were grown with the indicated concentration of IPTG. Ptac-slmA constructs contained a C-terminal triple FLAG tag, while Ptac-chiS constructs were untagged because the FLAG tag diminished the activity of this protein. Data are shown as the mean ± SD and are from at least three independent biological replicates. (E) Representative western blots of SlmA-3x FLAG and ChiS-FLAG overexpression using the same chromosomally integrated constructs used in A-D for SlmA and a ChiS-FLAG construct. (F) Representative phase contrast images to show the morphology of cells ectopically expressing WT SlmA or SlmA F63A using the same mutant constructs and concentrations of IPTG used in A-D.((PDF)Click here for additional data file.S11 Figchb-gfp reporter as well as the other mutations indicated. All strains were grown with 10 μM IPTG to overexpress the indicated SlmA mutant proteins. Consistent with a model where SlmA requires another coactivator, overexpression of these SlmA variants was not sufficient to induce expression of Pchb. Data are shown as the mean ± SD and are from at least three independent biological replicates.GFP fluorescence was assessed in strains that harbor a P(PDF)Click here for additional data file.S12 Figchb-gfp reporter with the indicated mutation to move the SBS sequence as indicated. Each SBS mutation was tested in a background where cbp is intact (+) or deleted (Δ). Data are shown as the mean ± SD and are from at least three independent biological replicates.All strains harbor a P(PDF)Click here for additional data file.S13 Figchb-gfp reporter with the indicated sigma factor deleted. Data are shown as the mean ± SD and are from at least three independent biological replicates.GFP fluorescence was assessed in strains that harbor a P(PDF)Click here for additional data file.S1 Table(PDF)Click here for additional data file.S2 Table(PDF)Click here for additional data file.


We show here that nucleosome depletion regions (NDRs) form in both proximal promoters and distal enhancers during NSCs differentiating into neurons in the early Drosophila embryonic development. NDR formation in the regulatory regions involves nucleosome shift and eviction. Nucleosome occupancy in promoter NDRs is inversely proportional to the gene activity. Genes with promoter NDR formation during differentiation are enriched for functions related to neuron development and maturation. Active histone-modification signals (H3K4me3 and H3K9ac) in promoters are gained in neurons in two modes: de novo establishment to high levels or increase from the existing levels in NSCs. The gene sets corresponding to the two modes have different neuron-related functions. Dynamic changes of H3K27ac and H3K9ac signals in enhancers and promoters synergistically repress genes associated with neural stem or progenitor cell-related pluripotency and upregulate genes associated with neuron projection morphogenesis, neuron differentiation, and so on. Our results offer new insights into chromatin remodeling during in vivo neuron development and lay a foundation for its epigenetic regulatory mechanism study of other lineage specification.Neurons are a key component of the nervous system and differentiate from multipotent neural stem cells (NSCs). Chromatin remodeling has a critical role in the differentiation process. However, its  The abnormal epigenomic changes disrupt genes that function in neural development and consequently lead to neurological and psychiatric diseases. Therefore, it is a prerequisite to characterize the in vivo chromatin states of neurons and NSCs for the analysis of epigenomic changes during NSCs differentiating into neurons.Neurons are a major and important cell type of the nervous system. Neurons differentiate from multipotent precursors called neural stem cells (NSCs). This indicates that an extensive epigenomic change occurs during differentiation.in vitro cell differentiation platforms. Overall, differentiated cells have a more condensed chromatin structure than stem cells.2 A bunch of studies showed changes in histone-modification patterns in embryonic stem cell (ESC) differentiation.5 A recent study profiled chromatin state dynamics across 16 stages of hematopoietic differentiation.6 Global chromatin states of mouse ESCs, neural progenitor cells, and embryonic fibroblasts were reported, as well as new bivalent chromatin marks.7 Both Foxa2 and H2A.Z mediate nucleosome depletion during ESCs differentiating into the endoderm.8 A recent study identified 85 core HM sites in mouse ESCs and 78 sites in neural progenitor cells.9 To date, there is no study reported focusing on chromatin dynamics during in vivo neuron development.There are many studies on the role of chromatin remodeling in cell fate specification using in vivo cell types to investigate epigenetic reprogramming during differentiation because cell cultures or in vitro differentiated cells lack the proper in vivo context. However, it is a technical challenge to isolate specific neurons and NSCs for epigenomic studies that require a large number of cells. Manual sorting10 is laborious and only useful for collecting a small number of cells. Laser capture microdissection11 is also not suitable for isolating large number of cells required by chromatin assays. Although fluorescence-activated cell sorting (FACS) can isolate large number of cells, it requires molecular markers specific to the cell type of interest that are not always available. FACS requires harsh treatment of the tissue and does not work efficiently in complex tissues such as the brain. As such, FACS is not optimal for genome-wide epigenomic profiling. Owing to an alternate approach, INTACT (isolation of nuclei tagged in specific cell types),12 it introduces into Drosophila genome a nuclear targeting fusion (NTF) gene consisting of 3xFLAG, BLRP , mCherry, and RanGap (expressed in the cytoplasm and outer nuclear envelope). The NTF gene is expressed under the control of a certain cell-specific gene promoter. Therefore, INTACT can capture cell-type-specific nuclei suitable for gene expression, epigenomic, and proteomic profiling within a tissue through affinity purification of the NTF protein.15 It allows the study of distinct cell types at different stages of development.It is critical to use specific Drosophila to isolate NSCs and neurons for their epigenome profiling including transcriptome, genome-wide nucleosome occupancy, and global core HM signals. Epigenomic analyses show that nucleosome depletion regions (NDRs) form in both promoters and enhancers through nucleosome shift and eviction. Nucleosome occupancy in promoter NDRs is inversely proportional to the gene activity. Genes with promoter NDR formation during differentiation are enriched for neuron development and maturation. Active H3K4me3 and H3K9ac are deposited in the promoters of genes in neurons related to neuronal functions in two modes: de novo establishment or increase from existing levels in NSCs. In addition, changes of H3K27ac and H3K9ac in promoters and enhancers synergistically upregulate genes with functional enrichment for neuron differentiation and downregulate genes with functional enrichment for neural progenitor cell-related pluripotency.Here we applied INTACT in 15 method to isolate neuronal nuclei, we generated a Drosophila strain containing Elav-Gal4>UAS-NTF (Elav>NTF). Elav is the commonly used neuronal marker gene.17 The neuronal nuclei were then affinity purified from stage 15 to 16 embryos (12–14 h AEL) using anti-Flag-coated magnetic beads . The promoter of NSC marker gene, Sca,18 drives the specific expression of NTF gene in the nuclear envelope of NSCs. Therefore, NSC nuclei were collected from stage 11 embryos (5–7 h AEL) through affinity purification. Moreover, the affinity-purified neuronal and NSC nuclei reached a purity of 98±2% and 90±3%, respectively  (To apply the INTACTic beads . Similarts each) .in situ hybridization by Berkeley Drosophila Genome Project.19 The results show that expression levels of neuronal genes are significantly higher compared with other tissue-specific genes in