Regulation of transcription is vitally important for maintaining normal cellularhomeostasis and is also the basis for cellular differentiation, morphogenesisand the adaptability of any organism. Transcription activators, whichorchestrate time and locus-specific assembly of complex transcription machinery,act as key players in these processes. One way in which these activators arecontrolled is by the covalent attachment of the conserved protein, ubiquitin(Ub), which can serve as either a proteolytic or non-proteolytic signal. For asubset of the activators, polyubiquitination-dependent degradation of theactivator controls its abundance. In these cases transcription activation canrequire protein synthesis as well as internal or external stimulus. In contrast,other activators have been reported to undergo mono- or oligoubiquitination thatdoes not lead to protein degradation. The mechanisms by which monoubiquitinationof transcription activators affect their activities have been poorly understood.In a recent study, we demonstrated that monoubiquitination of some transcriptionactivators can inhibit transcription by recruiting the AAA+ ATPase Cdc48 , which thenextracts the ubiquitinated activator from DNA. Met30 E3 ligase and rapidly degraded, and this process is coupledto transcription activation of genes placed behind a LexA operator.To study the mechanism by which monoubiquitination of activators regulatestranscription, we modified this model system by fusing monoUb to the N-terminus ofLV. Because wild-type Ub fusions are co-translationally processed bydeubiquitinating enzymes (DUBs), we mutated G76 of Ub to V76 to prevent anydeubiquitination. To our surprise, the irreversible attachment of monoUb to LV hadtwo effects: (1) it prevented rapid turnover of LV, and (2) Ub-LV no longeractivated transcription. These observations prompted us to investigate howmonoubiquitination and polyubiquitination of transcription activators can haveopposite consequences in transcriptional output.It has long been recognized that a subset of transcription activators containtranscription activation domains that overlap with degrons, sequences within theprotein that signal polyubiquitination and subsequent degradation by the proteasome.For some activators, such as Myc and Estrogen Receptor α, transcription activationand activator turnover are interdependent. Much of the initial characterization ofthis phenomenon utilized a model activator LexA-VP16 (LV), which is a fusion of theDNA binding domain from bacterial LexA and the activation domain from Herpes SimplexVirus protein VP16. LV expressed in yeast is polyubiquitinated by theSCFBy chromatin immunoprecipitation (ChIP) assays, we found that Ub-LV has low occupancyat the promoter of a reporter gene. A key experiment was to demonstrate that thisdefect was due to specific properties associated with Ub. It is well known that thehydrophobic patch of Ub, composed of residues L8, I44 and V70, is a commonly usedsurface for Ub-protein interactions. When we introduced point mutations in thehydrophobic patch, we found that these mutations abolished the inhibitory effect ofUb fusion. Ub(L8A)-LV behaves like LV, which potently activates transcription andhas a short half-life. Thus, one or more Ub-interacting proteins is responsible forpreventing Ub-LV from binding to DNA. Furthermore, these results indicated thatdeubiquitination of the activator is required for transcription. Thus, DUBs may playa crucial role in determining the residence time of an activator at the promoter inorder to finely tune the transcriptional response.Cdc48 is a hexameric AAA+ ATPase that has been termed an "Ub-dependentsegregase". Previous studies from multiple laboratories had shown that Cdc48can extract a number of ubiquitinated substrates from chromatin, such as yeasttranscription repressor α2, RNA Polymerase II, and replication licensing factorCdt1. In most cases, Cdc48 and its cofactors remove polyubiquitinated proteins fromchromatin to facilitate their subsequent degradation by the proteasome. Wespeculated that Cdc48 also can interact with a monoUb signal and that Cdc48 wasresponsible for removing Ub-LV from DNA. By using temperature-sensitive alleles ofCdc48 or its cofactors Ufd1 and Npl4, we demonstrated that inactivation of Cdc48,Ufd1 or Npl4 is sufficient to restore high promoter occupancy by Ub-LV and leads torobust transcription activation. Based on the properties of Cdc48, we envision thatCdc48 functions in this case by transiently unfolding LexA, thereby causing it todissociate from DNA and prevent transcription.in vivo, and competition by the large pool ofunconjugated Ub in the cell is expected to weaken the interaction. We also observedsubstantial amounts of Ub-LV that are not bound to chromatin. These are allpotential competitors for Cdc48:Ufd1:Npl4 complexes. Thus, chromatin-bound Ub-LVlikely contains additional signals that contribute to recruitment of Cdc48. Throughexperiments that are not detailed here, we ruled out some of the usual suspects,such as SUMO modifications or transcription coactivators that may bind to VP16. Wespeculate that Cdc48 or its cofactor(s) has intrinsic affinity for chromatin itself.Either naked DNA, which is commonly found at promoter regions in yeast, ornucleosomes might be additional targeting signals. Low-affinity, non-specificinteractions with chromatin combined with specific interactions with monoUb couldtarget Cdc48 efficiently to promoter-bound Ub-LV. However, we cannot rule out thatother components of the transcription machinery may also contribute to Cdc48recruitment. An increasing number of proteins have been found to interact withCdc48/p97/VCP, and a VCP-interacting motif (VIM) was recently identified in multipleCdc48-interacting proteins. The yeast genome encodes dozens of proteins that have aconsensus VIM, some of which are known players in the transcription process.Our ChIP data demonstrated that Cdc48 was recruited to the promoter of the reportergene in the presence of Ub-LV or Ub-LexA (without the VP16 activation domain), butnot by Ub(L8A)-LV, which contains a point mutation in the hydrophobic patch. It issurprising that a single Ub attached to the activator was sufficient to recruitCdc48 and elicit its segregase activity. After all, monoUb by itself is not a verydistinctive signal MET genes)involved in the synthesis of sulfur-containing metabolites; however, undernon-inducing conditions, cell proliferation requires inactivation of Met4, which isachieved through oligoubiquitination by the E3 ligase SCFMet30.Interestingly, Met4 ubiquitination without Met4 degradation is sufficient to turnoff MET gene expression. We hypothesized that Cdc48 may play a rolein maintaining the MET genes in a repressed state undernon-inducing conditions. By measuring promoter occupancy by Met4 and monitoringtranscript levels of several Met4 target genes, including MET17,MET3 and CYS3, we showed that inactivation ofCdc48 leads to increased promoter occupancy by Met4 and partial de-repression of theMET genes. Importantly, in yeast strains where Met4 is notubiquitinated, Cdc48 inactivation does not affect MET geneexpression. These results strongly suggest that, analogous to the Ub-LV system,Cdc48 prevents ubiquitinated Met4 from stably binding to promoter DNA, therebyinhibiting transcription activation.Based on the knowledge gained from studying LexA-VP16, we sought to identify nativetranscription factors that are regulated similarly by monoubiquitination. Previouswork from the Kaiser lab had shown that yeast transcription activator Met4 ismodified by a short oligoUb chain and that this modification does not lead toprotein turnover. Met4 is the master regulator of sulfur metabolism. In the absenceof methionine, Met4 activates multiple genes (named It was recently reported that in human cells receptor-activated Smads (R-Smads),including Smad2 and Smad3, undergo mono- and oligoubiquitination in the nucleus.This modification attenuates transforming growth factor β (TGF-β) signaling withoutpromoting R-Smads proteolysis. We took advantage of an inhibitor of mammalian Cdc48, called DBeQ, to investigate possible involvement of p97 inR-Smads-dependent transcription. Indeed, we found that DBeQ treatment increasedR-Smads-mediated transcription as well as activator promoter occupancy. Theseeffects were observed both in the absence or presence of TGF-β, although they aremore pronounced in the absence of the signaling ligand. Thus, Cdc48/p97 consistentlyplays a major role in maintaining target genes in repressed states throughmonoubiquitinated activators.Given the dynamic nature of Ub modification, it is often difficult to assess thedirect effect of ubiquitination on transcription activators. Without specificantibodies that differentiate the ubiquitinated from non-ubiquitinated pool ofactivators, determining which pool is chromatin-bound and responsible for drivingtranscription has generally not been possible. The Ub-LV fusion we employed allowedus to bypass these obstacles. It is satisfying that the mechanism revealed from thestudy of the artificial activator Ub-LV applies as well to native transcriptionactivators such as Met4 and R-Smads. We expect that additional activators will befound to be regulated by a similar mechanism in the near future.MET gene network, this mode of regulation couldbe essential, as new protein synthesis can be limiting in an environment wheremethionine levels are low. Currently, the DUB that deubiquitinates Met4 uponmethionine depletion has not been identified. In the case of R-Smads, Usp15 has beenfound to be necessary for transcription activation under inducing conditions. Inthese cases, how the DUBs are regulated in response to changing environments areinteresting topics for future studies.One remaining question is why these systems employ a non-proteolytic route to achievetranscription downregulation. An obvious possibility is that mono- oroligoubiquitination is readily reversible: rapid induction of transcription bydeubiquitination of the activator would not require new protein synthesis. In thecase of the yeast


Upon fertilization, the genome of animal embryos remains transcriptionally inactive until the maternal-to-zygotic transition. At this time, the embryo takes control of its development and transcription begins. How the onset of zygotic transcription is regulated remains unclear. Here, we show that a dynamic competition for DNA binding between nucleosome-forming histones and transcription factors regulates zebrafish genome activation. Taking a quantitative approach, we found that the concentration of non-DNA-bound core histones sets the time for the onset of transcription. The reduction in nuclear histone concentration that coincides with genome activation does not affect nucleosome density on DNA, but allows transcription factors to compete successfully for DNA binding. In agreement with this, transcription factor binding is sensitive to histone levels and the concentration of transcription factors also affects the time of transcription. Our results demonstrate that the relative levels of histones and transcription factors regulate the onset of transcription in the embryo.DOI:http://dx.doi.org/10.7554/eLife.23326.001 The DNA in a fertilized egg contains all the information required to form an animal’s body. In order for the animal to develop properly, particular genes encoded in the DNA are only active at specific times. The DNA is wrapped around proteins called histones, which allows the DNA to be tightly packed inside the cell. However, histones can block other proteins called transcription factors from binding to the DNA to activate the genes. Young embryos initially develop with all of their genes switched off, relying on the nutrients and other molecules provided by their mother. After some time, the embryo starts to switch on its own genes to take control of its own development, but it was not clear how this happens.Joseph et al. investigated how genes are activated in zebrafish embryos, which are often used as models to study how animals develop. The experiments show that competition between histones and transcription factors for binding to DNA controls when genes are switched on. In young fish embryos, there are so many histones present that transcription factors have no