In this study, Weng et al. provide new insight into the mechanism of piRNA transcription. They used functional proteomics approaches to identify an upstream sequence transcription complex (USTC) that is essential for piRNA biogenesis, and their findings demonstrate the repurposing of a general transcription factor complex, USTC, to aid in genome defense against transposons. Caenorhabditis elegans, most piRNA precursors are transcribed from two genomic clusters that contain thousands of individual piRNA transcription units. While a few genes have been shown to be required for piRNA biogenesis, the mechanism of piRNA transcription remains elusive. Here we used functional proteomics approaches to identify an upstream sequence transcription complex (USTC) that is essential for piRNA biogenesis. The USTC contains piRNA silencing-defective 1 (PRDE-1), SNPC-4, twenty-one-U fouled-up 4 (TOFU-4), and TOFU-5. The USTC forms unique piRNA foci in germline nuclei and coats the piRNA cluster genomic loci. USTC factors associate with the Ruby motif just upstream of type I piRNA genes. USTC factors are also mutually dependent for binding to the piRNA clusters and forming the piRNA foci. Interestingly, USTC components bind differentially to piRNAs in the clusters and other noncoding RNA genes. These results reveal the USTC as a striking example of the repurposing of a general transcription factor complex to aid in genome defense against transposons.Piwi-interacting RNAs (piRNAs) engage Piwi proteins to suppress transposons and nonself nucleic acids and maintain genome integrity and are essential for fertility in a variety of organisms. In  Caenorhabditis elegans, piRNAs preserve genome integrity in the germline by recognizing and silencing “nonself” genomic loci such as transposons or other foreign nucleic acids and induce chromatin modifications  are a class of small (21- to 30-nucleotide [nt]) RNAs that associate with Piwi proteins, a highly conserved subclass of Argonaute proteins, and play significant roles in fertility and genome stability . piRNAs ications . In addial cells .C. elegans piRNAs are also referred to as 21U-RNAs, given that they are predominantly 21 nt in length and have a strong bias toward 5′ monophosphorylated uracil , the biogenesis of piRNAs is a Dicer-independent process . In Drosunctions . The mouunctions —pachytenunctions .C. elegans encodes two Piwi proteins: PRG-1 and PRG-2. Whereas the function of PRG-2 is unknown, mature piRNAs associate with PRG-1 to conduct their functions  that initiates precisely 2 nt upstream of the 5′ end of mature piRNAs to generate 25- to 29-nt capped small RNA (csRNA) precursors . Next, tunctions . The bin animals , and los animals . Interes animals .C. elegans. Type I piRNAs are predominantly transcribed from two broad regions on chromosome IV and contain an 8-nt upstream Ruby motif (CTGTTTCA) and a small YRNT motif in which the T corresponds to the first U of the piRNA. Type II piRNAs are present outside of chromosome IV and lack the Ruby motif . These proteins significantly separate together from the PRDE-1 knockout immunoprecipitation , assuming 1:1:1:1 stoichiometry . Altogether, these results raise strong evidence that PRDE-1, SNPC-4, TOFU-4, and TOFU-5 function as a complex during piRNA transcription.Our previous study identified  elegans . Later,  elegans . To furt elegans , were ampitation B. Thus, hiometry C. To idehiometry , we wanttofu-4 and tofu-5 mutant phenotypes, respectively, advocating that the tagged proteins could recapitulate the functions of endogenous proteins . Both TOFU-4::GFP and TOFU-5::GFP exhibit distinct foci in the germline nuclei . Previous work showed that PRDE-1 colocalizes with SNPC-4 and forms distinct foci in germline cell nuclei . To validate that PRDE-1, TOFU-4, and TOFU-5 function as a complex, we crossed the mCherry::PRDE-1 transgene into TOFU-4::GFP- and TOFU-5::GFP-expressing animals  transgenic strains using the Mos1-mediated single-copy insertion (MosSCI) technology . The TOFl nuclei . We immu animals D. ConsisSupplemental Fig. S3). We found that all four factors have similar genome-wide binding profiles, with strong enrichment on chromosome IV piRNA clusters , although SNPC-4 did not appear enriched on the smaller piRNA cluster (cluster I). However, this might be a reflection of lower signal to noise ratio of the SNPC-4 ChIP-seq experiments . Consistent with our previous finding that PRDE-1 is not required for type II piRNA transcription . However, SNPC-4 and TOFU-5 were enriched at type II piRNA genes . We observed that all four factors exhibit robust enrichment around type I piRNA genes  and prde-1(mj207) mutants and found that TOFU-5 failed to form the subnuclear foci in germline cells in young adult animals  balanced mutant animals and found that TOFU-5 fails to form the subnuclear foci in snpc-4(tm4568) homozygous mutant young adult offspring of these animals (snpc-4(tm4568) mutants, TOFU-5::GFP accumulates in the germline syncytium, further confirmed by knocking down snpc-4 by feeding RNAi . Unlike wild-type TOFU-5, the TOFU-5(*SANT)::GFP fails to form the piRNA foci and is instead enriched in the germline syncytium  mutants and found that PRDE-1 was required for the formation of TOFU-4 piRNA foci (snpc-4(tm4568/hT2) and tofu-5(tm6408/hT2) and found that TOFU-4 fails to form piRNA foci in snpc-4(tm4586) and tofu-5(tm6408) homozygous mutants, which was further confirmed by knocking down snpc-4 by feeding RNAi . These results suggest that, in addition to their ancestral functions, including ncRNA transcription, USTC components might have acquired other functions, such as promoting piRNA biogenesis  genes . We therogenesis .Supplemental Table S2; Supplemental Fig. S7A,B; Supplemental Table S2). However, we did not identify any forkhead transcription factors in this screen, which had been shown previously to recognize the Ruby motif .To further understand the function of the USTC in promoting piRNA transcription, we searched for factors that are required for the formation of TOFU-5 subnuclear piRNA foci. We selected a number of candidate genes from our proteomic experiments and previous genome-wide RNAi screens and carried out a focused candidate RNAi screen for TOFU-5 foci  and a small YRNT motif. Type II piRNAs lack the Ruby motif and are present outside of chromosome IV . We speculate that tbp-1 and the Ruby motif may be required together for USTC binding to piRNA promoters. Here, TBP-1 may act as a bridge to bend the DNA so that the USTC comes closer to the piRNA promoter sequences. Further investigation of the roles of TBP-1 in piRNA biogenesis will shed light on the specific mechanistic understanding of the transcription regulation of piRNAs.The function of Both SNPC-4 and TOFU-5 contain SANT domains . SequencDrosophila, transcription of piRNA clusters is enforced through RNA Pol II preinitiation complex formation within repressive heterochromatin, in which the heterochromatin protein-1 variant Rhino recruits Moonshiner  and set-16 (a gene encoding a putative H3K4 methyltransferase) were identified. However, these two proteins and other chromatin modification factors were not required for the formation of piRNA foci . Therefore, we speculate that either the biogenesis of piRNAs is independent of chromatin modification processes or certain chromatin factors act together to promote the formation of piRNA foci. Further analysis of the properties of chromatin, including histone modifications, in the germline will be necessary to directly explore the relationship between the chromatin state, the USTC, and piRNA expression.Chromatin modifications play important roles in small RNA biogenesis. In onshiner . In C. e regions . In our Supplemental Table S3.Bristol strain N2 was used as the standard wild-type strain. All strains were grown at 20°C unless specified. The strains used in this study are listed in Antibodies against PRDE-1 and TOFU-5 were produced at the Laboratory Animal Resources . For this, New Zealand White female rabbits were immunized with the recombinant full-length proteins together with complete Freund's adjuvant. After the priming immunization, four further booster immunizations were performed every 3 wk. The harvested serum was purified by affinity purification using the recombinant full-length PRDE-1 or TOFU-5, respectively.3VO4) and lysed in a FastPrep-24 5G homogenizer. The supernatant of lysate was incubated with homemade anti-GFP beads for 1 h at 4°C. The beads were then washed three times with cold lysis buffer. Proteins were eluted with chilled elution buffer (100 mM glycine-HCl at pH 2.5). Eluates were precipitated with TCA or cold acetone and dissolved in 100 mM Tris (pH 8.5) and 8 M urea. The proteins were reduced with TCEP, alkylated with IAA, and finally digested with trypsin overnight at 37°C. The liquid chromatography-tandem MS (LC-MS/MS) analysis of the resulting peptides and the MS data processing approaches were conducted as described previously ::GFP were resuspended in an equal volume of 2× lysis buffer  and frozen in liquid nitrogen. Qproteome cell compartment kit (Qiagen) was used to purify nuclear fractions that were diluted with PBSCM buffer . Protein concentration was measured with BCA protein measurement kit (Pierce). Anti-PRDE-1 antibody KSOJ (0.88 mg/mL) was coupled to 5 µg of Dynabeads per 1 mg of beads according to the manufacturer's instructions. Uncoupled IgG beads were used as a control. Beads were incubated with nuclear extracts overnight at 4°C and washed eight times with PBSCM, and proteins were eluted in 80 µL of elution buffer  by boiling for 30 min at 60°C. The elutes were precipitated in TCA or cold acetone and dissolved in 100 mM Tris (pH 8.5) and 8 M urea. The proteins were reduced with TCEP and