neuronal nuclei  and calculated read count in each window as the nucleosome occupancy. Nucleosome occupancy changes spread out across the genome during differentiation and are statistically significant . We obta20 and obtained 523 516 nucleosomes in NSCs and 514 910 nucleosomes in neurons. The majority of nucleosomes (>92%) shift by 1 to 126 bp and only ~2% of nucleosomes do not shift during differentiation. In total, 5.3% of nucleosomes are lost in neurons, whereas 4.5% of nucleosomes are newly formed in neurons  is higher in NSCs than in neurons . It has n levels . Interesn levels . Consistn levels . GO termn levels . In contn levels . This sun levels . That is21 To understand how HMs alter and the resultant influence on gene expression during the differentiation, we first profiled HM signals in promoter regions (±1 kb of TSS). H3K4me3 levels in promoter regions take on a bimodal distribution   . A recenmmitment . This inTo understand how HMs change in promoters during the differentiation, we categorized promoters by HM content in neurons transited from promoters with a certain HM state in NSCs. The results show that the majority of promoters maintain their HM state during differentiation . We furtde novo established in neurons. It is unclear if these two modes are associated with different functions. To answer this question, we examined HM changes in the promoters of different gene sets classified by GO functional annotations. Intriguingly, genes associated with functions related to neuron development gain H3K4me3 and H3K9ac in promoters upon differentiation by increase from their existing levels in NSCs , poised (H3K4me1+ and H3K27ac−), and off (H3K4me1−). Approximately 65% of enhancers are active in both cell types . As expeK-means (K=3) revealed three patterns of NDR formation: nucleosome eviction and nucleosome shift to 5′ or 3′ direction  is initiated earlier and can indicate the differentiation potential of progenitor cells earlier than gene expression profiles during blood formation.6 It will be interesting to know whether openness within enhancers (measured by NDR) can serve as an indicator of lineage commitment during in vivo neural development in Drosophila. This goal will be achieved when clean marker genes for distinct intermediate cell types during NSCs differentiating to a wide variety of mature neural cells are available.Activity control of enhancers regulates cell-type-specific patterns of gene expression and has a critical role in development.egulated . This imDrosophila. NSCs differentiating to glial cells require low levels of H3K9ac. High levels of CREG-binding protein (a histone acetyl-transferase) result in high levels of H3K9ac and disrupt gliogensis in Drosophila embryonic neural development.34 In contrast, our results show that H3K9ac signals in promoters are increased in neuronal cells compared with NSCs. This is consistent with the observation that neurons have relatively high levels of H3K9ac compared with most glial cells.34 This implies that HM contents in promoters regulate the expression of genes critical for neural cell lineage specification. Intriguingly, we further revealed that the high levels of active H3K4me3 and H3K9ac come from both de novo deposition in neurons and increase from existing levels in NSCs 15 was kindly provided by Professor Steven Henikoff . This stock expresses the transgene (NTF) 3xFLAG-BLRP-mCherry-RanGap and BirA under GAL4 driver. Other lines such as Elav-Gal4;Tm3/Tm6B , Sca-Gal4 (II) and sp/cyo;Dr./Tm6B  were obtained from the Bloomington Stock Center .The transgenic line Elav-Gal4;Tm3/Tm6B virgin flies were crossed with w1118;p[UASRG]6 male flies to generate the F1 offspring Elav-Gal4/+p[UASRG]6/Tm3. The F1 male flies were backcrossed with Elav-Gal4;Tm3/Tm6B virgin flies to generate the F2 offsprings Elav-Gal4;p[UASRG]6/Tm3  and Elav-Gal4/+p[UASRG]6/Tm3 . Then, the F2 offsprings were self-crossed to generate the F3 offsprings that are homozygous, Elav-Gal4;p[UASRG]6  and Elav-Gal4/+p[UASRG]6 . The embryos of F3 flies were collected by self-crossing to isolate neurons.Sca-Gal4 virgin flies and sp/cyo;Dr/Tm6B male flies were crossed to generate the F1 offspring. We retained the ones with genotype Sca-Gal4/cyo;+/Tm6B (denoted as F1A) for the next cross. The sp/cyo;Dr/Tm6B virgin flies were crossed with w1118;p[UASRG]6 male flies to generate the F1 offspring. We retained the ones with genotype +/cyo;p[UASRG]6/Tm6B (denoted as F1B) for the next cross. Next, F1A virgin flies were crossed with sp/cyo;Dr/Tm6B male flies to produce the F2 offspring. We retained the ones with genotype Sca-Gal4/cyo;Dr/Tm6B (denoted as F2A) for the next cross. The sp/cyo;Dr/Tm6B virgin flies were crossed with F1B male flies to generate the F2 offspring. We retained the ones with genotype sp/cyo;p[UASRG]6/Tm6B (denoted as F2B) for the next cross. Later, the F2A virgin flies were crossed with F2B male flies to generate the F3 offspring. We retained the ones with genotype Sca-Gal4/cyo;p[UASRG]6/Tm6B (denoted as F3A) for the next cross. Finally, F3A flies were self-crossed to generate the F4 offspring. We retained the ones with genotype Sca-Gal4/cyo;P[UASRG]6 (denoted as F4A). The embryos of F4A flies were collected by self-crossing to isolate NSCs.First, The following antibodies were used for ChIP: H3K4me3 , H3K4me1 , H3K9ac , H3K27ac  and H3K27me3 . Antibodies used for immunostaining were as follows: rat-α-Elav  and goat anti-rabbit IgG H&L . Anti-Flag-coated M2 magnetic beads  were used for pull down.2HPO4, 1.4 mM NaH2PO4)+0.1% Triton X-100), and were washed with tap water to remove the yeast. Then, embryos were dechorionated with 50% hypochloric acid for 3 min and were crosslinked in a 1:3 mixture of ChIP–Fixed buffer (50 mM (pH 7.6) HEPES, 100 mM NaCl, 0.1 mM EDTA, 0.5 mM EGTA) with 1.8% formaldehyde and heptane for 15 min on a shaker with a speed of 300 r.p.m. The aqueous and organic phase was replaced with PBST containing 0.25 mM glycine to cease the crosslinking reaction. The fixed embryos were rinsed three times with PBST and then stored at −80 °C for future use.Embryos were collected on grape juice plates with a yeast paste from embryo collection cages for 2 h, and then aged at 25 °C for 3 and 10 additional hours. The harvested embryos are 5–7 and 12–14 h old, respectively. Embryos were transferred onto the mesh with PBST  Tris-HCl, 0.125 M sucrose, 0.5 mM spermidine, 0.15 mM spermine, EDTA, 0.5 mM EGTA, 1 × Complete protease inhibitor (PI)) and dounce homogenized. To monitor bead binding 0.5 ml of 5 mg/ml DAPI solution was added. The nuclei mixture was filtered through one layer of Miracloth into 50 ml conical tube and diluted to 40 ml with cold HB125 buffer; afterwards, 3 ml of OptiPrep (Sigma-Aldrich) was added and then centrifuged at 1000 × g for 10 min at 4 °C. The supernatant and OptiPrep cushion were discarded, leaving ~2 ml of HB125 containing nuclei concentrated at the interface. Isolated nuclei were suspended in HB125 with 60 μl of anti-Flag M2 magnetic beads slurry and incubated on a rotator for 2 h at 4 °C. Beads with affinity-bound nuclei were absorbed by the magnet and then washed three times using HB125. Purified nuclei were stored at −80 °C before proceeding with ChIP.Purification of tagged nuclei was performed using the INTACT technology as described previously.Embryos were dechorionated in 50% solution of bleach and fixed in a 1:1 mixture of 4% formaldehyde in PBS with 0.3% Tween-20 and heptane for 20 min on a shaker. The aqueous phase was discarded and replaced with methanol, and embryos were shaken for 3–5 min at 300 r.p.m. to burst vitelline membranes. Embryos were rinsed three times with methanol and then rinsed three times with PBS containing 0.3% Triton X-100. Fixed embryos were blocked with PBST supplemented by 5% normal donkey serum (NDS) for 30 min and then incubated with primary antibodies of various dilutions in PBST containing 5% NDS overnight at 4 °C. These embryos were washed as described above, followed by a 1–2 h secondary antibody incubation, and then were washed three times again to avoid nonspecific