opportunity to bind to DNA. Over time, however, the numbers of histones decrease, allowing transcription factors to bind to DNA and switch on genes.Histones and transcription factors regulate the activity of genes throughout the life of the animal. Therefore, competition between these two types of protein may also control gene activity in other situations. A better understanding of how gene activity is controlled could allow researchers to more easily grow different types of cell in the laboratory or to reprogram specific cells in the body. As such, these new findings may aid the development of therapies to regenerate organs or tissues that have been damaged by injury or disease.DOI:http://dx.doi.org/10.7554/eLife.23326.002 In many organisms, early embryonic development is directed exclusively by maternal products that are deposited into the female gamete during oogenesis. Following the clearance of a subset of these products , transcrXenopus embryos. This effect was maintained only when non-specific DNA was added to titrate chromatin assembly  and zebrafish   . RNA for, Nanog)   into embryos at the 1-cell stage and then analyzed the onset of transcription for the previously characterized set of genes  and 1B. To test whether the effect we observe upon injecting the histone cocktail required an increase in the level of all four core histones, we next removed one histone at a time from the cocktail. The total protein content was kept constant by raising the level of the other three histones. Removing any histone from the histone cocktail impaired the ability of the histone cocktail to delay the onset of transcription . These rSince the onset of zygotic transcription is known to be required for gastrulation , we analptx3-injected embryos at early 1K stage (one time-point prior to when genes are first induced in uninjected embryos) revealed that the level of expression is increased upon injection of ptx3 mRNA  content and, correspondingly, the number of molecules of core histones in embryos using a quantitative mass spectrometry approach we recently developed  . We obseBecause transcription takes place in the nucleus, we next wanted to investigate the concentration of non-DNA-bound histones in this compartment of the cell. First, we analyzed the dynamics of histone localization by lightsheet microscopy of living embryos . As expeWe next analyzed whether the decreased concentration of non-DNA bound histones in the nucleus is accompanied by a reduced density of nucleosomes on chromatin. We quantified the amount of histone H2B in the chromatin fraction of embryos ranging from 256-cell to high stage. Comparing the amount of histone H2B between stages revealed that the level of H2B scales with the amount of DNA . This isOur finding that the concentration of histones in the nucleus determines the onset of transcription without a significant change in global nucleosome density on DNA suggests that a simple depletion model cannot explain a role for histone levels in the timing of zygotic transcription. To explain the effect of histone levels on zygotic genome activation, we hypothesized that the transcriptional machinery (for simplicity referred to as transcription factors) competes with nucleosome-forming histones for binding to only a minimal fraction of the total DNA, corresponding to transcription-factor-binding sites . In suchIf competition between nucleosome-forming histones and the transcriptional machinery determines the onset of transcription, it would be predicted that the levels at which transcription factors are present could also affect the timing of zygotic transcription. To test this, we changed the level of Pou5f3, a transcription factor that has been identified as being required for the activation of a large set of genes during genome activation . To analTo reduce the level of Pou5f3, we used a previously characterized morpholino  and confapoeb and dusp6 at high stage  revealed that the level of expression is increased significantly upon injection of pou5f3 and sox19b mRNA for apoeb and dusp6  and identified co-precipitated DNA fragments by qPCR  or GAL4-VP16 resulted in premature transcription  , as wellDrosophila, Xenopus, and zebrafish  and transgenic embryos were dechorionated immediately upon fertilization, synchronized and allowed to develop to the desired stage at 28°C. Stage was determined by morphology and corroborated by cell counting. In terms of absolute time, the time between collected stages around ZGA was consistent between all conditions within an experiment and for all experiments. Histone cocktail and BSA  were injected into the yolk at the 1-cell stage at 22 ng per embryo. Pou5f3 anti-sense morpholino was injected at 6 ng per embryo, together with 1 ng of p53 morpholino (Zebrafish were maintained and raised under standard conditions. Wild-type (TLAB)  into the yolk was carried out. Human H4 cDNA was cloned into a pCS2+ vector with C-terminal sfGFP  . Human PTX3 cDNA was cloned into a pCS2+ vector with a C-terminal RFP. dge, MA) . mRNA eneif4g2a or input in ChIP-qPCR analysis. Relative mRNA expression levels were calculated via 1/(2∧(gene-eif4g2a)). Fold difference was calculated by dividing the relative mRNA expression level value of the test sample over control.Twenty-five embryos per developmental stage were snap frozen in liquid nitrogen. RNeasy Mini Kit  was used to extract RNA. For Tg(TRE:GFP) embryos, contaminating DNA was removed from RNA preparations using the DNA-free Kit . mRNA was converted to cDNA using the iScript cDNA Synthesis Kit . SYBR green  with Rox  was used as the qPCR master mix. Primers were used at a final concentration of 500 nM and sequences can be found in SCR_013672, Zeiss). Images were imported into the Imaris software  and the spot tool was used to calculate cell number.Embryos were fixed in 4% formaldehyde in Danieau’s solution at 4°C overnight. The next day, embryos were washed with Danieau’s solution and then permeabilized with 0.2% Triton X in Danieau’s solution for 30 min. Subsequently, embryos were incubated for 10 min in DAPI (1 µg/ml) and then washed several times with Danieau’s solution. Embryos were placed in an inverted agarose holder and covered with Danieau’s solution for imaging. An upright Zeiss LSM 780 NLO microscope equipped with a coherent Chameleon Vision II infrared laser was used for two-photon excitation . DAPI was excited with 780 nm and detected using a non-descanned GaAsP detector (BIG-Module) with BP450/60 or SP485. Samples were imaged with either a Zeiss W Plan-Apochromat 20 × 1.0 or 40 × 1.0 dipping objective. Images were acquired using a four tile scan of multiple z-sections (3–3.5 µm steps). Tiles were stitched with the ZEN software . Samples were boiled with SDS loading buffer at 98°C, run on 4–12% polyacrylamide NuPAGE Bis-Tris gels  and blotted onto a nitrocellulose membrane . Primary antibodies were incubated at RT for 1 hr or overnight at 4°C and secondary antibodies were incubated at RT for 45 min. Primary and secondary antibodies used are listed in Embryos were manually deyolked at the desired stage and snap frozen in liquid nitrogen. For all proteins, equal numbers of embryos were analyzed for each developmental stage (H4 and H2A eLife. Your full submission has been evaluated by Janet Rossant (Senior Editor) and three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the decision was reached after discussions between the reviewers. Based on our discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.Thank you for choosing to send your work entitled "Histone level determines the timing of zygotic genome activation in zebrafish" for consideration at As you will see, the reviewers broadly agreed on several key weaknesses in this work:1) The extensive overlap with previous work;2) The absence of a clear demonstration that adding excess histones alters nucleosome levels on DNA (or that depletion of H2B reduces nucleosome occupancy);3) Model that perhaps over-simplifies the scenario. Nothing in this manuscript or previous work pinpoints nucleosomes as the repressors. It remains entirely possible that it is a factor that binds nucleosomes – and not the octamer itself – that inhibits transcription.However, all reviewers also saw the potential in this system and encourage the authors to delve deeper into this interesting question to shed new light on zygotic genome activation.Reviewer #1:Work described in this manuscript tests the idea that histone levels define the timing of zygotic genome activation, using zebrafish as a model system.Xenopus oocytes and embryos . These studies involved injection of excess DNA, core histones, TBP and the activator VP16, leading to the conclusion that transcriptional quiescence before zygotic genome activation was due to a general repressive effect of histones and a defect in transcription activation. Work from the Wolffe lab also noted that they had not established that core histones were directly responsible for repression, but that nucleosome-binding proteins such as linker histones could functionally carry out repression. This possibility was recently supported by work in Drosophila  which showed that a specific variant of linker histone H1 affected the timing of zygotic genome activation .Background on the question: The idea that core histones repress zygotic transcription in the early embryo dates back to the early 90s  and was explored in detail in The current manuscript explores the role of histones in zebrafish genome activation. Quantitative mass spec nicely measures the histone levels across early development, confirming their gross excess over DNA. Injection of a cocktail of core histones is found to delay gastrulation and the onset of zygotic transcription (as measured by qPCR of 6 transcripts). All core histones were required for a maximal delay in genome activation, indicating that the formation of an entire nucleosome is essential for full repression. The reciprocal experiments were also performed, wherein the level of histone H2B is reduced to decrease the potential for nucleosome formation. These experiments show a modest premature activation of zygotic transcription. The model proposed is well in line with that from the Wolffe lab, involving both the titration of inhibitory nucleosomes and the involvement of an activator that can overcome such repression at selected sites.Overall, the manuscript is well written and the hypothesis being tested is clear. The experiments seem to be carefully performed and the results are generally convincing. I like the quantitative mass spec and the focus on activation of endogenous genes (previous work largely looked at activation of genes located on exogenous DNA). One caveat is that the effects of histone H2B depletion on gene activation appears small, and I would encourage the authors to consider trying to reduce more than one core histone to achieve stronger effects.Xenopus, reducing the impact of this work. Further, the authors should acknowledge that their work does not define core histones as the repressors, only that repression requires nucleosome formation (as in earlier work). I see this as a missed opportunity, given the very nice system developed. Why not target linker histones, specific transcription activators or repressors? That would allow them to go beyond what has been shown in Xenopus and Drosophila to define direct repressors and activators in their system.My primary concern is that the conclusions do not go much beyond the model established in the mid-90s in Reviewer #2:eLife.In the manuscript "Histone level determines the timing of zygotic genome activation in the zebrafish", the authors demonstrate that excessive quantities of histone proteins repress transcription during the early stages of zebrafish development. When injected with even more excess quantities of histone proteins, development and onset of transcription is delayed. These findings show are exciting and suggest a clever mechanism for regulating the timing of transition from maternal to zygotic transcription. If the authors could address the following issues, the manuscript would be of interest to the readers of 1) The authors give a compelling argument for the role of excess nucleosomes in preventing zygotic transition, however some direct evidence of the excess nucleosomes actually