alkylated with IAA. The samples were digested with trypsin overnight at 37°C, and each sample was run separately.The wild-type strain was var. Bristol N2 , and theC. elegans TOFU-4 protein fragment (amino acids 1–223), the full-length C. elegans TOFU-5 protein (amino acids 1–311), the full-length C. elegans PRDE-1 protein (amino acids 1–526), and the full-length C. elegans SNPC-4 protein (amino acids 1–928) (NCBI sequence reference NM_060011.4) were PCR-amplified and cloned into pB27 as a C-terminal fusion to LexA-DBD . Constructs were checked by sequencing the entire insert and used as bait to screen a random-primed C. elegans mixed-stage cDNA library constructed into pP6. pB27 and pP6 were derived from the original pBTM116  and L40:Gal4 (MAT) yeast strains as described previously  using a fully automated procedure. A confidence score  was attributed to each interaction as described previously , was injected into a Superose6 xK 16/70 . One-milliliter fractions (starting after the void volume of 40 mL) were collected, and proteins were precipitated with TCA, separated by gel electrophoresis, and transferred to a nitrocellulose membrane using a semidry transfer system (Bio-Rad).TOFU-4::GFP, a TOFU-4 promoter and CDS region were PCR-amplified with the primers 5′-GCAGATCTTCGAATGCATCGAATTGAAAATGAGAAAAAACTG-3′ and 5′-ATAGCTCCACCTCCACCTCCGAATTCTGCGTCTACCATCA-3′ from N2 genomic DNA. A GFP::3xFlag region was PCR-amplified with the primers 5′-GGAGGTGGAGGTGGAGCTATGAGTAAAGGAGAAGAAC-3′ and 5′-TCACTTGTCATCGTCATCCT-3′ from plasmid pSG085. A TOFU-4 3′ UTR (untranslated region) was PCR-amplified with the primers 5′-AGGATGACGATGACAAGTGAAAGAATTTTTATCCGAGTCAC-3′ and 5′-AGATATCCTGCAGGAATTCCGAAACTTGATTTTCAAAATTTA-3′ from N2 genomic DNA. The ClonExpress MultiS one-step cloning kit  was used to insert the TOFU-4::GFP::3xFlag fusion gene into the pCFJ151 vector. The transgene was integrated onto the C. elegans chromosome II by the MosSCI method  was used to insert the TOFU-5::GFP::3xFlag fusion gene into the pCFJ151 vector. The transgene was integrated onto the C. elegans chromosome II by the MosSCI system.For TOFU-5(*SANT)::GFP, plasmid was PCR-amplified with the primers 5′-TCGCAGCTTCATCGAAGAGATAAAGTAAGT-3′ and 5′-TCTCTTCGATGAAGCTGCGACCTTCTGCGA-3′ from TOFU-5::GFP::3xFlag plasmid. The transgene was integrated onto the C. elegans chromosome II by the MosSCI system.For RNAi experiments were conducted as described previously . Images ChIP experiments were performed as described previously with hypochlorite-isolated embryos or young adults . AnimalsThe DNA samples from ChIP experiments were sent to in-house sequencing for library preparation and sequencing. Briefly, 10–300 ng of ChIP DNA was combined with End Repair Mix and incubated for 30 min at 20°C followed by purification with a QIAquick PCR purification kit (Qiagen). The DNA was then incubated with A-tailing mix for 30 min at 37°C. The 3′ end adenylated DNA was incubated with the adapter in the ligation mix for 15 min at 20°C. The adapter-ligated DNA was amplified by several rounds of PCR amplification and purified using a 2% agarose gel to recover the target fragments. The average molecule length was analyzed on the Agilent 2100 bioanalyzer instrument (Agilent DNA 1000 Reagents). The library was quantified by qPCR (TaqMan probe). The libraries were further amplified on cBot to generate the clusters on the flow cell and sequenced with a single-end 50 method on a HiSeq1500 system.C. elegans genome using BWA version 0.7.7 . The same summed inputs were used for BEADS normalization and peak calls. We observed that despite using input files for MACS2  and filtq-value cut-off and a fragment size of 150 bp against summed ChIP-seq input. To generate sharp peak call sets, we obtained peak summits and extended them 150 bp upstream and downstream, creating 300-bp regions around summit calls. Furthermore, we combined ChIP-seq replicates by intersecting these regions and setting the final peak size back to 300 bp. Finally, peaks overlapping nonmappable (GEM mappability <25%) or blacklisted regions were discarded. This produced sharp uniform peaks suitable for further quantitative analyses.ChIP-seq peaks were called using MACS2 version 2.1.1  with a pPRDE-1, SNPC-4, TOFU-4, and TOFU-5 peak calls were classified using selected annotations from Ensembl version 92. The peak was assigned to a given class if it directly overlapped with annotated loci. The assignments were exclusive (a peak could be assigned to only a single class), giving the following order of priority: snRNA, snoRNA, transfer RNA (tRNA), miRNA, large intergenic ncRNA (lincRNA), ncRNA, promoter, and gene body. Promoters were defined as 500 bases upstream of annotated TSSs. The overlap significance was estimated using a hypergeometric test for overrepresentation.SeqPlots  software2-scaled signal in 1-kb bins divided into piRNA clusters and chromosomes I, II, III, and V. The signal was obtained using the bigWigSummary utility from the Kent library . Genomic coordinates of piRNA genes were obtained by SAMtools against the C. elegans ce10 genome assembly. Type II piRNA genes were obtained from a previous publication (piRNA annotations were downloaded from the piRBase online database (lication . Type I


Komagataella phaffii (Pichia pastoris) was attempted by engineering a strong methanol-inducible DAS1 promoter using Citrobacter braakii phytase production as a model case. Constitutive expression of KpTRM1, formerly PRM1—a positive transcription regulator for methanol-utilization (MUT) genes of K. phaffii,was demonstrated to produce phytase without addition of methanol, especially when a DAS1 promoter was used but not an AOX1 promoter. Another positive regulator, Mxr1p, did not have the same effect on the DAS1 promoter, while it was more effective than KpTrmp1 on the AOX1 promoter. Removing a potential upstream repression sequence (URS) and multiplying UAS1DAS1 in the DAS1 promoter significantly enhanced the yield of C. braakii phytase with methanol-feeding, which surpassed the native AOX1 promoter by 80%. However, multiplying UAS1DAS1 did not affect the yield of methanol-free expression by constitutive KpTrm1p. Another important region to enhance the effect of KpTrm1p under a methanol-free condition was identified in the DAS1 promoter, and was termed ESPDAS1. Nevertheless, methanol-free phytase production using an engineered DAS1 promoter outperformed phytase production with the GAP promoter by 25%. Difference in regulation by known transcription factors on the AOX1 promoter and the DAS1 promoter was also illustrated.The construction of a methanol-free expression system of  DAS1 promoter to develop an efficient methanol-free expression system in K. phaffii (P. pastoris).Engineering the methanol-inducible  AOXalcohol oxidaseDASdihydroxyacetone synthaseMUTmethanol-utilizationUASupstream activation sequenceURSupstream repression sequenceKomagataella phaffii (formerly Pichia pastoris), Ogataea methanolica (P. methanolica), O. polymorpha (Hansenula polymorpha) and Candida boidinii possess a common methanol utilization (MUT) metabolic pathway  that does not require methanol induction is also used in K. phaffii for protein production  De-repression through the depletion of repressive carbon source. Once the repressive carbon source is depleted, the AOX1 promoter is weakly induced. iii) Methanol induction. The AOX1 promoter is fully induced in the presence of methanol. Transcription factors responsible for i) are negative regulators while the ones responsible for ii) or iii) are positive regulators.The regulation of the cis-elements) in the promoter of a targeted gene and bind it through the DNA binding domain to regulate the transcription of the gene. The DNA binding domain is classified into three domains, namely, the zinc-containing domain, such as C2H2 zinc finger (Adr1 type) and Zn2Cys6 zinc cluster ; helix-turn-helix (Matα2 type); and the zipper-type domain (Gcn4 type)   in K. phaffii over 50 times. In a fungus Aspergillus oryzae, 12 tandem repeats of region III, which is a conserved region among amylase genes in A. oryzae, increased the expression of the agdA promoter >20-fold  containing the . et al.  reportedK. phaffii, using another strong methanol-inducible DAS1 promoter through engineering its transcription factors, namely KpTrm1p, which was applied to the production of an industrial enzyme in high-cell-density fermentation. Multiplication of the UAS of the DAS1 promoter was also tested in order to increase enzyme productivity with and without methanol. We also discuss our findings on the difference in the regulation of the AOX1 promoter and the DAS1 promoter by known transcription factors in K. phaffii.Here, we report on our attempt to develop a methanol-free expression system of K. phaffii (Pichia pastoris) GS115 , KM71  and expression vectors pPIC9K, pGAPZαA were purchased from Invitrogen . Escherichia coli cloning vector pCR2.1-TOPO  and pT7Blue-Novagen  and cloning host E. coli DH5α , TOP10 (Life Technologies), XL10  were purchased from suppliers. Yeast strains were grown on YPD  unless specified. Media used for phytase production were described below. E. coli strains were grown in LB  with relevant antibiotics. Other strains and plasmids used in this study are summarized in + and MutS represent methanol-utilization plus and methanol-utilization slow phenotype, respectively.Citrobacter braakii phytase G01651 and its expression plasmid with AOX1 promoter, pPICNoT-G01651, was described in US 8.236,528 B2  encoding dihydroxyacetone synthase was identified in the K. phaffii genome sequence provided by Integrated Genomics, Inc.  (DAS1 gene (GenBank GZ456654) was isolated from K. phaffii GS115 and used for construction of phytase expression plasmid pNo-DP3 , termed KpTRM1, formerly called PRM1 (GenBank GZ456640), and another positive regulation factor MXR1 . RD medium , 4 × 10−5% biotin, 0.005% each amino acid of L-glutamic acid, L-methionine, L-lysine, L-leucine, L-isoleucine) supplemented with 2% Difco Agar Noble or MD medium  supplemented with 1 M sorbitol and 2% agar noble was used for regeneration with his4 selection. In the case of Zeocin resistance selection, YPDS + Zeocin agar ) was used for the regeneration. Plates were incubated at 28 or 30°C for 3–4 days until colonies appeared. Screening for Mut+/MutS phenotypes was performed using MD (minimum dextrose) medium plates and MM (minimum methanol) medium plates following the published protocol .HIS4 in the host genome, and primer H2: 5’-GCCGCCCAGTCCTGCTCGCT-3’based on the sequence in the expression plasmids. The strains in which the expression cassette was integrated at HIS4 locus generated a 2.9 kb band.A small portion of a colony grown on agar medium was picked with a sterile toothpick and transferred into a small tube, then heated in a microwave oven for one minute. The dried cells were suspended in 50 μL sterilized water and subjected to PCR. The reaction mixture was 20 μL including 2 mM dNTP, 10 μM of each primer, 1 unit of Expand high fidelity plus , 1 × Expand high fidelity plus buffer and 1 μL of cell suspension as mentioned above. The PCR primers were primer H1: 5’-CTGCTCTAGCCAGTTTGCTG -3’ corresponding the sequence upstream of et al. Isolated transformants were grown in liquid medium to evaluate the phytase expression level. In the case of methanol induction, strains were grown in YPD with vigorous shaking at 30°C for 2 days, then methanol was added at the final concentration 0.5–2% (V/V) to induce the expression. Activity measurement was done using 3 days’ sample. In the case of expression without methanol, strains were grown in YPD with vigorous shaking. Glucose in YPD medium was usually consumed within 2–3 days in the tested condition. Two days’ or 3 days’ samples were used for activity measurement. Phytase activity was measured as described before  with some modification. Fermentation was conducted in 5 L-fermenters  starting with 2 liters of basal medium . Strains were pre-cultured in 110 mL of YPD at 30°C for 1 day, then transferred to the basal medium supplemented with PTM1 Trace Salts (Invitrogen). The fermentation was controlled at 30°C, pH above 5.0 and keeping dissolved oxygen (DO) level over 30% by adjusting agitation speed. Glycerol feeding started at 12 hours and the feeding rate was increased step-by-step until 40 hours. When the DO level decreased below 10%, feeding was stopped until the DO recovered. In the case of the methanol fermentation, methanol feeding was started at 40 hours. The methanol-feeding rate after 41 hours was adjusted aiming to maintain the methanol concentration in the medium at 0.8%. When the DO level decreased below 20%, methanol feeding was stopped until the DO recovered and then the feeding rate was decreased to keep DO level over 20%. In the case of glucose fermentation, initial carbon source in the basal medium and feeding substrate was glucose instead of glycerol. Glucose feeding started at 12 hours and feeding rate was increased step-by-step until 40 hours. After 40 hours, the feeding rate was controlled to keep the DO level between 10–30%. Residual glucose in the medium was usually below 0.2 g/L after ∼70 hours. Dried cell weight (DCW) was measured as follows; ∼2 mL of culture broth was transferred to a pre-weighed glass tube and the tube was again weighed to calculate a weight of the whole broth. Cells were collected from the broth by centrifugation at 3500 rpm for 10 minutes, then washed twice with Reverse Osmosis (RO) water. After discarding the washed water, cells in the tube were dried in drying oven at 105°C for at least 24 hours. After cooling in a desiccator, the tube was weighed to calculate the weight of dried cells. Dried cell weight (DCW/g) was obtained as a quotient of the weight of dried cells divided by the weight of whole broth. Phytase activity was measured using the culture supernatant after centrifugation.High-cell-density fermentation was performed following the manufacturer's protocol  phenotype and maintained the cell mass during the enzyme production phase  phenotype of the strain. However, the cell mass of strain DAS40 did not increase during the methanol feeding phase   . Surprisingly, the effect of the pGPrm on the DAS1 promoter was much more significant than its effect on the AOX1 promoter. Selected co-expressing strain DPrm11, producing phytase with the DAS1 promoter, was evaluated in high-cell-density fermentation with glucose feeding in order to compare it with reference strain DAS40  promoter was partially activated under de-repressed conditions without methanol feeding, while the DAS1 promoter was not activated under de-repressed conditions and required the addition of methanol. Yurimoto, Oku and Sakai  to the methanol-specific induction of the DAS1 promoter was smaller than that of Trm1p in C. boidinii. Our results suggest that it is probably the same in K. phaffii, i.e. the DAS1 promoter is mainly activated by methanol-induction specific KpTrm1p, and Mxr1p alone does not induce the DAS1 promoter . Their results indicate that Mit1p is the true methanol-induction-specific regulator for AOX1, rather than KpTrm1p. Wang . Li et al.  and deletion of the URS increased phytase expression. Ohi et al. (AOX2 promoter of K. phaffii (P. pastoris), at around −645 and −255, and that removing or mutating these URSs increased the expression level dramatically.Aiming to engineer a methanol-inducible promoter to enhance promoter activity Fig. , we attei et al.  reportedDAS1 promoter  and MRE2 (TGCATTCCTAAAATAG)—existed in the DAS1 promoter of C. boidinii and that MRE1 was closely related to the induction by Trm1p. Similar sequences were identified in the UAS1DAS1 in K. phaffii, termed PBS1 and PBS2, and their deletion was examined  showed higher phytase activity than the plasmids maintaining UAS1DAS1 (pPd-17∼19). The results suggest that UAS1DAS1 was not a cis-element of KpTrm1p for activating the DAS1 promoter in the tested methanol-free expression system. This unexpected result indicated that there must be another element in the DAS1 promoter for KpTrm1p to interact with. Careful analysis revealed that the region between −455 and −355 was more important to maximizing the effect of KpTrm1p . In fact, strains carrying these plasmids, except 19P-13 with pDd-19, showed similar levels of phytase productivity to DPrm11 carrying the wild type DAS1 promoter. The reason for the lower yield of 19P-13 is not known, but it might be related to the region deleted in plasmid pDd-19. Moreover, there must be another regulation factor that interacts with UAS1DAS1 to enhance methanol activity. Determining whether it is Mit1p requires further study.Co-expression of constitutive DAS1 promoter, with 3xUAS1DAS1 for phytase production. Selected strain 28P-14, derived from strain 28-2, and strain 2P-4, derived from reference strain 2-3, were evaluated in high-cell-density fermentations under methanol-free conditions (Fig. DAS1 does not interact with KpTrm1p in the tested methanol-free fermentation. Interestingly, the phytase yields obtained with these two strains were three times higher than the previously constructed DPrm11 derived from strain DAS40, carrying pNo-DP3 with the original DAS1 promoter of 1-kb for phytase expression. This might be because the truncated version of the DAS1 promoter was used for the construction of pDd-2 and pDd-28. One possible reason could be that the removed −855∼−1055 region contained URS—which affects the promoter activity negatively, even without methanol. Glucose repression could be an example of such a negative effect, which would have a significant impact, especially in glucose fermentation. Ohi et al. (AOX2 promoter were involved in catabolite repression, and the mechanism of their repressing effect was not defined. Another possible reason for the yield increase could be the integration locus of the expression plasmid. In the case of 2-3 and 28-2, plasmids pDd-2 and pDd-28 were integrated at the HIS4 locus, while in the case of DAS40, pNo-DP3 was integrated at the DAS1 promoter region. To elucidate mechanisms of improvement, further experiments are awaited. Nevertheless, the newly developed methanol-free expression system based on a truncated version of the DAS1 promoter produced 25% more phytase than the system based on the GAP promoter (Fig. DAS1 on the effect of a methanol-free system is awaited.Constitutive KpTrm1p was also tested in strain 28-2 carrying the ons Fig. . As expei et al. , howeverter Fig. . It was AOX1 and DAS1, in K. phaffii was different (Fig. DAS1, single transcription factor KpTrm1p was enough to activate the DAS1 promoter, while AOX1 required another transcription factor, such as Mit1p or Mxr1p, to fully activate its promoter. This difference made it possible to develop a methanol-free expression system using the DAS1 promoter and KpTrmp1, but not the AOX1 promoter.Results obtained during our research revealed that transcriptional regulation between two MUT genes, ent Fig. . In the DAS1 and removing URS in the DAS1 promoter enhanced methanol induction of the DAS1 promoter significantly. However, it turned out that UASDAS1 was not a cis-element of KpTrm1p, and its amplification did not affect methanol-free expression by constitutive KpTrmp1. Another region termed ESPDAS1 was identified instead as a potential cis-element of KpTrm1p. Nevertheless, the engineered DAS1 promoter combined with constitutive KpTrm1p successfully surpassed phytase production with the GAP promoter.Multiplying UASfoz059_Supplemental_FileClick here for additional data file.