binding. These stained embryos were mounted on slides with additional 50% glycerol in PBS. Slides were examined on a Zeiss Imager M2 microscopy . To examine bead binding, DAPI-stained total and affinity-purified nuclei were counted on a hemacytometer, respectively.g for 5 min and suspended with 500 μl of 37 °C preheated MNase  digestion buffer (10 mM (pH 7.5) Tris-HCl, 15 mM NaCl, 60 mM KCl, 2 mM CaCl2, 0.15 mM spermine, 0.5 mM spermidine, 1 × PI) with 12 U MNase  and incubated at 37 °C for 20 min. The reaction was terminated on ice by adding EDTA to a final concentration of 10 mM for 10 min. The supernatant was discarded and the pellet was washed with A2 buffer  HEPES, 1 mM EDTA, 0.5 mM EGTA, 0.1% Triton X-100, 0.1% sodium deoxycholate, 1 × PI) and resuspended in A2 buffer with 0.1% SDS. The nucleosome pellet was dissolved through sonication with three cycles of 20 s duration with at least 40 s pauses between cycles at the power setting of 6 (out of 20) on a Misonix sonicator XL-2000 . The supernatants with chromatin were kept for the next ChIP assay, or were reverse crosslinked to harvest nucleosomal DNA fragments as follows: chromatin was treated with RNase A at 37 °C for 0.5–1 h and followed by proteinase K treatment at 65 °C for 2 h. Later, the nucleosomal DNA was retracted by phenol–chloroform and precipitated with a 1:10:100:200 mixture of 20 mg/ml glycogen, 3 M (pH 5.3) NaOAc, nucleosomal DNA mixture, and cold 100% ethanol.Affinity-purified nuclei were pelleted by centrifugation at 1000 × μg was used for each ChIP reaction with HM antibodies of various doses as described in the specifications. Mixture containing chromatin, antibody, and ChIP buffer (16.7 mM (pH 8.1) Tris-HCl, 167 mM NaCl, 1.2 mM EDTA, 1% Triton X-100, 0.01% SDS) was incubated overnight on a rotator at 4 °C. Then, 20 μl of ChIP-Grade Protein G Magnetic Beads  was added to each IP reaction. The mixture was incubated for 2 h with rotation. Afterwards, beads were washed three times with low salt wash buffer  Tris-HCl, 0.1% SDS, 1% Triton X-100, 150 mM NaCl) and once with high salt wash buffer  Tris-HCl, 0.1% SDS, 1% Triton X-100, 500 mM NaCl) for 5 min each wash. Beads were suspended in 150 μl of ChIP elution buffer (50 mM (pH 8.1) Tris-HCl, 10 mM EDTA, 0.1% SDS) at 65 °C for 45 min.Chromatin of 10–15 The purified mononucleosomal DNA was subjected to massively parallel DNA sequencing on Illumina HiSeq2000 platform  using a 49 bp single-end protocol.Drosophila transcripts (FlyBase r5.43) using TopHat (v.1.3.1) with default parameter setting.38 The uniquely mapped reads were assembled into transcripts guided by reference annotation with Cuffdiff (v.1.3.0)38 to calculate gene expression levels that were normalized as fragment per kilobase per million mapped fragments. The DEGs were identified with FDR <0.05.Nuclear RNA was isolated form tagged nuclei of the affinity-purified embryos without crosslinking using the RNeasy Micro Kit . Genomic DNAs were removed with Turbo DNA-Free Kit . The RNA sequencing libraries were constructed using standard Illumina libraries prep protocols. RNA-seq was performed on Illumina HiSeq2000 platform. Sequencing reads were aligned to the in situ database BDGP.19 Genes expressed at stages 11–12 and 13–16 in different tissue-specific cells were collected.Tissue-specific gene lists were downloaded from the Berkeley Drosophila Genome Project Drosophila reference genome (dm3) using Bowtie,39 allowing maximal two mismatches. The uniquely mapped reads were used to identify genome-wide nucleosome positions through the peak-calling tool GeneTrack20 that also calculated read count for each nucleosome. The read count was normalized by total uniquely mapped reads as the nucleosome occupancy. Nucleosome fuzziness was calculated as the standard deviation of the coordinates of all reads defining the same nucleosome as described previously.40 It measures how delocalized a nucleosome position is. Each nucleosome was assigned to either of promoter, genic, or intergenic regions depending on in which region the midpoint of the nucleosome was located.Nucleosomal sequencing reads were aligned to the 40 Briefly, Drosophila transcript annotation was downloaded from FlyBase release 5.43. Nucleosomes located within ±1 kb of TSSs were collected. Nucleosome length equals to the fuzziness value and centers at the nucleosome midpoint that defines the nucleosome position. The region that a nucleosome length spreads out has the nucleosome occupancy. The difference in nucleosome organization around TSSs was measured by subtracting nucleosome occupancy in NSCs from that in neurons for each site. The difference clustered by K-means (K=5) and plotted as heatmap.Nucleosome organization change around the TSS was analyzed as in the previous study.The original composite distribution of nucleosome around TSS of the five clusters of genes in NSCs and neurons, respectively, was calculated by aggregating nucleosomal read count at each distance relative to the TSS as follows: each read represents a nucleosome by extending toward 3′ end to a length of 147 bp. The midpoint of extended read defines the nucleosome position. We summed total read counts at each site within ±1 kb of TSSs for the five gene clusters, respectively. The nucleosome occupancy equals to the read count normalized as RPKM. We further binned the nucleosome occupancy by a 5-bp interval of nucleosome distance to TSS, and smoothed it with 5-bin moving average and 1-bin step size.K-means clustering (K=2).Sequencing reads were mapped to the reference genome similar to the above nucleosomal read mapping. We defined promoters as the region ±1 kb of TSSs. HM levels in a promoter were calculated as all reads within the promoter and normalized as RPKM. The density distribution of H3K4me1, H3K4me3 and H3K9ac levels in promoters is a bimodal pattern. The levels at the valley were chosen as thresholds, which was 11.6 for H3K4me1, 16 for H3K4me3, and 16 for H3K9ac. For qualitative analysis, promoters with a higher level of a certain HM compared with the threshold are characterized as marked by this HM, otherwise not. The density distribution of H3K27me3 levels in promoters is a normal distribution. We grouped promoters into two classes with or without H3K27me3 by The composite distribution of HM around TSS was calculated in the same manner as nucleosome distribution described above.6 In brief, H3K4me1-enriched regions (peaks) were identified using HOMER41 using a 1000-bp sliding window with a false discovery rate of 0.1%. In addition, two adjacent peaks need to be separated by at least 1000 bp to avoid redundant detection. We next combined such valid peaks from two cell types as one set of peaks by merging overlapped peaks. The peak with highest HOMER score replaces the overlapped peaks. Then, we retained total 14 234 H3K4me1 peaks for the two cell types. To remove false enhancers, we calculated H3K4me3 signals in H3K4me1 peaks and observed a bimodal distribution. A Two Gaussian mixture model was used to fit the distribution to select a threshold (=4.55). The H3K4me1 peaks with H3K4me3 levels higher than the threshold were discarded. It resulted in 7728 H3K4me1 peaks as the final set of enhancers.Enhancers and their chromatin states were determined as described previously.To define chromatin states for enhancers, we first recalculated H3K4me1 and H3K27ac signals in the enhancers for the two cell types. We also randomly selected the same number (7728) of 1000-bp genomic regions and calculated H3K4me1 and H3K27ac signals. The smallest level of the top 10% signals was used as the false-positive threshold, which was 5.5 for H3K4me1 and 4.5 for H3K27ac, respectively. Enhancers are ‘on' with H3K4me1 level higher compared with 5.5 and ‘off' otherwise. Of ‘on' enhancers, those are ‘active' with H3K27ac level higher compared with 4.5 and ‘poised' otherwise. Finally, each enhancer was associated with a single gene based on the nearest RefSeq TSS.The RNA-seq, MNase-seq, and ChIP-seq data sets have been deposited in the Gene Expression Omnibus under the accession number GSE80458.