interacting with DNA is necessary to support their data. Showing maintenance of nucleosomes on the tested loci upon overexpression of histones and loss of nucleosome occupancy when the H2B morpholino is used would provide some evidence of the effect of the excess nucleosomes. As well, using ChIP to assay binding of transcriptional machinery to show differences with excess or loss of histone proteins.2) While the authors show a role for nucleosomes in the timing of transition, they do not address the idea that a repressive chromatin remodeling complex or some other factor may still contribute to the timing of transition as well. Hdac1 is a likely candidate for this role, but there are many repressors that could contribute. It would be helpful for the authors to test Hdac1 or other repressors or discuss the possibility that other repressors may exist.3) Authors estimate that the entire genome is fully bound once every 170bp with histones, with no mention of possible variations. While this is obviously an estimate, it would be helpful to at least note that this is an estimate based on the expectation of full occupancy of the genome with histones during the entire course of experiments.4) The histone calculations are a bit confusing, particularly with the phrase "2x1011 copies of each per embryo". It would be helpful to state histone proteins rather than copies. Is this actually per embryo or per cell? If it is per embryo, then the calculations for excess amounts of histones added for later experiments are confusing, since the measurement is taken at the 8 cell stage, so 8 there would be genomes worth of histones. It seems like the 1.8x1011 histone proteins should be divided by 8x the number of histones in one genome rather than one single genome, giving an excess of 636 rather than 5049.5) Calculations of the fold excess are slightly different using the numbers in the paper. Using the numbers from the paper, the fold excess at the 1000 cell stage calculates to 705 rather than 715. 1.8x1011 histone proteins divided by 35,425,427 comes out to 5081 rather than 5049.Reviewer #3:The question of how a developing embryo controls the transition from reliance on maternally loaded mRNA to activation of zygotic expression is a central question in developmental biology. The current manuscript examines this phenomenon in the context of early zebrafish development. The authors hypothesize that maternally loaded histones compete with the transcriptional machinery and that zygotic transcription ensues upon reaching a critical ratio of DNA to histones. They take the general approach of manipulation of the histone:DNA ratio by (1) microinjection of exogenous recombinant human histones or (2) depletion of endogenous H2B using morpholino technology, assessing (1) transcriptional states via RT-PCR of selected protein-coding genes as well as (2) developmental progression.Xenopus development via microinjection of DNA. Regardless of the difference in technique, the current manuscript and the Mechali/Wolffe studies come to an identical conclusion – that the ratio of histones to DNA in early development is a critical determinant of the onset of zygotic transcription. Mechali and colleagues state "Our data suggest that the large excess of histones represses gene activation during early development…". Given this precedent in the literature, it is unclear to me why the current manuscript states repeatedly that the importance of the histone:DNA ratio in zygotic gene activation has never been established .These experiments are highly reminiscent of papers published about 20 years ago by the Mechali  Cell 77: 439-449) and Wolffe  EMBO J 14: 1752-1765) labs in which the histone:DNA ratio was manipulated in early Mechanistically, I do not understand what exactly the current manuscript is measuring. The authors purchase recombinant human histones (NEB). These are supplied as purified, single polypeptides – not reconstituted into octamers or any other physiologically relevant complex. They then mix the histones together and inject the cocktail into embryos. It is not clear to me what the fate of the injected histones would be in this scenario. Recombinant histones at low salt concentration will be largely unstructured, certainly not present as octamers (and likely not as H3/H4 tetramers or H2A/H2B heterodimers). I encourage the authors to provide biochemical evidence that injected histones can associate in biochemical species similar to existing histone pools in the embryo – which are largely associated with histone chaperones. The conclusion by the authors that nucleosomes are the relevant species in their experiments competing with transcription factors for DNA (as opposed to non-specific interactions of unstructured basic proteins) seems poorly supported by the data.[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]eLife. Your revised article has been favorably evaluated by Robb Krumlauf (Senior Editor), a Reviewing Editor, and two reviewers. The manuscript has been considerably improved, and would now be suitable for publication once several major concerns below are addressed.Thank you for resubmitting your work entitled "Competition between histone and transcription factor binding regulates the onset of transcription in zebrafish embryos" for further consideration at Summary:eLife" if the following concerns could be addressed or explained properly in an updated manuscript.By utilizing efficient quantitative techniques, Joseph et al. present a very interesting study that supports a competitive interaction between histones and transcription factors to set the time for ZGA. They conclude that histones and critical transcription factors compete in binding to DNA, and eventually, the concentration of non-DNA bound core histone decides the timing of ZGA during zebrafish gastrulation. By injecting stoichiometric mixture of all four core histones they showed that increasing their levels delays the onset of zygotic transcription. On the other hand, decreasing levels of histones, by sequestering H4 with PTX3, shows an opposite effect. Using a quantitative mass spectrometry approach, the authors calculated the number of non-DNA bound histones per cell in genomes worth of histone, and showed that this parameter drops dramatically as embryos approach gastrulation. This provides opportunity for critical transcription factors (such as Pou5f3) to bind to DNA and initiate transcription. Following this argument, increasing and decreasing the level of Pou5f3 predates and delays the onset of ZGA, respectively. Finally, in the context of a TRE-GFP reporter construct, by injecting either a heterologous transcription factor (rTA-VP16) alone, or by co-injecting it with histone cocktails, a ChIP-PCR analysis provided further evidence for the competitive interaction between histones and transcription factors to bind to TRE elements in the GFP reporter. Altogether, this study provides a detailed mechanistic analysis to answer a few important issues related to a long-standing concept in the field. The quantitative techniques followed in this study highlights the usefulness of zebrafish to answer critical questions which could be otherwise difficult in other model systems. Such analyses would be beneficial for the zebrafish community as a whole. Taken together, this would be suitable for publication in "Major points that must be addressed in a revised manuscript:Sox2, Nanog) to see the effect on ZGA? What about non-developmental factors? According to their model, the competition between ZGA-related transcription factors and histones should not be specific to Pou5f3 only. Rather, knockdown of any other pluripotency factor should have the same ZGA-related phenotype. The specificity/ generality of this phenomenon should be tested by knockdown of at least one additional TF.1) In 2) The authors should clarify the nature of their competition model, through additional experiments, or clarifications in the text. At present (and as detailed below) it is not clear how the authors envision that excess histones are functioning, and the text and data are not consistent in this regard. For this work to have the desired impact, we encourage the authors to work towards clearly articulating and substantiating their model.A) The reviewers were concerned about the absence of proof for a competition model in the data presented, and alternative interpretations for the data presented are plausible: for example, histone 'excess' may affect replication timing, thereby affecting timing of transcriptional activation at ZGA. While the integrated transgene and VP16 experiments is strong, one would like to see it 'working' in an endogenous target: at the very least, the same ChIP experiment performed in excess of histone should be done for the Apoeb gene for Pouf3t and Sox19 transcription factors. Otherwise, it is difficult to reconcile it with the specificity in competition program discussed by the authors in the subsection “Competition for DNA binding between histones and transcription factors regulates the onset of transcription”.B) Perhaps an additional possibility to address whether the competition model is indeed occurring, would be to reduce histone chaperone activity at the 'early' 1K stage. This should increase non-DNA bound histones, and therefore should have the same effect as the injection cocktail.C) If the effect on transcriptional activation is assumed to be due to non-DNA bound histones (subsection “Onset of transcription coincides with a reduction in nuclear histone concentration” and Summary) – why is the addition of Histone cocktail effective considering that the authors conclude that 'this led to efficient chromatin incorporation of the histones injected as determined using Cy5 labelling' ?D) Along the same lines, in the Discussion, the authors state that, because the 4 core histones are necessary to see the effects of the injected cocktail on transcriptional activation, this suggests a role for the nucleosome for the repressor function of histones. This statement is not consistent with the interpretation that it is the 'non-DNA bound histones' that is effecting ZGA timing (e.g. nucleosome is composed by the histone octamer plus the DNA). Given the biochemical properties of the 4 histones, it is also unlikely that the histones injected remain as octamers in the cells.E) Similarly, related to the PTX3 experiments (F) The authors conclude that histones are effectively incorporated by doing immunofluorescence with Cy5-labelled H4. However, nuclear localisation is not a proof for incorporation – did the authors try the same immunofluorescence with e.g. either pre-extraction with tritron or salt?3) In  [Editors’ note: the author responses to the first round of peer review follow.]As you will see, the reviewers broadly agreed on several key weaknesses in this work:1) The extensive overlap with previous work;We agree with the reviewers that previous work using injected species of exogenous DNA had suggested that histones might play a role in transcription regulation in the embryo. We apologize for not doing enough justice to this work and now clarified this part of the manuscript.Questions that remained after previous studies, however, include:Whether histones regulate the timing of zygotic transcription (transcription of endogenous genes in the embryo).If so, which histones are involved (important for understanding the mechanism).How this all should work (the mechanism).In our revised manuscript, we address these questions. We first set up in an in vivo assay to analyze the onset of zygotic transcription  and show2) The absence of a clear demonstration that adding excess histones alters nucleosome levels on DNA (or that depletion of H2B reduces nucleosome occupancy);We show that histone levels regulate the time of zygotic transcription. Transcription starts when the nuclear concentration of histones drops, but this is not accompanied by a loss of nucleosome density on DNA. This suggests that the onset of transcription is regulated by the concentration of non-DNA bound histones in the nucleus. Based on this data, we propose that the onset of transcription in the embryo is regulated by a competition for DNA binding between non-DNA bound histones and transcription factors and we provide direct experimental