Vascular calcification is the deposition of hydroxyapatite crystals in the blood vessel wall. Osteogenic differentiation of vascular smooth muscle cells (VSMCs) plays a key role in this process. Increased expression of alkaline phosphatase (ALP) occurs in some in vitro models of VSMC calcification and is thought to be crucial for mineralization, however, little is known about the transcriptional regulation of ALP in VSMCs. Recently, ALP upregulation was shown to coincide with endoplasmic reticulum (ER) stress-mediated vascular calcification, specifically with expression of the transcription factor ATF4. As no direct links between ALP expression and ER stress have previously been demonstrated in VSMCs, the aim of this study was to investigate whether ATF4 interacts directly with the ALP promoter.The present study shows that ALP mRNA and activity were significantly increased by ER stress treatment of human primary VSMCs in vitro and that this was ATF4-dependent. Bioinformatics analysis predicted two ATF4 binding sites in ER-stress responsive regions of the ALP promoter (− 3631 to − 2048 bp from the first intron). However, we found that ATF4 does not bind within this fragment of the ALP promoter region.The online version of this article (10.1186/s13104-018-3582-4) contains supplementary material, which is available to authorized users. Vascular calcification is the deposition of hydroxyapatite crystals in the blood vessel wall , 2. The Alkaline phosphatase , a key inducer of mineralization in bone, catalyses dephosphorylation of pyrophosphate resulting in inactivation of this calcification inhibitor and providing phosphate ions for hydroxyapatite crystal formation. ALP is ubiquitously expressed in tissues and increased levels in serum and the vasculature have been linked to vascular calcification , 9. IncrThe endoplasmic reticulum (ER) is an organelle where secreted and transmembrane proteins are folded and mature. ER stress occurs when the influx of unfolded proteins to the ER is larger than its capacity to fold them, resulting in activation of a signalling pathway called the unfolded protein response (UPR) . The UPRExpanded “Materials and Methods” are available in Additional file Human primary VSMCs were of aortic origin, collected from an adult donor with local ethics committee approval , characterised and archived in the laboratory. All human materials were handled in compliance with the Human Tissue Act . Cells were treated with 0.2 μg/ml thapsigargin  or 0.4 μg/ml tunicamycin . SiRNA gene knockdown was carried out using HiPerfect (Qiagen) and 3 pmol siRNA oligonucleotide smartpool (GE Dharmacon).Gene expression was analysed with quantitative real time PCR  and western blotting .ALP activity was measured in cell lysates colorimentrically at 405 nm using pNPP substrate (Sigma), normalised to protein concentration and expressed as μM of pNP generated per minute.The sequence corresponding to the − 122 to − 4556 (counting from the first nucleotide of the initiation codon) of the ALP promoter was analysed with Matinspector (Genomatix) for the presence of transcription factor binding sites. ALP promoter constructs  and pRL-Samples were analysed using liquid chromatography–tandem mass spectrometry (LC–MS/MS). Raw mass spectrometry data were analysed in Proteome Discoverer  utilising the Mascot database. Samples were searched against Uniprot database to identify proteins.All results represent 3 independent experiments, unless stated otherwise. Graphs show mean with SEM. Where appropriate t-tests or one way ANOVA with Tukey’s or Dunnett’s post hoc tests were performed. Statistical significance is indicated with asterisks: * denotes p between 0.05 and 0.01, ** denotes p between 0.01 and 0.001, *** denotes p < 0.001.ALP mRNA levels were upregulated in VSMCs treated with ER stress inducers tunicamycin (TM) and thapsigargin (TG) Fig. a. ER strSiRNA knock-down of IRE1, PERK and ATF6 caused small, but insignificant decreases in ALP expression. In contrast, knock-down of ATF4 caused a twofold decrease in ALP mRNA expression  and which suppress (silencers) transcription of the ALP promoter Fig. b. PotentTo examine known ER stress responsive consensus binding sites within the ALP promoter bioinformatics analysis with Genomatix Matinspector was performed. Several binding sites for ATF6, ATF4, CHOP and XBP1 were predicted in the full length sequence Fig. The promoter fragment containing both potential ATF4 binding sites Fig. c despiteFor more sensitive detection we subjected eluted proteins to liquid chromatography–mass spectrometry (LC–MS/MS)  bioinformatics predictions may not be accurate, (2) ATF4 may not be a direct regulator of ALP expression, but could act via other, unknown downstream factors that were downregulated after ATF4 knock-down and (3) the ATF4-responsive element could be located outside the examined promoter fragment . This isOur results also suggest that tunicamycin and thapsigargin regulate ALP via different pathways and this is in line with studies showing that in VSMCs these two compounds can have differential effects on gene expression .The proteomics results also suggest indirect regulation by ER stress. The responsive fragments contained a TCF/LEF binding site  and β-cThe main limitation of this study is the lack of consistent data for ALP promoter activity after ATF4 knock-down. However, using a direct approach of proteomics we did not find ATF4 binding the examined fragment of ALP promoter. It is also possible that ATF4 is a regulator of the ALP promoter, but our analysis did not encompass the right region of the promoter.Additional file 1: Figure S1. The sequence of the ALP promoter with ER stress-related transcription factor binding sites. Figure S2. Coomassie gel used for proteomic analysis of proteins bound to the ALP promoter. Full description of materials and methods.Additional file 2. Proteomics results. List of proteins, that bind the ALP promoter, identified by mass spectrometry. Nuclear extracts were bound and eluted off the biotinylated DNA on beads and analysed by LC–MS/MS. Raw mass spectrometry data were analysed in Proteome Discoverer  utilising the Mascot database. Samples were searched against Uniprot database to identify proteins bound to the ALP promoter and compared with proteins that nonspecifically bound to unconjugated beads.