Expression of protein-coding genes in eukaryotes relies on the coordinated action of many sophisticated molecular machineries. Transcription produces precursor mRNAs (pre-mRNAs) and the active gene provides an environment in which the pre-mRNAs are processed, folded, and assembled into RNA–protein (RNP) complexes. The dynamic pre-mRNPs incorporate the growing transcript, proteins, and the processing machineries, as well as the specific protein marks left after processing that are essential for export and the cytoplasmic fate of the mRNPs. After release from the gene, the mRNPs move by diffusion within the interchromatin compartment, making up pools of mRNPs. Here, splicing and polyadenylation can be completed and the mRNPs recruit the major export receptor NXF1. Export competent mRNPs interact with the nuclear pore complex, leading to export, concomitant with compositional and conformational changes of the mRNPs. We summarize the integrated nuclear processes involved in the formation and export of mRNPs. The product of a protein-coding gene that is delivered to the cytoplasm is an export competent mRNA–protein (mRNP) complex . An mRNPAfter release from the gene, an mRNP enters the interchromatin compartment and moves by diffusion mainly through the interchromatin channel network of diploid cells . Some prWe will outline the processes at the active gene, in the interchromatin and at the NPC that together result in the formation and export of functional mRNPs. It is clear that there is a close physical and functional coupling between transcription, processing, and assembly of pre-mRNPs and mRNPs. The mRNP obtains a processed structure and a collection of proteins that will influence the downstream events. The intricate physical and functional couplings are likely to have evolved, because they enhance coordination, efficiency, and regulation of the multiple steps of gene expression Fig. . We willThe cell nucleus contains several functional compartments which are maintained despite the lack of an enclosing membrane and the dynamic exchange of components with the surrounding nucleoplasm . CompartTranscribing genes may represent transient microenvironments favourable for mRNP formation Fig. . Upon geThe functional significance of chromatin unfolding and the establishment of a microenvironment for transcription and pre-mRNA processing need to be further investigated. The relative contribution of chromatin modifications, the transcription process itself, and the presence of transcripts must be clarified. We need specific examples of endogenous genes in living cells to learn the rules and variations that appear to exist in different types of cells . Even if technologies have been developed, such as 3C (chromatin conformation capture), 4C-seq , and sopnrb/NonO [It is still not clear how recruitment of factors to active genes is brought about, although several mechanisms have been proposed Fig. . Many prnrb/NonO , 48 havenrb/NonO .Structural studies of purified spliceosomes are improving , but higThe study of mRNP formation must consider all the coordinated events and interacting molecules at the active gene. One such consideration is the influence of chromatin on the transcription rate and pre-mRNA splicing . The traNucleosome density and deposition of histone variants can modulate the movement of RNA pol II . NucleosA number of histone modifications have been shown to influence splicing. Some of these may work through affecting the RNA pol II transcription rate and others by influencing the recruitment of splicing factors. One example is the H3K36 methylation that plays a role in nucleosome positioning and in limiting elongation rate and thereby influencing splicing . H3K36 mThe close contact between the chromatin and the pre-mRNP enables functional interactions between them. It has been reported that a pre-mRNP can influence transcription by recruiting a histone acetyltransferase to the active gene .Chromatin structure is likely to affect transcription termination. Nucleosomes are enriched downstream polyadenylation sites , and mayAll RNAs, including pre-mRNA and mRNA, have the ability to fold in alternative ways, following energy rules. Folding starts during transcription , 80. The7G cap [The repertoire of proteins that associate with RNA is substantial and diverse . Large-s7G cap , the pol7G cap , and the7G cap . Some pr7G cap . They ca7G cap . It shouApproximately, 20 different proteins (named A1 to U) belong to the group of heterogeneous nuclear proteins (hnRNPs) , 95. TheMethodological improvements are needed to characterize the complete protein content of specific endogenous pre-mRNPs. One must consider that the composition of pre-mRNPs is dynamic over time and along a gene. The BR pre-mRNPs at the gene, and the BR mRNPs in both the interchromatin and at the NPCs, are morphologically identifiable. This fact has made it possible to decide, using immune-EM, where proteins associate with and dissociate from these specific endogenous transcripts. So far, 34 different proteins were identified within BR pre-mRNPs. These proteins have functions related to transcription, capping, 3´ end cleavage and processing, splicing, packaging, export, and quality control . Some prDuring transcription, the growing pre-mRNA is rapidly and continuously assembled into a pre-mRNP complex, probably as soon as the transcript emerges at the surface of RNA pol II . Many diThe molecular background for packaging of pre-mRNPs is unknown in almost all cases. It has been suggested that interactions between SR proteins and the EJC compact pre-mRNPs . DimerizIn perichromatin regions in mammalian cell nuclei, short fibrillar and granular structures presumably represent nascent pre-mRNPs . The pacThe evolution of packaging of the pre-mRNP and mRNP has presumably been important for a number of reasons. Packaging might reduce the formation of RNA–DNA hybrids, so-called R-loops, which can cause genomic instability . FurtherAt the gene, the pre-mRNA interacts with the processing machineries responsible for capping, splicing, and 3′ cleavage and polyadenylation. The processing machineries all contribute to the composition and structure of the pre-mRNP and all leave marks in the mRNP that are important for subsequent events in gene expression, including export.Capping enzymes recognize the 5′ end of the pre-mRNA, essentially as soon as it appears at the RNA pol II exit tunnel . The nucIntron excision requires the multicomponent spliceosome . SpliceoSR proteins are characterized by having one or more RNA-binding domains and a domain rich in serine-arginine repeats . SR protSplicing is not only necessary for obtaining correct open reading frames. Splicing also leads to deposition of the EJC core . EJCs asEssentially, all pre-mRNAs are cleaved and polyadenylated by the 3′ cleavage and polyadenylation machinery. The processes and responsible components are biochemically well characterized. Proper 3′ end formation is necessary for mRNA export . More thA part of the polyadenylation machinery is recruited to the transcript via the RNA pol II CTD , 43. IntSome genes are extremely long, emphasising that synthesis of pre-mRNAs is dependent on the processivity of RNA Pol II . SynthesStudies in yeast show that both Nab2p and Pab1p, the mammalian orthologs PABPN1 and PABPC1, bind poly(A) tails. Nab2p is predominantly located in the nucleus and Pab1p in the cytoplasm. Nab2p protects poly(A) RNA from degradation by the exosome  and promSynthesis and processing of pre-mRNPs are complex molecular events that are not always perfectly performed. Aberrant products can be made and the cell must be able to handle these. The presence of quality control systems has been demonstrated in cells having specific mutations in the RNA or by knocking down enzymes that normally degrade aberrant RNAs. Transcripts with defects in capping, assembly, splicing, and 3′ end formation have all been shown to be degraded . DegradaTranscripts with retained introns or defective 3′ ends have been shown to be retained at the gene –152. TheIn a broader sense, it can be argued that also the deposition of proteins in connection to pre-mRNA processing provides a quality control. For example, a non-correct EJC will not support downstream processes such as export and translation.An mRNP released from its gene