evidence for this model. We therefore propose that the effect of changing histone levels experimentally is exerted through changes in the non-DNA bound concentration of histones and not by changes in global nucleosome density.3) Model that perhaps over-simplifies the scenario. Nothing in this manuscript or previous work pinpoints nucleosomes as the repressors. It remains entirely possible that it is a factor that binds nucleosomes – and not the octamer itself – that inhibits transcription.This is where we have really expanded the manuscript. Our experiments show that the four core histones mediate the repressor effect. Moreover, our finding that histones are neither completely depleted from the soluble fraction, nor generally depleted from chromatin, argues against a model in which a global loss of nucleosome density on chromatin causes the onset of transcription. This strongly suggests that histones themselves function as the actual repressors of transcription and argues against a role for the depletion of proteins associated with core histones. Furthermore, the model presented in the earlier submission did indeed simplify the scenario. We now present data that the levels of activators are also important. We provide direct evidence for a model in which competition for DNA binding between histones and activators regulates the onset of transcription in the embryo.However, all reviewers also saw the potential in this system and encourage the authors to delve deeper into this interesting question to shed new light on zygotic genome activation.Reviewer #1:Work described in this manuscript tests the idea that histone levels define the timing of zygotic genome activation, using zebrafish as a model system.Background on the question: The idea that core histones repress zygotic transcription in the early embryo dates back to the early 90s  and was explored in detail in Xenopus oocytes and embryos . These studies involved injection of excess DNA, core histones, TBP and the activator VP16, leading to the conclusion that transcriptional quiescence before zygotic genome activation was due to a general repressive effect of histones and a defect in transcription activation. Work from the Wolffe lab also noted that they had not established that core histones were directly responsible for repression, but that nucleosome-binding proteins such as linker histones could functionally carry out repression. This possibility was recently supported by work in Drosophila  which showed that a specific variant of linker histone H1 affected the timing of zygotic genome activation .Xenopus embryo. We apologize and now discuss existing work more clearly and identify the aspects which needed further work.We agree with the reviewer that in our previous manuscript, we did not do enough justice to the work that has previously shown a role for histones in repressing transcription in the early The current manuscript explores the role of histones in zebrafish genome activation. Quantitative mass spec nicely measures the histone levels across early development, confirming their gross excess over DNA. Injection of a cocktail of core histones is found to delay gastrulation and the onset of zygotic transcription (as measured by qPCR of 6 transcripts). All core histones were required for a maximal delay in genome activation, indicating that the formation of an entire nucleosome is essential for full repression. The reciprocal experiments were also performed, wherein the level of histone H2B is reduced to decrease the potential for nucleosome formation. These experiments show a modest premature activation of zygotic transcription. The model proposed is well in line with that from the Wolffe lab, involving both the titration of inhibitory nucleosomes and the involvement of an activator that can overcome such repression at selected sites.Overall, the manuscript is well written and the hypothesis being tested is clear. The experiments seem to be carefully performed and the results are generally convincing. I like the quantitative mass spec and the focus on activation of endogenous genes (previous work largely looked at activation of genes located on exogenous DNA). One caveat is that the effects of histone H2B depletion on gene activation appears small, and I would encourage the authors to consider trying to reduce more than one core histone to achieve stronger effects.As suggested, we have changed our approach to reduce histones. Since we see by mass spectrometry that a large amount of histones is present as protein from fertilization on , we deciSox2 are highly translated during development) and because many of these activators might function cooperatively, they all need to be present at the right concentration at the right time.We also expand our Discussion, and indicate that due to a role of activator levels in our competition model, pushing genome activation back will always be harder than forward (with more histones). Activators might be limiting because they need to be translated . I see this as a missed opportunity, given the very nice system developed. Why not target linker histones, specific transcription activators or repressors? That would allow them to go beyond what has been shown in Xenopus and Drosophila to define direct repressors and activators in their system.With respect to the novelty of our work, and the identity of the repressor, we refer to the key points.Reviewer #2:In the manuscript "Histone level determines the timing of zygotic genome activation in the zebrafish", the authors demonstrate that excessive quantities of histone proteins repress transcription during the early stages of zebrafish development. When injected with even more excess quantities of histone proteins, development and onset of transcription is delayed. These findings show are exciting and suggest a clever mechanism for regulating the timing of transition from maternal to zygotic transcription. If the authors could address the following issues, the manuscript would be of interest to the readers of eLife.1) The authors give a compelling argument for the role of excess nucleosomes in preventing zygotic transition, however some direct evidence of the excess nucleosomes actually interacting with DNA is necessary to support their data. Showing maintenance of nucleosomes on the tested loci upon overexpression of histones and loss of nucleosome occupancy when the H2B morpholino is used would provide some evidence of the effect of the excess nucleosomes. As well, using ChIP to assay binding of transcriptional machinery to show differences with excess or loss of histone proteins.To address these points, we did the following:We have chemically tagged H4 and show that it incorporates into chromatin . We haveExactly as suggested, we used ChIP-qPCR to assess the binding of a transcriptional activator in response to changing histone levels. Increasing histone levels causes a reduction of binding of the activator to its target site . In agre2) While the authors show a role for nucleosomes in the timing of transition, they do not address the idea that a repressive chromatin remodeling complex or some other factor may still contribute to the timing of transition as well. Hdac1 is a likely candidate for this role, but there are many repressors that could contribute. It would be helpful for the authors to test Hdac1 or other repressors or discuss the possibility that other repressors may exist.Here, we show that histone levels in combination with transcription factor levels are sufficient to change the timing of transcription. We agree with the reviewer, however, that chromatin remodeling complexes might also contribute to genome activation. We have now included a discussion on how the activity of chromatin remodelers might also be mediated by histone levels.3) Authors estimate that the entire genome is fully bound once every 170bp with histones, with no mention of possible variations. While this is obviously an estimate, it would be helpful to at least note that this is an estimate based on the expectation of full occupancy of the genome with histones during the entire course of experiments.The number we used is based on MNase-Seq experiments and the average nucleosome repeat length derived from that . We have clarified this in the text. Of note is that the nucleosome repeat length that we use is not critical for our model. Even variations of nucleosome occupancy from 160 to 200 do not substantially affect our calculations of the total nor nuclear histone concentration.4) The histone calculations are a bit confusing, particularly with the phrase "2x1011 copies of each per embryo". It would be helpful to state histone proteins rather than copies. Is this actually per embryo or per cell? If it is per embryo, then the calculations for excess amounts of histones added for later experiments are confusing, since the measurement is taken at the 8 cell stage, so 8 there would be genomes worth of histones. It seems like the 1.8x1011 histone proteins should be divided by 8x the number of histones in one genome rather than one single genome, giving an excess of 636 rather than 5049.We apologize for the confusion and provide more clarity with our calculations in the new manuscript  Calculations of the fold excess are slightly different using the numbers in the paper. Using the numbers from the paper, the fold excess at the 1000 cell stage calculates to 705 rather than 715. 1.8x1011 histone proteins divided by 35,425,427 comes out to 5081 rather than 5049.This discrepancy was due to rounding our numbers. There is more clarity in our new Reviewer #3:The question of how a developing embryo controls the transition from reliance on maternally loaded mRNA to activation of zygotic expression is a central question in developmental biology. The current manuscript examines this phenomenon in the context of early zebrafish development. The authors hypothesize that maternally loaded histones compete with the transcriptional machinery and that zygotic transcription ensues upon reaching a critical ratio of DNA to histones. They take the general approach of manipulation of the histone:DNA ratio by (1) microinjection of exogenous recombinant human histones or (2) depletion of endogenous H2B using morpholino technology, assessing (1) transcriptional states via RT-PCR of selected protein-coding genes as well as (2) developmental progression.For a summary of the data in the revised manuscript we refer to the key points.These experiments are highly reminiscent of papers published about 20 years ago by the Mechali  Cell 77: 439-449) and Wolffe  EMBO J 14: 1752-1765) labs in which the histone:DNA ratio was manipulated in early Xenopus development via microinjection of DNA. Regardless of the difference in technique, the current manuscript and the Mechali/Wolffe studies come to an identical conclusion – that the ratio of histones to DNA in early development is a critical determinant of the onset of zygotic transcription. Mechali and colleagues state "Our data suggest that the large excess of histones represses gene activation during early development…". Given this precedent in the literature, it is unclear to me why the current manuscript states repeatedly that the importance of the histone:DNA ratio in zygotic gene activation has never been established .We apologize for not clearly reviewing existing data previously, and included a thorough discussion of existing work in our revised manuscript.With respect to the novelty of our work, we refer to the key points.Mechanistically, I do not understand what exactly the current manuscript is measuring. The authors purchase recombinant human histones (NEB). These are supplied as purified, single polypeptides – not reconstituted into octamers or any other physiologically relevant complex. They then mix the histones together and inject the cocktail into embryos. It is not clear to me what the fate of the injected histones would be in this scenario. Recombinant histones at low salt concentration will be largely unstructured, certainly not present as octamers (and likely not as H3/H4 tetramers or H2A/H2B heterodimers). I encourage the authors to provide biochemical evidence that injected histones can associate in biochemical species similar to existing histone pools in the embryo – which are largely associated with histone chaperones. The conclusion by the authors that nucleosomes are the relevant species in their experiments competing with transcription factors for DNA (as opposed to non-specific interactions of unstructured basic proteins) seems poorly supported by the data.To address these points, we did the following:We provide proof that the human NEB histones are functional. We have chemically tagged H4 that shows that it localizes to the nucleus and incorporates into chromatin .Additionally, the experiments that remove one histone from the cocktail and elevate the levels of the other three, argue against a non-specific effect of an unstructured basic protein. If the delay in transcription was a result of randomly injecting unstructured protein, then the histone minus one experiment should also have the same effect. Which they don’t.