CLIC4 between tumor cells and the surrounding stroma during cancer progression has been suggested to have a tumor-promoting effect. However, little is known about the transcriptional regulation of CLIC4. To better understand how this gene is regulated, the promoter region of CLIC4 was analyzed. We found that a high GC content near the transcriptional start site (TSS) might form an alternative G-quadruplex (G4) structure. Nuclear magnetic resonance spectroscopy (NMR) confirmed their formation in vitro. The reporter assay showed that one of the G4 structures exerted a regulatory role in gene transcription. When the G4-forming sequence was mutated to disrupt the G4 structure, the transcription activity dropped. To examine whether this G4 structure actually has an influence on gene transcription in the chromosome, we utilized the CRISPR/Cas9 system to edit the G4-forming sequence within the CLIC4 promoter in the cell genome. The pop-in/pop-out strategy was adopted to isolate the precisely-edited A375 cell clone. In CRISPR-modified A375 cell clones whose G4 was disrupted, there was a decrease in the endogenous CLIC4 messenger RNA (mRNA) expression level. In conclusion, we found that the G4 structure in the CLIC4 promoter might play an important role in regulating the level of transcription.The differential transcriptional expression of  Chloride intracellular channel 4, CLIC4, is a multifunctional protein. In addition to its diverse physiological functions ,2,3,4, tCLIC4 promoter [2+-induced keratinocyte differentiation [CLIC4 promoter is still unclear.The regulation of gene expression includes multiple mechanisms such as transcription factor binding  and epigpromoter ,14 and Sntiation . NeverthGuanine-rich (G-rich) sequences containing four runs of G-tracts can adopt four-stranded structures, named the G-quadruplex (G4). Four guanine bases can assemble into a square planar structure via Hoogsteen-type hydrogen bonds, and this is called a G-quartet. A stack of G-quartets is stabilized by monovalent cations, such as potassium or sodium, and this forms a tetrahelical G4 structure. The loop sequences connecting G-tracts determine the types of G4 topology . G4 archCLIC4 promoter, we found that high GC contents could form G4 structures. We further evaluated the effect of the G4 structure in regulating CLIC4 gene expression.G4s were found predominately in telomeres, and in the gene regulatory region of the human genome . ParticuCLIC4 promoter, a series of truncated CLIC4 promoter sequences from −1700 to +285, relative to the transcription start site  were cloned into the secreted embryonic alkaline phosphatase (SEAP) reporter vector pSEAP2-basic, and transfected into A375 cells to estimate the promoter activity of each region. As shown in To examine the regulatory region of the CLIC4 promoter region, we argued that DNA secondary structure might exist to interfere with the PCR and the sequencing process. Therefore, this prompted us to analyze its promoter sequence with DNA secondary structure. The QGRS program [CLIC4 p is shown in During the reporter plasmid constructions with different promoter regions, we found that the PCR and sequencing reactions were hampered. Since high GC contents were found in the  program  predicteCLIC4 promoter region, we examined three G-rich sequences with a higher potential to form G4 structures, and named them as putative G-quadruplex-1, -2, and -3 , as shown in CLIC4 promoter is presented in + to induce G4 formation, implying the formation of G4 structure in all of these three PG4s. Furthermore, in NMR spectra  comprising five G-tracts in the antisense strand of the CLIC4 promoter. Three mutants were designed to disrupt the quadruplex formation. Mutant No. 1 was generated by changing 78G and 52G in the first and last G-tracts of the PG4-3 sequence to 78T and 52T, respectively. Mutant No. 1 was unable to form G4 as revealed by CD spectra with the absence of ~265 nm or ~290 nm bands in 150 mM K+ solution  that we had constructed. As shown in Despite the disappearance of the G4 NMR signal in the synthetic Mutant No. 2 oligonucleotide, the reporter construct containing Mutant No. 2 sequence still exhibited normal transcription activity. We therefore further examined this contradictory result between the reporter construct and the synthetic nucleotide. Except for the five G-tracts in PG4-3, we found another G-tract at +39 to +41 immediately adjacent to PG4-3. We argued that this additional G-tract might be incorporated with the remaining three G-tracts, and it may have participated in forming another G4 structure in the reporter construct when the first two G-tracts were mutated in Mutant No. 2. To address the possibility that the Mutant No. 2 reporter construct still contained a G4-forming sequence, the sequence from +67 to +39, which included three remaining G-tracts and the additional G-tract, was synthesized and analyzed by NMR. As shown in CLIC4 transcription, we designed another mutant that we named loop-3T, in which only the non-G-tract sites: 76A, 70G, and 56C in the loops were mutated, respectively, to 76T, 70T, and 55T, which would not destroy G4-forming elements. The NMR result in CLIC4 transcription.To further examine the importance of the PG4-3 G4 structure in regulating CLIC4 expression, we used software to predict the possible candidate which could binds to the nucleotide sequences of PG4-3. SP1 and MAZ are the two candidate genes predicted to bind on this sequence and also have been reported to bind on G4 structure [CLIC4 mRNA expression nor transcription activity . A lower level of endogenous mRNA expression was found in genome-edited cell clones carrying Mutant No. 3 that disrupted the PG4-3 structure -mediated transcription. With regard to this, the formation of G4 could be the obstacle for RNAP, and the disruption of the G4 structure would result in the upregulation of transcriptional activity. However, as shown in VEGF) and endonuclease III-like protein 1 (NTHL1) promoters were modifiable by 8-oxo-7,8-dihydroguanine (OG) [CLIC4 is also a gene that can be regulated by reactive oxygen species (ROS)-induced oxidative stress. PDT-induced ROS downregulated CLIC4 transcription [CLIC4 transcription [There are five G-tracts in the originally identified PG4-3 sequence. In Mutant No. 2, we noticed that the additional G-tract might incorporate in the G4 formation when the first two G-tracts were mutated. The G4-forming sequence with a fifth G-tract has been observed in many oncogene promoters, and the ‘spare tires’ hypothesis was proposed . For insine (OG) ,36, and cription , while Tcription . Whether2.Human melanoma A375 cells  were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), and grown at 37 °C under 5% CO+ concentration of 150 mM. The observed signals were baseline subtracted.CD experiments were conducted using a spectropolarimeter  with a bandwidth of 2 nm at a scan speed of 50 nm/min and a step resolution of 0.2 nm over the spectral range of 210–350 nm. The DNA sample concentrations were 4 μM in 10 mM Tris-HCl (pH 7.5), and a stock solution of 3 M KCl  was added to the DNA samples to reach a final K2O in 10 mM Tris-HCl (pH 7.5) or 150 mM K+ conditions with an internal reference of 0.01 mM DSS , and they were analyzed by Bruker AVIII  500 MHz spectrometers equipped with a prodigy probehead, and on a Bruker AVIII 800 MHz NMR spectrometer equipped with a cryoprobe at 25 °C. 1D imino proton NMR spectra were recorded using a WATERGATE for water suppression.The unlabeled oligonuleotides synthesized by Bio Basic  were prepared to 100 μM in 10 mM Tris-HCl (pH 7.5) with or without 150 mM KCl, followed by denaturing at 95 °C for 5 min and slowly annealed to 25 °C. The strand concentrations of the NMR samples were 100 μM containing 10% DCLIC4 sequence from −1700 to 285 relative to the transcription start site (+1) was generated from genomic DNA of A375 cells, and cloned into the pSEAP2-Basic vector . After transfection in A375 cells with TurboFect  for 48 to 72 h, the culture medium was collected and analyzed for SEAP activity by measurement of the hydrolysis of p-nitrophenyl phosphate (pNpp) with a spectrophometer at OD405. MTT assay was used for the normalization of cell numbers.The CLIC4 sgRNA targeting near CLIC4 PG4-3 was selected and synthesized by in vitro transcription. HDR-1 donor templates containing different fluorescence cassettes: GFP, mCherry, and BFP, driven by a CMV promoter with a lacZ sequence on both ends (kindly provided by Dr. Steve Lin) flanking a 1 kb sequence of CLIC4 homology arms upstream and downstream of the PG4-3 Mutant No. 3 mutation site were generated by Gibson assembly. Cas9, CLIC4 sgRNA, and three fluorescence donor templates were introduced to A375 cells by nucleofection with the 4D-NucleofectorTM system and SF kit  under the FF-120 program. Single cells co-expressing three fluorescence markers were isolated by FACS for clonal expansion, and the genomic sizes of the sequences containing the fluorescence cassettes were confirmed by PCR. In the pop-out step, Cas9, sgRNA targeting the lacZ sites, and the HDR-2 donor template only containing the CLIC4 homology arms with the mutation sites, were again nucleofected in the pop-in cell clone. Single cells without fluorescence were sorted by FACS. The genomic DNA edited into the Mutant No. 3 sequence was analyzed by PCR and Custom TaqMan® Gene Expression Assays, SM ID: APFVMGD, which was designed to specifically anneal to the CLIC4 mutation site. Cell clones harboring the mutant sequence were further confirmed by sequencing.® Green Master Mix Kit  with the ABI StepOne system (Thermo Fisher Scientific). The primer sequences were as follows: CLIC4 (sense), 5′-GCAGTGATGGTGAAAGCATAG-3′; CLIC4 (anti-sense), 5′-TATAAATGGTGGGTGGGTCC-3′; GAPDH (sense), 5′-GACCACAGTCCATGCCATCA-3′; GAPDH (anti-sense), 5′-GTCCACCACCCTGTTGCTGTA-3′.Total RNA was extracted using TRIzol reagent  following the manufacturer’s instructions accordingly. A total of 1 µg RNA was used to synthesize complementary DNA (cDNA) by reverse transcription. The cDNA product was used as a template for real-time PCR analysis using the ABI Fast SYBRt-test was used to compare the differences between pairs of means. p < 0.05 was considered significant.All results were obtained from three independent experiments, and each value was expressed as the mean ± SD. The two-tailed Student’s CLIC4 promoter region may act as a regulatory element in regulating CLIC4 gene transcription, as shown in the reporter assay, as well as in the CRISPR-modified A375 cell clone with mutated PG4-3.The G4 structure formed by PG4-3 in the