is in several aspects likely to be ready for export to the cytoplasm. It is folded in a compact but flexible manner. It is equipped for downstream processes, with a CBC, a large number of different proteins, a poly(A) tail, and it is spliced to have a translatable open reading frame. The question is what more an mRNP must contain to be export competent in the sense that it can be channelled through an NPC. An obvious answer is that each mRNP requires export adaptors and export receptors. Evolution has resulted in multiple different export adaptors and their recruitment to mRNPs is promoted by transcription and processing. The two main export receptors, NXF1 and CRM1, interact with different sets of export adaptors Fig. . Many adThe TREX complex is a conserved complex that has functions in transcription elongation, mRNA export, and genome stability , 158. ItTREX-2 is another conserved protein complex that has been shown to interact with the NPC basket , 167 andSR proteins can serve as export adaptors , 174. SREven if many processes in mRNP formation occur at the gene, important events such as completion of processing, binding of export receptors, and intranuclear transport take place in the interchromatin compartment Fig. . The proSplicing can occur post-transcriptionally. Fractionation of cell nuclei, followed by quantification of the catalytically active SF3b155 protein, estimated that about 20% of splicing takes place in the nucleoplasmic fraction . SeveralClusters of granules in the interchromatin, called interchromatin granule clusters at the EM level and speckles at the light microscope level, contain high concentrations of spliceosome components . The fulSynthesis of the poly(A) tail can be completed in the interchromatin. In BR genes, isolation by microdissection showed that polyadenylation is initiated at the gene and can be completed after release into the interchromatin .So far, we know very little about the structure of endogenous gene-specific mRNPs. The best characterized examples are the BR1 and BR2 mRNPs. These mRNPs, containing approximately 40 kb-long mRNAs, have a well-defined three-dimensional structure, and this structure has a hierarchy of folding that is established largely during transcription. At a first level, a 7 nm mRNP fibre is further folded, forming a 26 nm ribbon. This ribbon is folded into an overall ring-like complex .In yeast, EM analyses of purified mRNP populations suggest that mRNPs have an extended shape with lateral restrictions . The lenHigh sensitive mass spectrometry analyses of proteins that UV-crosslink to and co-purify with poly(A) RNAs have revealed a large number of proteins in direct contact with mRNAs. In mammalian cells, approximately 800 proteins are associated with the mRNA population , 185. EnStudies of endogenous gene-specific mRNPs are rare, but required for knowledge of the fate and behaviour of mRNPs. Experimental advantages make it possible to analyse properties of the BR1 and BR2 mRNPs. Already at release from the gene, these mRNPs are packaged and equipped with the majority of its associated proteins. BR1 and BR2 mRNPs have a protein content of about 60%, presumably corresponding to about 500 individual proteins . Some prBR mRNPs retain their morphology at the ultrastructural level from the release from the gene until translocation through the NPC. Even so, immune-EM has demonstrated that BR mRNPs recruit some proteins in the interchromatin. One example is the RNA helicase Ct-hrp84 . Most noN6-methyladenosine, pseudouridine, 5-methylcytosine, N1-methyl6adenosine, and 5-hydroxymethylcytosine modifications of coding RNAs were identified in eukaryotic cells. The m6A modification is the one most thoroughly analysed and described to date. The modifications can induce conformational changes of the RNA. They can also interfere with RNA–protein interactions by either blocking or facilitating the binding of specific proteins.Modified bases are important for the function of non-coding RNAs, such as tRNAs and rRNAs. More recently, it has been discovered that also mRNAs are subjected to chemical modifications that can influence and regulate their function and fate –193. So The enzymes that mediate and the enzymes that remove the modifications are mostly found in the nucleus, but can, in some cases, also be localized to the cytoplasm. Thus, both nuclear and cytoplasmic pre-mRNAs and mRNAs can be targeted and, perhaps, influenced in different ways, depending on the location. Studies of the modifications identified so far have reported consequences for splicing, export, translation, and stability of the pre-mRNPs and mRNPs , 195.For export, mRNPs depend on association with export receptors Fig. . These rThe major protein exporter CRM1, in association with Ran-GTP , is alsoA number of mRNAs have a 50 nucleotides-long sequence in their 3′ UTR that is sufficient to direct these mRNAs into an eIF4E and CRM1-dependent export pathway . This soThe NXF family member NXF3 can also recruit CRM1 to mRNPs.Several observations show that interactions between export adaptors and receptors are more diversified than previously thought. Binding of the main mRNP export receptor, NXF1, to export adaptors shows variation. For example, in addition to SR proteins and ALY/REF, the neuron-specific HuD protein is an adaptor for NXF1 . ComponeThe combination of various ways of recruiting different export adaptors, dependent on processing and sequence specificity, and the subsequent binding of the export receptors, show that there are alternative export pathways. Through such diversified interactions, export of selected subpopulations of mRNAs can occur , 211. DuDiploid cell nuclei contain chromosome territories and different compartments , 214. AcEM analyses have demonstrated that BR mRNPs in polytene nuclei, largely devoid of chromatin, move in all directions after release from the genes . This obQuantitative measurements and calculations for BR mRNPs show that the interchromatin in polytene nuclei contains a pool of BR mRNPs. No or little degradation of BR mRNPs takes place in the nucleus . In vivoIn the nuclei of mammalian cells, it appears that pools of poly(A) RNAs with retained introns are frequent , 228. ItIt is likely that a balance between intranuclear processing, degradation, and diffusion regulates mRNP export and thus influences the size of the pool of specific mRNPs. Retention and degradation of incompletely spliced mRNAs may be involved in neuronal development. It has been proposed that the protein PTB represses splicing, and that nuclear degradation of the incompletely spliced mRNA follows. The exosome and Tpr are involved . SpecifiThe NPC mediates almost all bidirectional transport of molecules between the nucleus and cytoplasm. In special circumstances, unconventional transport pathways may be used. For example, viruses can rebuild the NPC, allowing transport of viral mRNPs  and largAs shown in the BR system, an mRNP is largely produced at the gene and appears only modestly reorganized or modified in the interchromatin compartment . AdditioThe structure and composition of NPCs are well characterized , 241. ThTransport of mRNPs through the NPC occurs in three steps: docking onto the nuclear basket, translocation through the central channel, and release from the cytoplasmic fibrillar structures. Measurements of fluorescently labelled single mRNPs regarding the time for NPC interaction vary, ranging from 12 ms to several seconds , 244. ThAt the ultrastructural level, the interaction between the NPC and endogenous gene-specific mRNPs has been best characterized for the BR mRNPs. The process lasts for 65 ms to several seconds . In vivoWhen export occurs, a BR mRNP first interacts with the closed basket . The basFor the BR mRNPs, a partial unfolding then follows and its bent 26 nm ribbon becomes more straight as it moves through the central channel. The 5′ end leads the way , while tAt the cytoplasmic side of the NPC, there is additional unfolding of the BR mRNP into a 7–10 nm mRNP fibre. Other studies have shown that upon exit from the central channel, mRNPs associate with the cytoplasmic part of the NPC . A recenIn the cytoplasm, the mRNP is again remodelled . For exaGiven the importance of mRNP formation and export for gene expression, it is not surprising that a great number of