[Editors' note: the author responses to the re-review follow.]Major points that must be addressed in a revised manuscript:1) In We apologize for the confusion. We agree with the reviewers that, according to our model, the competition between ZGA-related transcription factors and histones should not be specific to Pou5f3. We have clarified this in the text.Following the reviewers’ suggestion, we now show that knockdown of Sox19b and FoxH1 also results in a delay in the transcription of target genes  The authors should clarify the nature of their competition model, through additional experiments, or clarifications in the text. At present (and as detailed below) it is not clear how the authors envision that excess histones are functioning, and the text and data are not consistent in this regard. For this work to have the desired impact, we encourage the authors to work towards clearly articulating and substantiating their model.Thank you for raising this point. We have detailed below what experiments we have done and what changes we have made to the text to address it.A) The reviewers were concerned about the absence of proof for a competition model in the data presented, and alternative interpretations for the data presented are plausible: for example, histone 'excess' may affect replication timing, thereby affecting timing of transcriptional activation at ZGA.That is a good point. Replication timing is not affected in our experiments, and we have now clarified this in the text .While the integrated transgene and VP16 experiments is strong, one would like to see it 'working' in an endogenous target: at the very least, the same ChIP experiment performed in excess of histone should be done for the Apoeb gene for Pouf3t and Sox19 transcription factors. Otherwise, it is difficult to reconcile it with the specificity in competition program discussed by the authors in the subsection “Competition for DNA binding between histones and transcription factors regulates the onset of transcription”.This is an excellent point. We used the transgene because the regulation of endogenous genes is complex and the data is therefore harder to interpret. However, we now show that the binding of Pou5f3 on apoeb , is also reduced in the presence of more histones  Perhaps an additional possibility to address whether the competition model is indeed occurring, would be to reduce histone chaperone activity at the 'early' 1K stage. This should increase non-DNA bound histones, and therefore should have the same effect as the injection cocktail.This is in principle a good idea. However, the lack of information on chaperones in zebrafish, as well as the – related – lack of specific drugs make this experiment difficult to perform and interpret. Especially in combination with the fact that in the best-case scenario this experiment would recapitulate the histone cocktail effect without providing more (quantitative) insight in the competition model, we decided not to try these experiments.C) If the effect on transcriptional activation is assumed to be due to non-DNA bound histones (subsection “Onset of transcription coincides with a reduction in nuclear histone concentration” and Summary) – why is the addition of Histone cocktail effective considering that the authors conclude that 'this led to efficient chromatin incorporation of the histones injected as determined using Cy5 labelling' ?This experiment was designed to show that the histones we inject can incorporate in chromatin, and thus become part of, and behave like, the endogenous pool of histones available for competition. Like endogenous histones in the early embryo, not all histones will incorporate into chromatin due to their excess levels. We have now clarified this in text .D) Along the same lines, in the Discussion, the authors state that, because the 4 core histones are necessary to see the effects of the injected cocktail on transcriptional activation, this suggests a role for the nucleosome for the repressor function of histones. This statement is not consistent with the interpretation that it is the 'non-DNA bound histones' that is effecting ZGA timing (e.g. nucleosome is composed by the histone octamer plus the DNA). Given the biochemical properties of the 4 histones, it is also unlikely that the histones injected remain as octamers in the cells.We apologize for the confusion. Our model is based on the knowledge that the four core histones are required to form a histone octamer (and nucleosome). Of course, the histones only form a histone octamer when associating with DNA and it is therefore expected that the concentration of soluble histones dictates how likely a nucleosome is formed on an available piece of DNA. Thus, we propose that the soluble histones compete with transcription factors for binding, but ultimately it is either the histone octamer or transcription factor that occupies DNA. We have clarified this in the text (throughout).E) Similarly, related to the PTX3 experiments . Therefore, when histones are not available to bind to DNA (because they are bound by PTX3), this lowers their effective concentration. We have clarified this in the text (subsection “Decreasing the pool of available histones causes premature transcription”).F) The authors conclude that histones are effectively incorporated by doing immunofluorescence with Cy5-labelled H4. However, nuclear localisation is not a proof for incorporation – did the authors try the same immunofluorescence with e.g. either pre-extraction with tritron or salt?We apologize for the confusion. The conclusion that histones are incorporated was not based on the nuclear localization of the labeled histone but rather on the fact that we see the label in chromatin during metaphase. We have now clarified this in the text  and in 3) In We assume that there is an increase in histone binding but we have not looked at this. We have now clarified this in the text (subsection “Transcription factor binding is sensitive to histone levels”) in the section on Pou5f3 binding , which i


Escherichia coli K-12 MG1655 and a Bacilli cassette linked with lactose degradation. Six genes from both cassettes had similar functions related to carbohydrate metabolism, namely, hydrolase, aldolase, kinase, isomerase, transporter, and transcription factor. The Escherichia coli sulphoglycolysis cassette was thus predicted to be associated with lactose degradation. This prediction was confirmed experimentally: expression of genes coding for aldolase (yihT), isomerase (yihS), and kinase (yihV) was dramatically increased during growth on lactose. These genes were previously shown to be activated during growth on sulphoquinovose, so our observation may indicate multi-functional capabilities of the respective proteins. Transcription starts for yihT, yihV and yihW were mapped in silico, in vitro and in vivo. Out of three promoters for yihT, one was active only during growth on lactose. We further showed that switches in yihT transcription are controlled by YihW, a DeoR-family transcription factor in the Escherichia coli cassette. YihW acted as a carbon source-dependent dual regulator involved in sustaining the baseline growth in the absence of lac-operon, with function either complementary, or opposite to a global regulator of carbohydrate metabolism, cAMP-CRP.Comparative genomics analysis of conserved gene cassettes demonstrated resemblance between a recently described cassette of genes involved in sulphoquinovose degradation in  Many bacterial species can assimilate a wide range of carbohydrate substrates, adjusting to environmental changes, and are able to quickly switch between pathways. With multiple energy sources in the environment, a bacterial cell goes through metabolic assessment and, in most cases, selects a specific preferred energy source, e.g. glucose3, over a hundred thousand genes related to carbohydrate metabolism have been described for over six hundred genomes of various bacterial species. Moreover, some of these genes are known to encode proteins that have multiple functions and can be involved in different metabolic pathways. For example, thermophilic glycoside hydrolase CoGH1A from Caldicellulosiruptor owensensis has a high capability of saccharification, being able to hydrolyze various substrates4, polysaccharide lyase Smlt1473 from Stenotrophomonas maltophilia is able to catalyze chemical bond cleavage in a variety of carbohydrate molecules5, and hexokinase Glk/TM1469 from Thermotoga maritima switches between substrates under different temperature conditions6.Genes encoding proteins responsible for carbohydrate transport, degradation, and synthesis are very diverse. According to the IMG database8. CCR allows the cell to select a preferred source over a non-preferred one, maintaining a proper energy balance9. In a glucose-free environment or under stress conditions bacteria switch to the next best carbon source available11. In Escherichia coli and related species, this switch is often controlled by cAMP-CRP as a global regulator, and a local regulator encoded by a gene usually located either in the same operon with genes encoding enzymes and/or transporters necessary for growth on alternative carbohydrates, or in a divergent orientation14.Carbohydrate metabolism in most bacteria is controlled by the carbon catabolite repression (CCR) mechanism16. It functions as a dimer in the form of the CRP-cAMP complex and may activate or repress transcription. In most cases, CRP downregulates transcription when its operator region is situated upstream of the transcription start site18. CRP-mediated activation occurs when cAMP-CRP binds upstream of the target promoter and directly contacts RNA polymerase19. It can also bind DNA in a cAMP-independent manner, resulting in a weak target repression17. An additional, substrate-specific layer of regulation is provided by the binding sites of local transcription factors from six major families  that are located close to the CRP binding sites, or even overlap them. In most cases, these factors play a role opposite to CRP and act as dimers. Dimerisation caused by the ligand binding leads to a metabolic switch20.cAMP-CRP is one of the most important transcription factors that regulates transcription initiation for more than a hundred genes of carbohydrate catabolismE. coli, supplementing the lac operon described in the pioneer work of François Jacob and Jacques Monod21. The lac operon consists of three genes, encoding β-galactosidase, β-galactoside transacetylase, and β-galactoside permease. Until now, no other pathways for lactose utilisation in E. coli have been known.Here, we identify and explore the second lactose catabolism system in et al.22 described the E. coli gene cassette consisting of 10 yih genes and, according to that study, completing the biogeochemical cycle of sulphur. Four reactions of sulphoquinovose (SQ) catabolism were demonstrated using purified, heterologously expressed enzymes; other genes from this cassette were predicted to be involved in regulation, transport, hydrolysis, and other processes of sulphoglycolysis. The yih genes have not been linked to the lactose catabolism, and their transcription profile has not been characterised.In a recent biochemical study, Denger 29. We compared functional composition of the yih cassette with other carbohydrate metabolism gene cassettes and found a match with a cassette involved in the lactose catabolism of Bacilli. Six genes