TIMING OF CAB EXPRESSION 1 (TOC1) Arabidopsis clock gene. However, the chromatin-related modifiers responsible for these marks remain largely unknown. Here, we uncover that the chromatin modifier HISTONE DEACETYLASE 9 (HDA9) and the Evening complex (EC) component EARLY FLOWERING 3 (ELF3) directly interact to regulate the declining phase of TOC1 after its peak expression. We found that HDA9 specifically binds to the TOC1 promoter through the interaction with ELF3. The EC-HDA9 complex promotes H3 deacetylation at the TOC1 locus, contributing to suppressing TOC1 expression during the night, the time of EC function. Therefore, we have identified the mechanism by which the circadian clock intertwines with chromatin-related components to shape the circadian waveforms of gene expression in Arabidopsis.Circadian clocks are conserved time-keeper mechanisms in some prokaryotes and higher eukaryotes. Chromatin modification is emerging as key regulatory mechanism for refining core clock gene expression. Rhythmic changes in histone marks are closely associated to the  TOC1. This study illustrates how the circadian clock cooperates with histone deacetylase to regulate the circadian gene expression in Arabidopsis.Kyounghee Lee et al. show that a chromatin modifier HDA9 and a component of the Evening Complex, ELF3, cooperatively regulate the declining phase of Arabidopsis clock gene  Arabidopsis, two single-MYB transcription factors, CIRCADIAN CLOCK–ASSOCIATED 1 (CCA1) and LATE ELONGATED HYPOCOTYL (LHY), repress transcription of the TOC1/PSEUDO RESPONSE REGULATOR 1 (PRR1) that in turn represses CCA1 and LHY expression, constituting the central loop2. The central loop is interlocked with additional transcriptional loops. In the morning, a transcriptional repressing wave includes additional members of the PRR family including PRR5, PRR7, and PRR9 and CCA1 and LHY4. Evening-expressed clock components such as TOC1 and the EC contribute to the repression of the morning-expressed genes8. Additional transcriptional regulation also underlies circadian frameworks11. Moreover, accumulating evidence suggests that multiple layers of regulation such as alternative splicing, controlled protein turnover, and posttranslational modification further contribute to a precise rhythmic oscillation14.Circadian clocks generate biological rhythms with a period of 24 h and control plant growth and development in synchronization with the environmental cycles. Multiple transcriptional feedback loops define the basic architecture of the plant circadian clock. In 15. In particular, histone acetylation is implicated primarily in transcriptional activation16. Histone acetyltransferases (HATs) catalyze the addition of acetyl groups to lysine residues at histones, neutralizing positive charges and thus decreasing the affinity of histones to DNA17. This facilitates the accessibility of transcriptional regulators and other chromatin modifiers18. In contrast, histone deacetylases (HDACs) antagonize the action of HATs by removing the acetyl groups from histone19.Chromatin conformation influences the accessibility of transcriptional regulator(s) to the associated DNA regions. Chemical modifications at histone N-terminal tails contribute to proper packing of chromatinArabidopsis circadian clock identified the circadian changes in Histone 3 acetylation (H3ac) at the TOC1 promoter. The antagonistic action of CCA1 at dawn and another single MYB clock-related transcription factor REVEILLE8/LHY-CCA1-LIKE5 (RVE8/LCL5) was found to define the hypo-acetylated and hyper-acetylated states of H3 at the TOC1 promoter during the day10. Indeed, CCA1 facilitates a repressive chromatin conformation at dawn either by interfering with HAT accessibility or by recruiting HDAC activity20. During the day, RVE8/LCL5 antagonizes the CCA1 repressive function, favoring H3ac10. These counteracting functions precisely shape the waveform of TOC1 expression10. The rhythmic accumulation of different histone marks was also observed at the promoters of other core clock components, including CCA1, LHY, LUX ARRHYTHMO (LUX), and PRRs, and correlates with their transcript accumulation21.The first study relating chromatin modification and the TOC1 locus, the chromatin remodeling factor(s) contributing to the rhythms of histone acetylation and its inner working mechanism remain elusive. Here, we report that HDA9, a member of the reduced potassium dependency 3 (RPD3)/HDA1 family class I HDAC23, is involved in the circadian regulation of TOC1 by suppressing its expression. HDA9 specifically interacts with an EC component, ELF3, and the physical interaction enables HDA9 to bind to the TOC1 promoter. HDA9 might facilitate a closed chromatin structure at the TOC1 promoter, contributing to its declining phase of expression during the night period. These results provide an insight into how temporal regulation of histone acetylation is achieved in order to stably maintain circadian activity.Despite the clear rhythms in histone acetylation at the 24. As pharmacological inhibition of the RPD3/HDA1 family class I HDAC activities25 with TSA (Trichostatin A) affects the circadian oscillation25, we investigated their possible function within the circadian clock. Among the four members of this family 26, we focused on two members not previously studied, HDA7 and HDA9.Previous studies have shown that histone deacetylation is important for the rhythmic oscillation in histone acetylation at the core clock promotersHDA7 and HDA9-deficient mutants, hda7–2 and hda9–1, and analyzed rhythmic expression of the circadian oscillator gene, CCA1. Quantitative real-time RT-PCR (RT-qPCR) analysis revealed that the circadian expression of CCA1 was unaffected in hda7–2 mutant compared with wild type  and CHLOROPHYLL A/B BINDING–PROTEIN 2 (CAB2), and several circadian oscillator genes displayed a similar pattern with a period shortening and advanced rhythmic phase  in the promoter regions of selected clock genes, which include key clock-related cis-elements, such as CCA1-binding site (CBS), evening element (EE), G-box, and/or LUX binding site (LBS)  assays using 35 S:BS) Fig. 5,7,27,28ter Fig. . HDA9 spcus Fig. , but notHDA9 did not significantly oscillate throughout a circadian cycle   Fig. , when TO23. Since HDA9 binds to the TOC1 promoter  might contribute to the rhythmic function of HDA9. As HDA9 itself does not have binding selectivity on specific DNA regions, DNA-binding proteins may be required for specific HDA9 binding to the TOC1 promoter. To examine this hypothesis, we carried out yeast-two-hybrid assays with expression constructs containing core clock transcription factors. The HDA9-GAL4 DNA binding domain fusion construct was co-expressed with a construct expressing a clock gene fused in-frame to the 3’-end of GAL4 activation domain in yeast cells. Cell growth on the selective medium revealed that HDA9 specifically interacted with ELF3 , and the ELF3 gene was fused in-frame to the 5’-end of a sequence encoding the N-terminal half of YFP (nYFP). The fusion constructs were then transiently co-expressed in Arabidopsis protoplasts. The HDA9-ELF3 combination was able to visualize yellow fluorescence, which was exclusively detected in the nucleus  analysis using eus Fig. . Moreoveeus Fig. . These r30, HDA9 could be associated with the EC. This is in agreement with the temporal association of HDA9 to TOC1, because EC function is highly relevant at ZT1630. However, it is currently unknown whether the EC indeed associates with the TOC1 promoter. To test this possibility, we employed pELF3:ELF3-MYC/elf3–1, pELF4:ELF4-HA/elf4–2, and pLUX:LUX-GFP/lux-4 transgenic plants30. ChIP assays were performed with epitope-tagged transgenic plants immunoprecipitated with corresponding antibodies. DNA bound to epitope-tagged proteins was analyzed by qPCR assays. The ChIP-qPCR analysis showed that the TOC1 promoter was enriched by ELF3, ELF4, and LUX   Fig.  and 12. TOC1 promoter was tested using 35S:MYC-HDA9 transgenic plants crossed with the elf3–8 mutant. ChIP assays using an anti-MYC antibody showed that HDA9 binding to the TOC1 locus was reduced in the elf3–8 mutant background compared with the wild-type background /ARABIDOPSIS TRITHORAX–RELATED 3 (ATXR3) protein establishes H3K4me3 mark at core clock gene loci to promote expression. The H3K4me3 mark inhibits clock repressor binding at the core clock promoters, facilitating transcriptional repression to target genes only at specific time-of-dayArabidopsis circadian system. PRR5, PRR7, and PRR9 directly repress CCA1 and LHY34. Notably, PRRs interact with TOPLESS/TOPLESS RELATED PROTEINS (TPL/TPRs), as well as HDA6 and HDA1934. Consistently, inhibition of the TPL activity compromises transcriptional repression activities of PRR5, PRR7, and PRR9, lengthening circadian period34. The potent HDAC inhibitor TSA activates CCA1 expression, but this activation is impaired in the tpl-1 mutant, demonstrating that HDAC regulation is largely dependent on TPL34.A couple of HDACs have been shown to be implicated in the TOC1 expression. HDA9 binds to the TOC1 promoter and stimulates H3 deacetylation to suppress TOC1 expression. HDA9 interacts with the EC component ELF3, and rhythmic binding of HDA9 to the TOC1 promoter is defined by the EC. Accordingly, although HDA9 has no obvious rhythmic expression pattern, the EC-HDA9 interactions facilitate temporal regulation of TOC1. Consistent with the fact that the EC is expressed highly during night period35, the EC-HDA9 complex is relevant in the declining phase of TOC1 expression. Nonetheless, many questions remain to be answered. Considering the variable alteration patterns of circadian gene expression in hda9–1 mutant, EC and HDA9 are extensively interconnected with other circadian and chromatin-related components, accounting for dynamic contribution