defects in these processes have been connected to diseases. An important field of investigation is how dysregulation of splicing can be involved in cancerogenesis , 259. SpDysregulation of export components, including CRM1, THOC1, Rae1, and eIF4E, is described in different types of cancer cells . In accoReplication of viruses depends on host cell gene expression machineries and can thereby affect mRNP formation and transport. Viruses are studied for the obvious medical implications for developing treatments against viral infections. A second reason is that viral infections often provide experimental advantages and short cuts in understanding cellular processes. This is exemplified by the fruitful studies of splicing of adenovirus pre-mRNAs .During viral infection, the host cells activate defense mechanisms and the viruses interfere with the host cells to maximize viral replication. A common viral strategy is to shutoff host cell gene expression. In this way, the cellular immune response is evaded and resources are directed towards viral replication. Down-regulation of host cell gene expression can take place at most levels and individual viruses can use more than one mechanism.Transcription can be inhibited. For example, the 3C protease of the poliovirus cleaves TFIID  and SindPre-mRNA processing is also targeted. The influenza virus protein NS1 binds CPSF and inhibits PABP binding, thus interfering with 3′ cleavage and polyadenylation , 270. NSMany viruses, including herpes, influenza, and corona viruses, use viral or cellular endonucleases to degrade host cell mRNAs . At the Export of viral mRNAs from the nucleus to the cytoplasm has attracted considerable interest. In general, viruses take over cellular export factors, for example, the NXF1 and CRM1 export receptors, using a variety of strategies. Several viruses produce mRNAs that contain elements that can bind TREX  or that A deeper understanding of the processes of gene expression in the intact cell nucleus will have a great impact on many aspects of biology, ranging from evolution to cell differentiation and neurobiology. It will also be essential for diagnosis and treatment in most disciplines of medicine. It is important to continue to identify and characterize processes and the involved individual components, probably with an emphasis on multi-molecular complexes and the dynamic changes of such complexes. The most important and, at the same time, most challenging area is how everything operates in the living cell. Knowledge about the compositional and conformational dynamics of complexes and their interactions must be studied in the perspective of functional importance for endogenous genes in intact cells. This goal is demanding and exciting, and will require that new methods are developed. Such methods should be able to map the interplay within and between RNA–protein and protein–protein complexes, including the temporal and spatial dimensions. As an example, it is already evident that the pre-mRNP, the processing machineries, and the chromatin are all in very close contact. It is desirable to learn which interactions are the functionally important ones. Furthermore, we need to know more about how various components are recruited to sites of action and thereafter recycled, in the intact cell.


RNA‐seq methods that enable the simultaneous measurement of: (i) the states of internal gates, (ii) part performance , and (iii) impact on host gene expression. This is applied to a three‐input one‐output circuit consisting of three sensors, five NOR/NOT gates, and 46 genetic parts. Transcription profiles are obtained for all eight combinations of inputs, from which biophysical models can extract part activities and the response functions of sensors and gates. Various unexpected failure modes are identified, including cryptic antisense promoters, terminator failure, and a sensor malfunction due to media‐induced changes in host gene expression. This can guide the selection of new parts to fix these problems, which we demonstrate by using a bidirectional terminator to disrupt observed antisense transcription. This work introduces RNA‐seq as a powerful method for circuit characterization and debugging that overcomes the limitations of fluorescent reporters and scales to large systems composed of many parts.Genetic circuits implement computational operations within a cell. Debugging them is difficult because their function is defined by multiple states  that vary in time. Here, we develop  An ability to construct synthetic networks would enable the spatiotemporal control of biological processes , enable the measurement of genomewide mRNA levels with nucleotide resolution  to allow for many samples to be efficiently pooled and sequenced together  is produced that defines the expected curvature at each end of a transcription unit. Because the curvature is localized and fully captured by the first 500 nt of the hypothetical profile, this region is extracted and normalized by its maximum value to generate a correction factor profile C(x).To address this problem, a probabilistic method was developed to correct for curvature at the ends of each transcription unit, which utilizes the fragment length distribution as the only input variable. We begin by generating a fragment length distribution by directly analyzing the sequenced fragments. Using a Monte Carlo approach, large numbers of fragment lengths are drawn from this distribution. Fragments of these lengths are randomly mapped to positions falling within the boundaries of a hypothetical transcription unit 2,000 nt long. By counting the number of mapped fragments covering each nucleotide P(x) generated by counting the number of mapped fragments covering each nucleotide. Unwanted curvature is corrected for by dividing the value of P(x) for the first and last 500 nt of each transcription unit by C(xn), where xn is the distance in nucleotides to the nearest end of the transcript. Specifically, the corrected transcription profile is given by Next, the RNA‐seq data are used to generate the transcription profile for each transcription unit within the circuit. Fragments mapping exclusively within transcription unit boundaries are selected and a transcription profile Because the correction factor is only based on the length of the fragments and does not consider their sequence, the same correction factor profile is used for both the 5′‐ and 3′‐end, while the middle of a transcription unit, where no curvature is present, remains unmodified. The correction factor is only applied to known transcripts internal to the genetic circuit. It is not applied to other regions where part function does not have to be calculated; for example, read‐through between transcriptional units, transcripts from internal promoters, antisense transcription, and genomic transcription. Further details regarding the correction method are provided in i.e., fractional abundance of the total sequenced fragments), potential differences introduced during sample preparation or sequencing require correction to allow for comparison between samples. This is performed by calculating between‐sample normalization factors using a trimmed mean of M‐values (TMM) approach  is the number of RNAPs passing nucleotide position x per second. Here, we assume that all RNAPs that pass a nucleotide lead to an mRNA transcript and that all transcripts within the circuit degrade at the same rate. With these assumptions, the flux at a position x is given by the steady‐state number of transcripts M(x) at that position .Promoters cause sharp increases in the transcription profile Fig B. The acM(x) at each position x. However, the profiles cannot be quantified in units of transcripts and are thus presented in arbitrary units (au) and the fluxes are in au/s. The change in the number of transcripts at position x is given by ɣ = 0.0067 s−1 is the degradation rate of mRNA /dt = 0 and the flux of RNAP J(x) = ɣM(x). The activity of a promoter can be quantified as the change in flux δJ that occurs over the length of the part . To reduce the effect of fluctuations in the profile, an averaging window is applied immediately before and after the part boundaries dt=Jgiven by dM(x)dt=JTS has been previously defined as the fold decrease in gene expression before and after the terminator  and the output is the control of a promoter (turning RNAP flux on or off). For gates, the inputs and outputs are both promoters and the response function captures how the output changes as a function of the input at steady‐state. Unlike genetic parts, whose function can be extracted from a single profile, characterizing a sensor or circuit requires sampling the device in different states, extracting the activities of the input/output promoters, and then fitting these data to a mathematical model of device performance.