from these two cassettes encoded proteins with similar functions, namely, hydrolase, aldolase, kinase, isomerase, transporter, and transcription factor, allowing us to suggest involvement of the E. coli yih-cassette in the lactose catabolism. We then confirmed it experimentally using expression analysis. We also mapped promoters for the yih genes, and studied how the switch to lactose utilization is regulated, thus describing a possible additional pathway for lactose utilization in E. coli and suggesting multi-functional features for the respective proteins.Unknown functions of a gene can be predicted using comparative genomics analysis, based on gene proximity in the chromosome and functional similarity of their protein productsE. coli and present in some other Enterobacteriaceae, and a lactose catabolism cassette in several Bacilli species  and Staphylococcus spp. .Comparison of the combinations of gene functions in cassettes related to bacterial carbohydrate metabolism revealed a six-gene match between a cassette linked with SQ degradation in ies Fig. . Namely,3. The genes coding for aldolases also belonged to one and the same cluster of orthologous genes (COG3684). Such co-localization of genes linked with functions similar to those required for the Bacilli lactose pathway , aldolases (EC 4.1.2), kinases (EC 2.7.1), and isomerases (2.3.1), and also transporters and transcription factors , kinase (yihV), isomerase (yihS), alpha-glucosidase (yihQ), and a local transcription factor (yihW). To obtain a comprehensive view on the transcription pattern in this locus, the yihU and yihQ genes coding for aldolase and aldose epimerase that are not associated with the suggested pathway, were also incorporated in the analyses  revealed the presence of promoters capable of efficient binding with RNAP in the intergenic regions yihU/V, yihV/W, and yihS/R  relative to the annotated ATG codon. RNA synthesis from yihUP1, yihUP2, yihUP4, and yihUP5 was also confirmed with primer yihU/yihV_F  is not involved in the lactose metabolism. For yihV (putative kinase), several transcription starts were also mapped, with the major one located at –25/–27 relative to the starting ATG (a large cluster of predicted promoters with the highest score of 11.35 at position 4071737 in U00096.2). This promoter was activated during growth on lactose , or their regulatory role in the yihT transcription32. For yihW, only one promoter was mapped, with transcription start located at −27/−28 relative to the ATG codon , aldolase (yihT), and isomerase (yihS) were activated during growth on galactose and lactose, and this activation was more pronounced during growth on lactose. yihW expression, conversely, was approximately 2-fold repressed during growth with lactose suggesting either its function as a repressor for the yih genes at this time point, or a self-repressor function. The yihU gene, encoding a putative reductase, did not respond to differences in carbon sources , encoding epimerase and glucosidase, respectively, was also independent (yihR) or barely dependent (yihQ) on the carbohydrates used; this result supports our previous assumption about their independent transcription  to get more pronounced effects of the carbon source. The experiment showed that yihT expression on both glucose and lactose is totally controlled by YihW that functions as a sugar-dependent dual regulator , aldolase (YihT), and isomerase (YihS)  Figs  and 2. Iage Fig.  and S3.yihW gene, encoding a predicted DeoR-family transcriptional regulator, was also lactose-activated during exponential growth but repressed upon transition to starvation where the substrates start depleting. This observation led us to an assumption that YihW may act as a dual regulator for the yih genes, activating them in the presence of extra lactose and repressing when lactose is lacking. Stimulated growth of lac-minus cultures with lactose in the absence of YihW, that is even more pronounced on galactose  medium supplemented with 5% or 10% (v/v) LB, 0.2% (w/v) of the appropriate carbon source - D-glucose, D-galactose, lactose or glycerol. Cultures were grown aerobically at 37 °C under constant shaking. Cells were harvested after 4.5 hours of growth . To express the cAMP-CRP protein, E. coli BL21* (DE3) cells44 were chemically transformed with the pET_CRP plasmid constructed based on pET28b (Invitrogen), with the crp gene inserted between the NdeI and Bpu1102 sites. Purified transformants were grown on LB or Terrific broth (TB) at 37 °C till OD650 = 0.3, and the protein synthesis was induced by addition of a very low IPTG concentration (20 μM) to avoid the effects of potential protein toxicity. The induction rate of CRP after 3 hours of IPTG induction was ~70% of the total protein  was added, and cells were further sonicated 3 × 30 seconds on ice. Then, the lysate was cleared by 20 min centrifugation at 15,000 rpm, +4 °C, and loaded onto cAMP-agarose (Sigma). Then, the protocol described in19 was applied. Final yield of purified native CRP was ~20 mg of protein for 200 ml culture. All steps of purification are illustrated in Supplementary Fig. To obtain pure CRP, BL21*(DE3) cells with overproduced CRP  were harvested by centrifugation at +4 °C, washed with 1xPBS and lysed with 2 ml BugBuster protein extraction reagent (Novagen) for 20 minutes at +4 °C. Due to the very high induction level, all protein was in insoluble fraction even when very low IPTG concentration and low temperatures were used. To extract CRP from inclusion bodies, two-step lysis was performed: after initial lysis with BugBuster, 6 ml of ice-cold Lysis bufferyih cassettes from the Escherichia coli, Enterobacter cloacae, Salmonella enterica, Cronobacter turicensis, and Pantoea anantis . Regions were then extended in both directions by 100 nucleotides allowing for possible errors in the original open reading frame assignments. The resulting nucleotide sequences were obtained from GenBank45 and aligned using the T-Coffee multiple alignment tool46. Promoter-like motifs in the E. coli K-12 MG1655 genome (U00096.2) were found by the software PlatProm30 and its unified version PlatPromU47 where the impact of a sigma factor in promoter recognition by RNAP was depleted. Transcription starts were localized by band-shift assays with σ70-RNA polymerase, single-round transcription in vitro as previously described48 and by measuring the lengths of primer extension products in vivo. For primer extension, cells were grown in MS in the presence of 10% LB and 0.2% glucose, lactose, or glycerol, and harvested at OD650 = 0.4. RNA was extracted from 10 ml of cell culture with TRIzol reagent  and treated with DNAse I  following the manufacturers’ instructions. 10 µg of total RNA was incubated with 4 pmol of the [γ-32P]ATP-labeled appropriate primer . Gels were scanned using PMI Molecular Imager  and autoradiographed.Four intergenic regions with lengths allowing for promoter or transcription factor site mapping, e.g., exceeding 40–50 nucleotides, were selected for genes of the 49, with respect to the mapped transcription start sites, within the region −250/+50. The resulting sites were ranked according to their scores and conservation among the Enterobacteriaceae species. Highly conserved sites with best scores were retained for further analysis.Candidate binding sites for transcription factors were identified in the intergenic regions using phylogenetic footprinting and the Virtual Footprint databaseyih cassette. DNA fragments containing potential promoter regions of separate genes were PCR amplified  was added. After further 30 min at 37 °C, complexes were loaded onto a 5% polyacrylamide gel that had been pre-warmed to 37 °C. Protein complexes were separated from free DNA by electrophoresis in 1xTBE buffer. The DNA fragment containing known σ70-promoter of the hns gene amplified with primers hns_Bgl_263 and hns_Xba (Table hns (primers RT-PCR) where no promoters were predicted was used as a negative control.Electrophoretic mobility shift assays  were used to test the efficiency of the RNA polymerase binding to the regulatory regions of the genes belonging to the ba Table  was usedyih genes. This approach allowed us to skip addition of extracellular cAMP in the sample in vitro. CRP was produced in BL21*(DE3) cells, then cells (10 ml) were harvested, washed and resuspended in 3 ml of 0.5× Transcription buffer with 10 mM PMSF. Cell lysate was prepared using standard 3 × 30 seconds rounds of sonication, and a total protein concentration was determined using the Bradford reagent . Approximate molar concentration of CRP in the lysate was calculated, and 1–8 molar excess of the protein was used in band-shifts.The same approach, but with cell lysate with the overproduced protein, was used to test the ability of cAMP-CRP to bind the regulatory regions of the 15, and 20 μg/ml heparin was added to some samples to prove or disprove binding specificity.To estimate the efficiency of binding, free DNA fragments and proteins were loaded on separate lanes. Gels were stained by ethidium bromide to reveal bands containing DNA. Western blotting was used for specific detection of CRP in the complexes. In brief, band-shifts were performed as above in duplicate, one of which was stained with ethidium bromide and the other one was transferred onto a PVDF membrane  using the Mini trans-blot protocol . The membrane was then blocked with 0.5% skimmed milk  and incubated with anti-CRP (T14) antibodies  in Tris-buffered saline (TBS) supplemented with 0.1% Tween-20 and 0.5% skimmed milk for 2 hours at 37 °C. After 3 × 10 min washes with TBS-T, secondary antibodies  were added and incubation was allowed for 1 hour at room temperature. Membranes were stained with Western-blue stabilized substrate for alkaline phosphatase  and scanned. To prove specific binding of CRP, band-shifts were also made with the purified protein with or without addition of cAMP. Band-shifts were made exactly as above, but 200 μM cAMP was added to the samples, to the gel and the running 1xTBE bufferhns and ysaA-aRNA51 were used as controls. Data obtained from at least three biological samples and analyzed in three statistical replicates were calculated by the ΔΔCt method. The error bars indicate the standard deviations of the respective mean values.DT-322 thermocycler  and SYBR Green I as a fluorescent dye  were used for quantitative PCR (qRT-PCR). Primers used for reverse transcription (−RT) and amplification (−PCR) are listed in Supplementary Table Data generated or analyzed during this study are included in the present manuscript, additional datasets analyzed during the current study are available from the corresponding author by request. All strains and plasmids are also available by request.http://www.bioinformatics.org/sms/multi_align.html). Graph in Fig. Schemes in Figs Supplementary Figures and Tables