to circadian oscillation. Moreover, it is sometimes observed the anti-phase of binding regions and histone modifications36. While HDA9 binds to a proximal region of the TOC1 promoter, changes in histone acetylation are drastically observed in a distal region. This may be due to propagation and expansion of epigenetic modification of chromatin. Chromatin remodeling components that regulate epigenetic contexts on chromatin would also be involved, and thus the responsible protein components should be further elucidated. Future studies will give a comprehensive view of circadian-chromatin networks. Altogether, these results exemplify a mechanism by which chromatin factors engage with clock components to rhythmically modulate chromatin conformation and hence transcriptional oscillation.We here show that HDA9 regulates circadian oscillation by modulating HDA9 is highly expressed in dry seeds and represses seedling traits to maintain seed dormancy37. The hda9 mutants show reduced seed dormancy with higher expression of genes responsible for photosynthesis and photoautotroph, such as RuBisCO and RuBisCO activase (RCA)37. The HDA9 action is antagonistic to HDA6 and HDA19, which repress embryonic properties in germinating seeds37, balancing the proper transition from seed to autotropic seedling.The HDA9 protein is implicated in plant developmental processes. HDA9-deficient mutants also exhibit early flowering phenotypes under short day conditions39. HDA9 binds to the AGAMOUS–LIKE 19 (AGL19) promoter and negatively regulates its transcription through the alterations in H3K9 and H3K27 acetylation levels at the AGL19 locus39. Consequently, the HDA9-AGL19 module mainly regulates FLOWERING LOCUS T (FT) expression, in parallel with photoperiod or autonomous pathways to prevent precocious flowering in short days39. The epigenetic regulation of AGL19 by HDA9 is also relevant, in part, during vernalization process. Given that HDA9 is functional when AGL19 is actively expressed, such in the adult stage or after vernalization39, this HDAC protein is able to associate with transcriptionally active genes to reset the chromatin state. Accumulating evidence further supports that the HDA9-POWERDRESS (PWR) complex stimulates H3 deacetylation at a genome-wide level and is implicated in additional developmental processes, such as floral stem cell fate, stress responses, and aging23.The 40. Homologs of each component are present in Arabidopsis: six SIN3-Like (SNL1-SNL6), four RPD3 HDACs, one SAP18, and two SAP30 function-relateds (AFR1 and AFR2)40. The Arabidopsis Sin3-HDAC complex is crucial for diurnal control of FT40. This complex accumulates at dusk and is recruited to FT promoter to ensure periodic histone deacetylation. Dynamic cycles of H3 acetylation and deacetylation ensure adequate levels of the gene transcription and thereby prevent precocious flowering in the premature inductive conditions40. This study further supports periodic functions of HDA9 in controlling circadian gene expression. H3ac states can be reversible and account for stably oscillating gene expression. Taken together, the HDA9 protein seems to integrate temporal information to regulate oscillator gene expression and a variety of physiological output processes in order to optimize plant growth and development.Notably, HDA9 activity can be diurnally gated. HDACs often form a massive protein complex containing for instance Sin3, RPD3-type HDAC, SIN3-ASSOCIATED POLYPEPTIDE 18 (SAP18) and SAP30Arabidopsis thaliana (Columbia-0 ecotype) was used for all experiments unless otherwise specified. Plants were grown under neutral-day conditions  with cool white fluorescent light (100 μmol photons m−2 s−1) at 22 °C. The elf3–8, elf4–101, hda7–2, and hda9–1 were previously reported43. The lux-6 mutant (SALK-022315) was obtained from Arabidopsis Biological Resource Center (ABRC). To produce transgenic plants overexpressing the HDA9 gene, a full-length cDNA was subcloned into the binary pBA002 vector under the control of the CaMV 35 S promoter. Agrobacterium tumefaciens-mediated Arabidopsis transformation was then performed.Plants grown under conditions/ZT indicated were harvested and ground in liquid nitrogen. Total RNA was extracted by mixing the tissue powder with 700 μl of TRI reagent . The extraction mixture was incubated at 65 °C for 5 min. After mixing with 700 μl of chloroform:isoamyl alcohol (24:1), the extraction mixture was centrifuged for 20 min at room temperature. The RNA in the liquid phase was precipitated at 4 °C in a final concentration of 2 M LiCl. RNA pellet was washed by 70% ethanol, briefly air dried, and suspended in distilled water. Reverse transcription (RT) was performed using Moloney Murine Leukemia Virus (M-MLV) reverse transcriptase  with oligo(dT18) to synthesize first-strand cDNA from 2 μg of total RNA. Total RNA samples were pretreated with an RNAse-free DNAse to remove DNA contamination.T method was used to determine the relative gene expression, with the expression of Eukaryotic translation initiation factor 4A1 (eIF4A) gene (At3g13920) as the internal control. All RT-qPCR reactions were performed with biological triplicates using total RNA samples extracted from three independent replicate samples.Quantitative RT-PCR reactions were performed on the Step-One Plus Real-Time PCR System (Applied Biosystems). The PCR primers used are listed in Supplementary Table LacZ and His reporter genes was used. PCR products were subcloned into the pGBKT7 and pGADT7 vectors. The expression constructs were co-transformed into yeast AH109 cells, and transformed cells were selected by growth on SD/-Leu/-Trp medium and SD/-Leu/-Trp/-His/-Ade.Yeast two-hybrid assays were performed using the BD Matchmaker system . The pGADT7 vector was used for the GAL4 activation domain fusion, and the pGBKT7 vector was used for the GAL4 DNA binding domain fusion. The yeast strain AH109 harboring the HDA9 gene was fused in-frame to the 5′ end of a gene sequence encoding the C-terminal half of EYFP in the pSATN-cEYFP-C1 vector (E3082). The ELF3 cDNA sequences were fused in-frame to the 5′ end of a gene sequence encoding the N-terminal half of EYFP in the pSATN-nEYFP-C1 vector (E3081). Expression constructs were co-transformed into Arabidopsis protoplasts. Expression of the fusion constructs was monitored by fluorescence microscopy using a Zeiss LSM510 confocal microscope .The 3) followed by the digestion with 2.5 ml Proteinase-K  for overnight at 65 °C. DNA was purified using phenol/chloroform/isoamyl alcohol and sodium acetate (pH 5.2). The level of precipitated DNA fragments was quantified by quantitative real-time PCR using specific primer sets (Supplementary Table eIF4a). Values for control plants were set to 1 after normalization against eIF4a for quantitative PCR analysis.The epitope-tagged transgenic plant samples were cross-linked with 1% formaldehyde, ground to powder in liquid nitrogen, and then sonicated . The sonicated chromatin complexes were bound with corresponding antibodies. Anti-MYC , anti-H3ac , anti-HA , and salmon sperm DNA/protein A agarose beads  were used for chromatin immunoprecipitation. The immunoprecipitated chromatin complexes were purified by incubating with 50 μl slurry of Protein-A Sepharose  pre-equilibrated with 1 mg/ml salmon sperm DNA and 1 mg/ml BSA for 3 h at 4 °C. The affinity beads bound chromatin complexes were washed with 850 μl nuclei lysis buffer for three times, LNDET buffer  for three times and TE buffer  for three times. Chromatin complexes were eluted from the beads with use of 200 μl elution buffer (1% SDS, 0.1 M NaHCOArabidopsis protoplasts, reporter and effector plasmids were constructed. The reporter plasmid contains a minimal 35S promoter sequence and the GUS-coding gene. The TOC1 promoter was inserted into the reporter plasmid. The coding regions of HDA9, ELF3, ELF4, and LUX were subcloned into the effector vector for transient overexpression under the control of the CaMV 35 S promoter. An amount of 2.5 × 105 protoplasts was co-transfected with 4 μg reporter construct, 4 μg effector construct, and 2 μg CaMV 35 S promoter-luciferase construct as the internal transfection control. Transfected cells were incubated at 22 °C overnight and harvested for LUC and GUS reporter assays.For transient expression assays using Further information on experimental design is available in the Supplementary InformationReporting Summary


Transcriptional regulation is a critical biological process that allows the cell or an organism to respond to a variety of intra- and extra-cellular signals, to define cell identity during development, to maintain it throughout its lifetime, and to coordinate cellular activity. This highly dynamic mechanism includes a series of biophysical events orchestrated by a huge number of molecules establishing larger networks and occurring through multiple temporal and functional steps that range from specific DNA-protein interactions to the recruitment and assembly of nucleoprotein complexes. Essentially, the key transcription levels include the recruitment and assembly of the entire transcription machinery, the initiation step, pause release and elongation phases, as well as termination of transcription. Additionally, these steps are interconnected with governing chromatin accessibility , and other epigenetic mechanisms . Finally, various RNA maturation events, such as the splicing that occurs with transcription, constitute an additional level of complexity. Numerous molecules and molecular factors, including transcription factors, cofactors (both coactivators and corepressors), and chromatin regulators, are known to participate to this process . EssentiAdditionally, the high complexity of transcriptional regulation is also derived from the involvement of non-coding RNAs (ncRNAs). Indeed, research over the last two decades has revealed new classes of ncRNAs, including microRNAs (miRNAs), small nucleolar RNAs (snoRNAs), long ncRNAs (lncRNAs), circular RNAs (circRNAs), and enhancer RNAs (eRNAs), each with different regulatory functions and altogether belonging to a larger RNA communication network ultimately controlling the production of the final protein .Recent advances in “omics” and