δJon and δJoff, respectively  at steady‐state. For example, a NOT gate has one input promoter that drives the expression of a repressor that turns off an output promoter. The response function of this gate is Jin and δJout . Under these different conditions, the magnitude of the input promoter activity to the gate varies because of changes to the remainder of the circuit. We utilize those changes to plot data points for out Fig D and thiout Fig  to extraet al, et al, We applied our characterization method to a three‐input one‐output combinatorial logic circuit Fig A. Three 5 and 1.8 × 107 mapped fragments . To convert RPUs to arbitrary units that are compatible with the transcription profiles, the activity of the YFP output was measured in RPUs and compared to the average transcription profile across the ated Fig E with a ated Fig . A correated Fig F. Note tet al, Next, every promoter and terminator in the circuit was characterized across each of the eight states. An idiosyncrasy in our gate designs required a modification to the approach to characterize promoters. Normally, the rise in the transcription profile would be observed just after the promoter part. However, we use ribozymes as part of our gate design to insulate against contextual effects caused by changing the upstream promoters (Lou Tac−PTet1). Therefore, we calculate the dual promoter as a single promoter part using equation The use of ribozymes also makes it impossible to resolve the individual activities of multiple promoters in series (because the cleaved 5′‐RNA from transcripts from either promoter are lost during processing). Due to the use of multiple NOR gates, there are many examples of this in the circuit , we used the measured promoter strengths , the promoters cannot be individually resolved directly from the RNA‐seq data. To separate the activities of the two promoters, the individual outputs of the first and second promoters are defined as δJ1,on and δJ2,on, or δJ1,off and δJ2,off, where on and off refers to the presence or absence of the inducer. Then, for each state, the measured combined activity of the promoter pair δJ1+2 is equated to the expected activities for different combinations of inducers. For example, when the inducers that activate both sensors are present, δJ1+2 = δJ1,on + δJ2,on. This yields a set of algebraic equations to which δJ1,on, δJ1,off, δJ2,on, and δJ2,off can be fitted . However, PBAD2 produces a lower than expected on state, which manifests as a nearly flat transcription profile , and the output is the activity of the output promoter δJout. It is simple to calculate when this appears as a single promoter. If it is part of two promoters in series, the individual promoter activities are calculated as described for sensors, above  were expressed , and 125 significantly differentially expressed genes  with enrichment for functions broadly related to energy generation, anaerobic respiration, and fermentation , but without the remainder of the circuit. These data were used for differential gene analysis to search for potential differences in host gene expression. Expression of endogenous genes in cells containing the circuit for input states expressing only two or three genes (−/−/− and +/−/−) was highly correlated with the baseline  and aTc (PTet) sensors, with similar response functions across conditions . Differential expression analysis of the broken input states highlighted significant changes for 179 genes (P < 0.01), with enrichment of transport‐related functions for arabinose, xylose, and maltose  saw significant eight‐ to 56‐fold upregulation in culture tubes for the three broken input states  and bi‐cistronic insulators  7697 araD139 fhuA ΔlacX74 galK16 galE15 e14‐ϕ80dlacZΔM15 recA1 relA1 endA1 nupG rpsL (StrR) rph spoT1 Δ(mrr‐hsdRMS‐mcrBC) was used for cloning and measurements . Cells were grown in LB Miller broth  for harvesting plasmids. Cells were grown in MOPS EZ Rich  defined medium with 0.2% glycerol for circuit performance measurements. To select for the presence of plasmids, 50 μg/ml kanamycin  and 50 μg/ml spectinomycin  antibiotics were used.The Chemicals used to induce input promoters were isopropyl β‐D‐1‐thiogalactopyranoside , anhydrotetracycline hydrochloride , and L‐arabinose . Individual colonies were inoculated into MOPS EZ Rich Defined Medium  with 0.2% glycerol carbon source and 50 μg/ml kanamycin  and grown overnight for 16 h at 37°C and 1,000 rpm in V‐bottom 96‐well plates  in an ELMI Digital Thermos Microplates shaker incubator . The following day, cultures were diluted 178‐fold  into EZ Rich glycerol with kanamycin and grown under the same ELMI shaker incubator conditions for 3 h. For culture tube assays , cells were diluted 658‐fold (4.56 μl into 3 ml) into EZ Rich glycerol with kanamycin and inducers. For Erlenmeyer flask assays , cells were diluted 658‐fold (76 μl into 50 ml) into EZ Rich glycerol with kanamycin and inducers. Eight inducer combinations were used that cover the presence or absence of 0.5 mM IPTG, 10 ng/ml aTc, and 5 mM L‐arabinose. Culture tubes and Erlenmeyer flasks were then grown in an Innova 44 shaker  at 37°C and 250 rpm for 5 h. Finally, 40 μl of cell culture was placed into 160 μl of phosphate‐buffered saline (PBS) containing 2 mg/ml kanamycin to arrest translation and cell growth. These growth‐arrested cells were incubated in the PBS with kanamycin for one hour before fluorescence was measured using flow cytometry.Fluorescence of individual cells was measured using an LSRII Fortessa flow cytometer  controlled by the BD FACSDiva software. More than 20,000 gated events were collected for each state, and analysis of flow cytometry data was performed using FlowJo .E. coli DH10B cells harboring the genetic circuit plasmid and cultured under the circuit induction assay condition described above. Cultures were spun down at 4°C, 15,000 g for 3 min. Supernatants were discarded after centrifugation, and cell pellets were flash‐frozen in liquid nitrogen for storage at −80°C. Cells were lysed with 1 mg of lysozyme  in 10 mM Tris–HCl (pH 8.0) (USB 75825) supplemented with 0.1 mM EDTA (USB 15694). RNA was extracted with PureLink RNA Mini Kit  and further purified and concentrated with RNA Clean & Concentrator‐5  to ensure sufficient RNA concentrations (> 280 ng per sample). The purified RNA samples were analyzed using a Bioanalyzer , and Ribo‐Zero rRNA Removal Kit for bacteria  was used to deplete rRNA from the samples. We also checked the quality of the RNA extracted by calculating the RNA integrity number (RIN), which ranges from a value of 10 if all RNA is intact to 1 if the RNA is totally degraded  correction. Characterization of promoters, ribozymes, and terminators was performed using custom Python scripts that took a GFF reference of the circuit defining all part locations, their types, and any further information . To ensure termination strength and ribozyme cleavage were not underreported, measurements were filtered if there was a low level of RNAP flux (< 1 au/s) entering the terminator or leaving the ribozyme. Characterization of genetic parts was performed using a window size of n = 10 bp in equations Raw reads were mapped to the host genome (NCBI RefSeq: NC_01473.1) and circuit reference sequences using BWA  was used for all circuit simulations. For inputs, “low RPU” values of PTac = 0.0034, PTet = 0.0013, and PBAD = 0.0082, and “high RPU” values of PTac = 2.8, PTet = 4.4, and PBAD = 2.5 were used and the Eco1C1G1T1.UCF.json UCF file describing the gate response functions. Genetic circuit visualizations in SBOLv format −log]2. Here, S is the set of states , δJi is the measured activity of the output promoter for state i, and δJout,i is the expected output promoter activity calculated according to the gate's response function. For NOT gates, the response function