Schizosaccharomyces pombe, we examine the role of Cdk9, and related Mcs6/Cdk7 and Lsk1/Cdk12 kinases, on transcription at base-pair resolution with Precision Run-On sequencing (PRO-seq). Within a minute of Cdk9 inhibition, phosphorylation of Pol II-associated factor, Spt5 is undetectable. The effects of Cdk9 inhibition are more severe than inhibition of Cdk7 and Cdk12, resulting in a shift of Pol II toward the transcription start site (TSS). A time course of Cdk9 inhibition reveals that early transcribing Pol II can escape promoter-proximal regions, but with a severely reduced elongation rate of only ~400 bp/min. Our results in fission yeast suggest the existence of a conserved global regulatory checkpoint that requires Cdk9 kinase activity.Post-translational modifications of the transcription elongation complex provide mechanisms to fine-tune gene expression, yet their specific impacts on RNA polymerase II regulation remain difficult to ascertain. Here, in  While post-translational modifications of the RNA polymerase II complex regulate gene expression, the exact mechanism remains unknown. Here, the authors use altered-specificity kinase mutations to examine the role of Cdk9, Mcs6/Cdk7, and Lsk1/Cdk12 on transcription at base-pair resolution in yeast. For example, the concerted action of Cdk7, Cdk9, and Cdk12 are required for the early transition of Pol II from an initiating to a productively elongating and RNA processing complex6. Cdk7, the kinase subunit of the TFIIH general transcription factor complex , phosphorylates the largest subunit of Pol II within the C-terminal domain (CTD) at the ser5 and ser7 positions of heptad amino acid sequence, YSPTSPS, which is repeated 29 times in S. pombe9. The modification of Pol II by Mcs6 is critical for recruitment of the 5′ RNA capping machinery and release of the mediator complex at the earliest stages of gene transcription13. Ser2 of the CTD heptad repeat is also phosphorylated as Pol II elongates further into the gene body14, and both Cdk9 and Lsk1  have been suggested to modify this residue, possibly to facilitate 3′ RNA processing15.Multiple kinases modify RNA polymerase II (Pol II) and associated pausing and elongation factors to regulate Pol II transcription and pre-mRNA processingSpt4 and Spt5 comprise a highly conserved elongation factor -sensitivity inducing factor, or DSIF in metazoans)16. Spt5 possesses an unstructured C-terminal repeat (CTR) domain, with specific residues actively targeted by Cdk917. Moreover, the structure of eukaryotic Spt4/5 bound to transcribing RNA Pol II reveals contacts with upstream DNA and nascent RNA exiting Pol II18, and depletions of Spt5 in fission yeast globally reduce levels of elongating Pol II19. Cdk9 (Bur1 in S. cerevisiae) positively impacts transcription elongation20, and has been suggested to convert Spt5 from a negative to positive elongation factor in humans21. Such observations may reflect regulated recruitment of elongation-coupled factors, as association of capping enzymes and polymerase-associated factor 1 (Paf1) with the transcription complex are influenced by the phosphorylation status of Spt525.Similar to the Pol II CTD, targets within elongation factors can also mediate essential regulation of transcription elongation. Spt5 and negative elongation factor (NELF) by positive transcription elongation factor b , is thought to be required for the release of elongating Pol II from a promoter-proximal pause site27. This Cdk9-regulated process (referred to hereafter as pausing) is now known to occur on the vast majority of genes in mammals28, serving to regulate gene expression, or to ensure appropriate processing of RNA and/or maturation of the elongation complex29. In budding yeast, which lack homologs of all NELF subunits, such pausing has not been observed, yet perturbation of Cdk9 and Spt5 impair normal transcription by Pol II31. In contrast, pause-like distributions of Pol II have been found in the fission yeast S. pombe, where they depend on Spt432. In addition, S. pombe Cdk9 interacts with the triphosphatase component of the 5′ RNA capping apparatus (Pct1), possibly to alleviate a Spt4-5-induced checkpoint that ensures proper pre-mRNA processing17. Still, such a checkpoint in yeast remains largely speculative, and the relation to Cdk9-regulated pausing in metazoans is unknown.In most metazoan systems, the phosphorylation of 33. The use of such analog-sensitive (AS) strains in fission yeast revealed correlated changes in global mRNA levels upon inhibition of Mcs6 or Cdk9 within minutes of inhibition34. Inhibitions of an AS TFIIH kinase resulted in a shift of Pol II levels toward the 5′ ends of genes in budding yeast after 1 h35 and impaired pausing and termination in humans after a day of inhibition36. However, even within just 1 h, the primary effects of kinase inhibition on Pol II dynamics may be masked, especially in yeast where genes are short and transcribed in a matter of minutes37.Investigations of the direct influence of kinase activity on the dynamics and regulation of transcription in vivo requires highly selective inhibitors, which can be difficult to obtain. Mutant kinases have been designed with exquisite specificity for bulky ATP analogs that selectively inhibit their activityS. pombe to rapidly and selectively inhibit the activities of Cdk9, Mcs6, and Lsk1 individually or in combination. Using precision run-on sequencing analysis, we observe global changes in transcriptional dynamics within minutes of treatment, indicative of the general influence of these kinases on transcription by Pol II. While almost no instantaneous changes in Pol II distribution result from the loss of lsk1as activity, inhibition of cdk9as and/or mcs6as dramatically alter the transcription landscape within 5 min, despite non-overlapping substrates. Spt5 phosphorylation becomes undetectable after just 1 min of cdk9as inhibition; however, unlike metazoans, elongating Pol II is not trapped at the promoter-proximal pause site28. Instead, a fine-scale time course of cdk9as inhibition reveals that promoter-proximal Pol II has a severely compromised elongation rate, while downstream Pol II appears less affected. Our results support the existence of a Cdk9-dependent early elongation checkpoint in fission yeast, which may reflect a primitive form of Cdk9-regulated promoter-proximal pausing in metazoans.In this study, to gain an insight into the impact of transcription complex-targeting kinases on Pol II elongation, we use AS kinase mutants in the fission yeast 32, yet whether escape from such pausing is regulated through kinase activity is not known. In metazoans, kinase inhibitors, including DRB and flavopiridol (FP), which primarily inhibit Cdk9, have been instrumental in the discovery of promoter-proximal pausing as a major regulatory hurdle for most genes28. However, off-target effects of chemical inhibitors make selective ablation of Cdk9 activity in vivo nearly impossible38.Recently, we reported promoter-proximal pause-like distributions of Pol II on many genes in fission yeastCdk9 activity on Pol II pausing and elongation at single-nucleotide resolution in fission yeast, we performed PRO-seq in an AS mutant strain, cdk9as, which is vulnerable to inhibition by the addition of 3-MB-PP139. Phosphorylation of Spt5 became undetectable after 1 min of drug addition in cdk9as  or an equivalent volume of DMSO (untreated). As a control, we prepared PRO-seq libraries from equivalently treated wild-type (wt) cells. Comparison of untreated cdk9as profiles with wt profiles revealed no obvious differences , which measures the enrichment of Pol II in the promoter-proximal region (TSS to +100 nt) relative to the gene body (“+” indicates downstream and “−” indicates upstream). Although we observed striking increases in early gene body PRO-seq signal (TSS to +1 kb) resulting from 5 min of cdk9as inhibition, pausing did not explain this effect  was as sensitive to treatment with 3-MB-PP1 as cdk9as  was also inhibited by 10 μM 3-MB-PP1  after inhibition. We also profiled distributions of Pol II in strains containing combinations of AS kinases. Combined inhibition of either lsk1as and cdk9as, or mcs6as5 and cdk9as produced transcriptional defects resembling those of cdk9as or mcs6as5 alone, although possible basal effects of the combined mutations limited further interpretation from these strains  the promoter-proximal region (TSS to +100), (2) the early gene body (+150 to +450 from the TSS), and (3) the late gene body (−300 to CPS)  revealed global kinase-dependent effects at both ends of transcription units  Fig. . Most stnit Fig. .cdk9as activity, Pol II may exhibit a compromised processivity, preventing transcription elongation beyond a certain distance from the TSS37. Alternatively, the altered transcription profiles may reflect a reduced rate of early transcribing ECs incapable, or delayed in, accelerating to the natural productive speed. Assuming an unchanged initiation rate, the increased density of elongating Pol II at the 5′ end would be compatible with a reduced elongation rate of Pol II, which within 5 min may not have allowed Pol II to transcribe fully across the longest genes.Several mechanisms might explain the observed gene length-dependent effects on transcription. Consistent with a loss of signal from the 3′ ends of longer genes . Intriguingly, average profiles for these genes reveal a rapid recession of Pol II density toward the CPS  across the entire length of genes with increasing time of Cdk9 inhibition excludes premature termination as the sole explanation for effects on Pol II elongation. However, we cannot rule out a combination of defects in the elongation rate and processivity of affected Pol II, which might account for non-uniform increases in Pol II density seen at later time points after cdk9as inhibition  resulted in higher density across genes due to reduced elongation rate, similar to the phenotype of strains lacking the Spt5 binding partner Spt432. Importantly, we cannot rule out effects being a consequence of another known or unknown Cdk9 target. Considering that Cdk9 is capable of targeting additional transcription-coupled substrates, including the Pol II largest subunit CTD49, it is possible that the effects we observe are attributable to other targets, or combinations thereof. Alternatively, the discrepancy of effects on Pol II transcription may represent separable functions of Spt5. The depletion of Spt5 might directly influence transcription elongation and processivity through loss of known interactions with Pol II, template DNA, and the nascent RNA50, whereas inhibiting-specific phosphorylation of the unstructured, CTR of Spt5 might only affect transcription rate, perhaps by impairing recruitment of additional factors51.Surprisingly, our Pol II distributions upon loss of Spt5 phosphorylation at the early stages of elongation may reduce association of factors, which help to negotiate nucleosomes52. Transcription elongation is known to both, modulate and be influenced by the chromatin environment54. For example, inability to recruit Paf1 and the facilitates chromatin transcription (FACT) complex could reduce the ability of the elongation complex to progress through nucleosomes55. Alternatively, Pol II slowing may occur to ensure recruitment of 5′ capping components, similar to pre-mRNA processing steps at gene 3′ ends56. Inhibiting phosphorylation of Spt5 might prevent or delay the association of capping enzymes, potentially reducing elongation rates across entire transcription units. Nonetheless, further work comparing the importance of Spt5 phosphorylation with other Cdk9 substrates will be crucial for understanding the mechanisms by which Cdk9 exerts influence over the kinetics of transcription elongation in fission yeast.The loss of Cdk9 inhibition appears directly comparable to clearing waves observed in metazoan systems following the inhibition of P-TEFb28. But whereas most other transcribing Pol II is trapped at the promoter-proximal pause site in flies and mammals28, these early gene body elongation complexes continue through the gene with severely impaired rates in S. pombe. We propose that NELF, which is a critical pausing factor in Drosophila and mammals59, may serve to exploit an early elongation checkpoint, observed here in fission yeast, creating a rate limiting, Cdk9-dependent, mechanism to control gene expression. Indeed, inhibiting the ability of NELF to associate with Pol II in flies has been found to reduce the effects of Cdk9 inhibition on pause intensity60. Further work will be required to determine if the four-subunit NELF complex is sufficient to induce Cdk9-regulated pausing in other eukaryotes, and whether reduced Pol II elongation rates would result from pause escape in the absence of both NELF and Cdk9.The observation that Pol II farther downstream of the TSS (~1 kb) was less, or possibly unaffected by Yeast strains are listed in Supplementary Table mcs6 and cdk9 AS mutant strains, each with single substitutions of the gatekeeper residues, Leu87 and Thr120, respectively, with Gly34. Although growth of both strains was sensitive to 3-MB-PP1, the mcs6L87G strain was ~5-fold less sensitive than was the cdk9T120G strain34. By standard methods for gene replacement in S. pombe61, we introduced structure-guided second site mutations to attempt to optimize performance of mcs6as alleles62 and obtained a double-point mutant, mcs6N84T/L87G (referred to hereafter as mcs6as5), which had a growth rate indistinguishable from that of wild-type cells in the absence of 3-MB-PP1, and was sensitive to growth inhibition by 3-MB-PP1 with an IC50 ≈ 5 µM, which was nearly identical to that of cdk9as  lysis method63. Briefly, cultures were grown to a density of ~1.2 × 107 cells/mL in YES media at 30 °C prior to treatment. After the specified duration of treatment with of 3-MB-PP1 or DMSO, ~ 6 × 107 cells were transferred into 500 μl 100% (w/v) TCA and collected by centrifugation. Pellets were resuspended in 20% (w/v) TCA and protein extracts were prepared using a bead beater to mechanically lyse cells with glass beads. Antibodies used in this study recognized Spt5-P (used at 1:2000) or total Spt5 (used at 1:500)64, total Pol II , Pol II Ser2-P , Pol II Ser5-P , Pol II Ser7-P , and α-tubulin . Raw images of all scanned western blots used in this work are provided , before treatments. Prior to treatment, all culture volumes were adjusted (based on OD600) to have an equal number of S. pombe cells (in 10 mL volume). After normalization for cell number, a fixed amount of thawed S. cerevisiae (50 µL OD600 = 0.68) culture was spiked in to each sample. Treatments were performed by adding either 10 µM 3-MB-PP1 (stock concentration = 40 mM) or an equivalent volume of DMSO. Sample treatments were terminated by pouring into 30 mL ice-cold water, treated with diethyl pyrocarbonate DEPC. Samples were immediately spun down and processed for cell permeabilization as described below (see PRO-seq section).All samples were grown and treated in YES medium at 30 °C. Biological replicates were prepared by picking distinct colonies of a particular strain and growing them in separate liquid cultures. For each experiment, cultures were grown overnight and diluted to an optical density ODThe time course experiment was performed in two biological replicates, where all time points for a given replicate was performed on cells from the same large culture. Each time point treatment was performed as described above. In order to limit differences in time on ice before permeabilization, treatment time points were carried out in reverse order. The 2.5, 1, and 0.5-min treatments were performed separately to ensure accuracy of timing. The 0-min time point was treated with DMSO for 20 min to account for any possible effects of adding the solvent for the maximum duration.S. cerevisiae, cells were spun down and washed in cold H20. Cells were then permeabilized in 10 mL 0.5% sarkosyl at 4 °C, and incubated on ice for 20 min, then spun at a reduced RCF (400 × g) for 5 min at 4 °C. For run-on reactions, yeast pellets were resuspended in 120 μL of 2.5× run-on reaction buffer  with 6 μL 0.1 M DTT and 3.75 μL of each 1 mM biotin-NTP added immediately before use. The volume of the run-on reaction mix was brought to 285 μL with DEPC-treated H20. 15 μL 10% sarkosyl was then added to the reaction and the run-on was allowed to run on for 5 min at 30 °C. RNA was extracted using a hot phenol approach65; after the run-on reaction cells were pelleted at 400 × g for 5 min at 4 °C and quickly resuspended in 500 μL acid phenol. An equal volume of AES buffer  was added and placed at 65 °C for 5 min with periodic vortexing, followed by 5 min on ice. 200 μL chloroform was added and mixed followed by centrifugation at 14,000 × g for 5 min (4 °C). 3 M NaOAc was added to the aqueous layer (200 mM) followed by ethanol precipitation with 3× volume of 100% ethanol. The RNA pellet was air dried before being resuspended in 20 μL DEPC-treated water. The standard PRO-seq protocol66 was then followed, beginning with base hydrolysis, through to sequencing. For base hydrolysis of RNA, 5 μL 1 N NaOH was added to the resuspended RNA and left on ice for 10 min before being neutralized with 25 μL 1 M Tris-HCL pH 6.8. After passing hydrolyzed RNA through a P-30 column (Bio-Rad), RNA was incubated with streptavidin-conjugated magnetic beads  for 20 min at room temperature, then washed with 500 μL of each of the following: high salt solution , bead binding solution , low salt solution . Note that prior to incubation with sample RNA, M280 beads were washed once with a solution of 50 mM NaCl and 0.1 N NaOH, and then twice with 100 mM NaCl. After bead binding and washing, enriched nascent RNA was extracted using TRIzol (Ambion) and ethanol precipitated. An RNA adapter was then ligated to the 3′ ends of enriched RNAs (see below) using T4 RNA ligase 1 (NEB), followed by another round of bead binding, washing, and TRIzol extraction, as before. RNA 5′ ends were then biochemically prepared for adapter ligation by first removing any cap structures with RNA 5′ pyrophosphohydrolase , then restoring mono-phosphates to base hydrolyzed RNA 5′ ends with T4 polynucleotide kinase . T4 RNA ligase 1 was then used to ligate the 5′ RNA adapter (5′-CCUUGGCACCCGAGAAUUCCA-3′) to enriched RNAs. Following another round of bead binding and washing, as before, cDNA was prepared by reverse transcription using superscript reverse transcriptase III  and the primer, RP1 . Libraries were then amplified via PCR using Phusion (NEB) polymerase and Illumina oligos, RP1 and one indexed RNA PCR Primer . PCR products were run on a native polyacrylamide gel, size selected (~130–400 bp), extracted, quantified, and submitted to sequencing. More details regarding preparing PRO-seq libraries can be found in the published protocol for precision run-on and sequencing66.After experimental treatments and spiking in mcs6as5, or cdk9as, excluding the time course experiment, were prepared according to the standard procedure described above66. All libraries prepared from lsk1as mutant and mcs6as5cdk9as double mutant strains received a novel 3′ RNA adapter during the first RNA ligation step. This 3′ adapter contains a random 6 nt unique molecular identifier at the 5′ end that, although not used here, can be used during read processing to remove PCR duplicates (5′-/5Phos/NNNNNNGAUCGUCGGACUGUAGAACUCUGAAC/Inverted dT/-3′). For all time course samples prepared in cdk9as, as well as lsk1ascdk9as samples, a different 3′ RNA adapter was used. Here, the RNA oligonucleotide possessed a known hexanucleotide sequence preceded by a guanine at the 5′ end, which differed for each library prepared (5′-/5Phos/GNNNNNNGAUCGUCGGACUGUAGAACUCUGAAC-/Inverted dT/-3′). The ligation of this adapter with distinct barcodes to each library permitted the pooling of all libraries after the 3′ end ligation step and thus facilitated handling. After pooling, the remainder of the library preparation was carried out as described above, but in a single tube. After sequencing, the inline barcode was used to parse reads based on their sample of origin.For specific batches of experiments slight modifications to the library preparation were made. Samples prepared with wt, http://hannonlab.cshl.edu/fastx_toolkit/). Raw reads from each sample were processed by removing any instances of partial or complete matches to the 5′ adapter sequence (5′-TGGAATTCTCGGGTGCCAAGG-3′) with the fastx_clipper function. Next, if a 3′ molecular, or sample barcode was included during 3′ adapter ligation, this length was removed from the beginning of each read. All reads were then trimmed to a maximum remaining length of 36 nt using the fastx_trimmer function. With the fastx_reverse_complement function we retrieved the reverse complement sequence of each read. All downstream alignments were performed using Bowtie (version 1.0.0)67. Reverse complemented reads derived from ribosomal RNA genes were then removed through alignment to a genome consisting of ribosomal RNA genes from Saccharomyces cereivisiae (spike-in) and then S. pombe. To parse reads based on organism of origin, while eliminating reads of ambiguous origin, non-ribosomal DNA reads were then aligned to a combined genome, containing each chromosome from S. cerevisiae (sacCer3 = S288C_reference_genome_R64-1-1_20110203) and S. pombe (version: ASM294v2). Only uniquely mapping reads were considered for downstream analysis. Ultimately, the normalization factor for each library was calculated as: total spike-in mapped reads/105. Unique read alignments to the fission yeast genome were processed using the Bedtools68 function, genomeCoverageBed, to generate bedgraph formatted files, reporting the number of read 3′ ends (the last base incorporated) at each position across the genome. Bedgraph files were further converted to bigwig format for downstream analysis. Sequencing, alignment, and batch information for each sample can be found in Supplementary Table All sequencing was performed on an Illumina NextSeq 500 device. If samples were pooled during the library preparation using the novel 3′ RNA adapter described above, raw reads were parsed into their respective samples based on the inline barcode using fastx_barcode_splitter function from the FASTX-toolkit . The selected base was required to have >4 reads over background, while accounting for library size (annotations available at https://github.com/gregtbooth/Pombe_PROseq). All genes used in this work were required to have an observed TSS (n = 4654), and this observed TSS was used throughout. We then filtered genes for “activity” in our untreated, wild-type data. A gene was considered active if the read density within the gene body was significantly higher (p < 0.01) than a set of intergenic “background” regions, based on a Poisson distribution with λ = background density . Finally, to minimize the possible influence of read-through transcription from upstream genes, we filtered genes based on the relative amounts of PRO-seq signal directly upstream of the TSS. Thus, we only considered genes with more downstream (+250 to +550) relative to immediately upstream .Using previously published PRO-cap data43. Rather than using default between-sample normalization approaches, we supplied our own spike-in-based normalization factors  at each time point up to 5 min. Wave calling parameters were adjusted to accommodate the smaller genes of yeast. For calling advancing waves, the upstream region used was set to −500 to the observed TSS for each gene. Approximate wave distances, used to initialize the model, were set based on estimates derived from inspecting the fold-change heat maps . Wave distances were determined based on the difference in signal within 50-bp windows between treated and untreated. The wave quality was determined based on the criteria used previously46. We further required that for each gene, a wave must be called for all used time points (maximum 5 min) using the combined replicate data, and that the calls must not recede toward the TSS with time (n = 43).Both advancing and clearing waves were identified using a three state Hidden Markov Model (HMM) that was previously developed and implemented on GRO-seq data from a human cell linen = 42). Again, 50-nt windows were used to tile across genes; however, an additional smoothing parameter in the model arguments (TSmooth) was set to 5 as a way of restricting the effect of windows with outlying differences between treated and untreated. For each gene, we required that the model be able to identify a distinct clearing wave  at each time point that did not move toward the TSS with time (n = 32).To identify regions containing the faster population of Pol II , we adjusted the HMM to identify the region between the advancing and clearing waves. For this task, we used the first 1000 nt downstream of the observed TSS (within the advancing wave) to initialize the first state of the HMM. Approximate distances of the start of the clearing wave from the TSS were used to initialize the model and were set based on estimates derived from inspecting the fold-change heat maps . Unlike the model used to call the advancing wave, which assumed the upstream state to be normally distributed, all three states were presumed to behave according to distinct gamma distributions. Because of the fleeting nature of this population of polymerases observed in our time course, we were further limited to analysis of filtered genes longer than 6 kb , giving a distribution of means. A more conservative, gene-by-gene estimate of variance was derived simply from the distribution of rates based on individual measurements for each gene.https://github.com/gregtbooth/Pombe_PROseq.Custom scripts and alignment pipelines have been made publicly available through the following GitHub repository; The raw and processed sequencing files have been submitted to the NCBI Gene Expression Omnibus under accession GSE102308.Supplementary InformationPeer Review File