is given by equation δJoutmin,δJoutmax, K, and n. For each state, to calculate the expected output promoter activity of the gate, the total input RNAP flux Jin was extracted from the transcription profile and used as input to equation To fit the response function of a gate when the output promoter was in isolation, we defined a least squares error function E2=Σi∈S[log−log]2. Here, δJ1+2,i is the measured combined activity of both output promoters for state i, and δJ1,i and δJ2,i are the expected individual output promoter activities calculated using the response functions of the first and second sensor/gate, respectively. For sensors, the response function was equated to either δJon or δJoff depending on the inducers that were present for that state. For example, given all combinations of inducers for two sensors in series: −/−, −/+, +/−, and +/+, then the expected combined output activity δJ1 + δJ2 would be given by: δJ1,off + δJ2,off, δJ1,off + δJ2,on, δJ1,on + δJ2,off, and δJ1,on + δJ2,on, respectively. For gates, the combined activity of two output promoters in series δJ1 + δJ2 was calculated as δJ1,out + δJ2,out, where the output promoter activity of each gate  was given by equation δJoutmin,δJoutmax, K, and n. For each state, to calculate the expected output promoter activity of a gate, the total input RNAP flux Jin was extracted from the transcription profile and used as input to equation δJon and δJoff) and gates  were then fitted to minimize the adapted error function. To extract the individual contributions of each promoter to the combined measured activity of the output promoters δJ1+2,i ], and for promoter 2 as δJ1+2,i × .When the output promoters of two sensors or gates were found in series, we characterized their individual behaviors by employing an adapted error function i∈S[logδJ+2,i−log, where pc is the cleavage efficiency  compatible with the experimentally measured transcription profiles by using the conversion factor 1 RPU = 2,895 au. The circuit DNA is scanned from the 5′‐ to 3′‐end to trace out the predicted profile. Upon reaching a promoter, the height of the profile is raised at the transcription start site by the predicted promoter strength (from Cello in au) multiplied by  was measured using a Cary 50 Bio spectrophotometer . For culture tube assays , cells were diluted 658‐fold (4.56 μl into 3 ml) into EZ Rich glycerol with kanamycin and inducers. For Erlenmeyer flask assays , cells were diluted 658‐fold (76 μl into 50 ml) into EZ Rich glycerol with kanamycin and inducers. To calculate the initial OD600 post‐dilution, the aforementioned OD600 value was divided by 658. Culture tubes and Erlenmeyer flasks were then grown in an Innova 44 shaker at 37°C and 250 rpm for five hours. The final OD600 for each vessel/inducer condition was measured for 1 ml of culture in the manner described above. The doubling time was calculated as (5 h)/log2.Individual colonies were inoculated into MOPS EZ Rich Defined Medium  with 0.2% glycerol carbon source and 50 μg/ml kanamycin  and grown overnight for 16 h at 37°C and 1,000 rpm in V‐bottom 96‐well plates  in an ELMI Digital Thermos Microplates shaker incubator . The following day, cultures were diluted 178‐fold (16.9 μl into 3 ml) into EZ Rich glycerol with kanamycin and grown in an Innova 44 shaker  at 37°C and 250 rpm for 3 h. Next, 1 ml of culture was added to a VWR disposable cuvette  and the optical density at 600 nm . Cultures were grown in culture tubes under different induction conditions, and RNA‐seq samples were prepared by following the protocols described above.New genetic parts were synthesized as gBlock gene fragments  and assembled into full genetic circuit using Gibson assembly . The modified circuit was sequence verified and transformed into https://www.ncbi.nlm.nih.gov/geo/) under the accession numbers GSE88835 and GSE98890. Python scripts that implement the characterization pipeline are released as open‐source software under the MIT license (Computer https://github.com/VoigtLab/MIT-BroadFoundry).RNA‐seq data collected in this study were deposited to Gene Expression Omnibus (CAV, TEG, YP, and AAKN conceived of the study. CAV, TEG, AEB, and YP wrote the manuscript. TEG, AEB, BSD, and DBG developed algorithms and performed simulations. AAKN, YP, and JZ carried out the experiments.The authors declare that they have no conflict of interest.AppendixClick here for additional data file.Dataset EV1Click here for additional data file.Dataset EV2Click here for additional data file.Code EV1Click here for additional data file.Review Process FileClick here for additional data file.


The correlation between eRNA production and enhancer activity has stimulated studies on the potential role of eRNAs in transcriptional regulation. Additionally, eRNA has also served as a marker for global identification of enhancers. Here I review the brief history and fascinating properties of eRNAs.Enhancer-derived RNAs ( NGS has not only made it possible to exhaustively catalog the transcripts from coding sequences (cis-regulatory elements (CREs) such as promoters and insulators, enhancers contain binding sites—DNA sequence motifs varying in length between 6 bp and 20 bp—of various transcription factors (TFs) Of the ncRNA family, a latecomer and somewhat unconventional member is the enhancer-derived RNAs (eRNAs), which are transcribed at the loci of enhancers While enhancers and TFs are the primary players of gene regulation, other factors capable of chemically modifying DNA and histones are gradually being incorporated into the overall scheme of gene regulation, many of which modulate the accessibility of chromatin to allow for direct contact between TFs and enhancers cis-regulatory activity of their template enhancers—that is, context-dependent, enhancer-stimulated mRNA synthesis of nearby genes—indicating an intimate association between enhancer function and eRNA production Although transcription from active enhancers was reported in the early 1990 s Because the abundance of eRNAs is 19–34-fold lower than that of gene transcripts eRNAs are not as stable as mRNAs e.g., H2AZ) or modifications  can be applied in eRNA detection experiments To assist with the enrichment of eRNAs, chromatin immunoprecipitation (ChIP) using antibodies against histone variants  to deplete several eRNAs in human cells found evidence for a causal role of eRNAs in transcriptional activation cdkn1bcdkn1b, is largely intact A more rigorous approach to investigate the function of the eRNA is to interfere with its synthesis by inserting a polyadenylation cassette near its transcription start site, which triggers premature transcription termination. In a recent study, this approach was applied on the locus of an enhancer regulating the expression of Since the completion of the Human Genome Project, a major focus of the scientific community is to develop effective means to precisely map the estimated millions of CREs orchestrating distinct gene expression patterns of each cell type and developmental stage e.g., JQ1) Indeed, thousands of enhancers have been identified through analyzing the transcripts from non-coding sequences, which exhibit excellent overlap with the enhancer maps generated by histone mark ChIP-seq e.g., in clinical settings) As the cost of high-throughput sequencing rapidly decreases, protocols allowing for accurate determination of eRNAs become more feasible over time. Given that RNA-seq is now routinely used in genomic characterization of cells and tissues, using eRNAs to predict enhancer activity can obviate the need for additional experiments to identify enhancers. This is an advantage over other enhancer mapping techniques such as ChIP-seq, DNase-seq, and assay for transposase-accessible chromatin (ATAC)-seq, all of which use protocols different from RNA-seq. Thus, eRNA analysis may be especially useful in situations where specimen supply is limited  and mutations that appear to be enriched in putative enhancer elements Finally, the growing need to decipher the epigenome in both normal and disease tissues is expected to demand ever more sensitive and comprehensive methodologies to characterize enhancers in various cellular contexts. Notably, the past decade has seen the documentation of a plethora of disease-associated genetic variants (The author declares no competing interests.