computational biology have provided novel tools that allow one to integrate different layers of information from biophysical, biochemical, and molecular cell biology studies. In turn, these novel strategies provided a fuller understanding of how DNA sequence information, epigenetic modifications, and transcription machinery cooperate to regulate gene expression. Of note, most of the new molecular biomarkers and therapeutic targets for several human pathologies derive from transcriptome profiling studies, and their number is continuously increasing. Next Generation Sequencing (NGS), mainly RNA-Sequencing (RNA-Seq), has completely revolutionized transcriptome analysis, allowing the quantification of gene expression levels and allele-specific expression in a single experiment, as well as the identification of novel genes, splice isoforms, fusion transcripts, and the entire world of ncRNAs at an unprecedented level .It is well known that many human disorders are characterized by global transcriptional dysregulation because most of the signaling pathways ultimately target transcription machinery. Indeed, many syndromes and genetic and complex diseases—cancer, autoimmunity, neurological and developmental disorders, metabolic and cardiovascular diseases—can be caused by mutations/alterations in regulatory sequences, transcription factors, cofactors, chromatin regulators, ncRNAs, and other components of transcription apparatus ,2,3,4. TIn this special issue, a total of 19 excellent and interesting papers consisting of 11 original research studies, seven reviews, and one communication are published ,21,22,23CLIC4). This is a multifunctional protein with diverse physiological functions. Differential expression of CLIC4 between cancer cells and the surrounding stroma has been reported in various tumor types [CLIC4 transcription levels [Huang et al. analyzed the transcriptional regulation of the gene coding for the Chloride intracellular channel 4  transcription factor in the winged pearl oyster Pteria penguin. More precisely, this study investigated the role of PpPax3 in melanin synthesis and used RNA interference to provide evidence that this function is exerted in this important marine species through the tyrosinase pathway [FOXO1 (Forkhead Box O1) as a strong transcription factor interacting with other key genes associated with lethal heart disorders in PS. [Regarding transcription factors, a comprehensive review summarized the structures and functions of these regulators in both model and non-model insects, including Drosophila, and appraises the importance of transcription factors in orchestrating diverse insect physiological and biochemical processes . An orig pathway . A bioins in PS. .As expected in the NGS era, transcriptome analysis by RNA-Seq has been widely used in many studies to elucidate the most varied mechanisms of pathophysiology as well as other relevant biological processes in diverse organisms ,9,20,21.In the field of cancer research, an interesting pathogenetic mechanism involving dysregulation of transcription is represented by the destabilization of the messenger RNAs of critical genes implicated in both tumor onset and tumor progression exerted by tristetraprolin (TTP). Indeed, as reviewed in a paper of this special issue, the tumor suppressor TTP can negatively regulate tumorigenesis. In turn, TTP expression is frequently downregulated in several tumors by various mechanisms .Akt1), a gene related to metabolism [Several papers have described novelties in the field of ncRNAs. For instance, a study investigated the possible role in cell metabolism of miR-25-3p. This miRNA is highly conserved in mammals and was previously found to be involved in many biological processes and in some cancer and cardiovascular related diseases. Specifically, in the C2C12 cell line derived from mouse muscle myoblasts, it is positively regulated by the transcription factor AP-2α and enhances cell metabolism by directly targeting the 3′ untranslated region of AKT serine/threonine kinase 1  gene summarizes all the possible (epi)-genetic alterations leading to diseases. This gene encodes the p57Kip2 protein, the third member of the CIP/Kip family, and its alterations are known to cause three human hereditary syndromes characterized by altered growth rate. Interestingly, CDKN1C is positioned in a genomic region characterized by a remarkable regional imprinting that results in the transcription of only the maternal allele. Moreover, this gene is also down-regulated in human cancers. Of note, its transcriptional regulation is linked to several mechanisms, including DNA methylation and specific histone modifications. Finally, ncRNAs also play important roles in controlling p57Kip2 levels [Numerous epigenetic mechanisms other than regulation by ncRNAs take place during RNA polymerase II-transcription and may be involved in human pathophysiology. An outstanding review on the Cyclin Dependent Kinase Inhibitor 1C (Selenium-related transcriptional regulation is the topic of a comprehensive review . SeleniuIt is well-known that DNA methylation contributes to the gene expression regulation without changing the DNA sequence. Abnormal DNA methylation has been associated with improper gene expression and may lead to several disorders. Both genetic factors and modifiable factors, including nutrition, are able to alter methylation pathways. An interesting review of this special issue carefully describes molecular mechanisms underlying the link between diet and DNA methylation .Finally, we hope the readers enjoy this Special Issue of IJMS and the effort to present the current advances and promising results in the field of transcriptional regulation and its involvement in all of the relevant biological processes and in pathophysiology.


The nucleolus as site of ribosome biogenesis holds a pivotal role in cell metabolism. It is composed of ribosomal DNA (rDNA), which is present as tandem arrays located in nucleolus organizer regions (NORs). In interphase cells, rDNA can be found inside and adjacent to nucleoli and the location is indicative for transcriptional activity of ribosomal genes—inactive rDNA (outside) versus active one (inside). Moreover, the nucleolus itself acts as a spatial organizer of non-nucleolar chromatin. Microscopy-based approaches offer the possibility to explore the spatially distinct localization of the different DNA populations in relation to the nucleolar structure. Recent technical developments in microscopy and preparatory methods may further our understanding of the functional architecture of nucleoli. This review will attempt to summarize the current understanding of mammalian nucleolar chromatin organization as seen from a microscopist’s perspective. The nucleolus is the site of the highest RNA synthesis rate in cell nuclei giving rise to more than 60% of the entire RNA pool. Compared to highly transcribed nucleoplasmic gene foci, the nucleolus can be considered as particularly active transcription hub or transcription factory. While there are common principles as to the spatial organization of nucleoplasmic and nucleolar transcription  are surrounded by electron dense fibrillar stretches, dense fibrillar component (DFC), which may extend away from FCs. These fibrillar structures lie embedded in a large body of granular nature, granular component (GC), which dictates the overall shape of the nucleolus. In addition to these three components, patches of heterochromatin reside at the nucleolar periphery interspersed by euchromatin Fig. a, b. FroIn the past, intense research efforts of several workgroups tried to correlate the structure of rDNA with the nucleolar components. Consensus has been reached in the finding that rDNA is located within the fibrillar parts of nucleoli, in the perinucleolar heterochromatin, and in intranucleolar stretches. In addition, ribosomal RNA was unequivocally localized to DFC and GC, and nascent transcripts identified by incorporated labeled precursor nucleotides were identified in the fibrillar parts of nucleoli. Together, these results demonstrated that, outside the nucleoli, there reside transcriptionally inactive rDNA repeats, whereas rRNA transcription occurs in the fibrillar areas. It should be noted that inactive rRNA genes might also be found in the intranucleolar chromatin as part of a loop of rDNA emanating out of perinucleolar  rDNA into the nucleolar interior. rRNA processing, which starts co-transcriptionally, is found in DFC and GC, in the latter of which assembly of rRNA with ribosomal proteins takes place. Unfortunately, although similar technical approaches were used, no consensus was reached on the exact localization of rDNA within the two fibrillar areas and as to where the site of transcription precisely is located  revealed, however, that maintenance of the large-scale nucleolar structure is based on surprisingly dynamic behavior of its constituents. It was found that Pol I components are continuously exchanged at sites of rRNA transcription and thus only transiently localize to fibrillar components of nucleoli  made it difficult to evaluate the arrangement of entire rDNA repeats or even array segments thereof within the nucleolus. It should be kept in mind that an elongated Christmas tree is larger than the diameter of the average FC/DFC complex (4.4 µm versus approx. 1 µm). Clearly, rDNA fibers have to be present within nucleoli in a conformation providing a significant packaging while concomitantly ensuring massive transcriptional activity.Combinatorial studies using histochemistry and immunogold methods, volumetric imaging, or ChIP, 3C data sets, and modeling led to the proposal of several models of the 3-D arrangement of rDNA within nucleoli. The DNA-cloud model proposes an inner core of DNA fibers located within FCs originating from condensed intranucleolar chromatin and further expanding in a radial fashion towards the periphery. These peripheral fine chromatin fibers overlap with DFCs where the transcriptionally active genes are located  is long known to be an essential factor for rRNA transcription and has recently been identified as factor responsible for defining the chromatin state of actively transcribed mammalian rRNA genes , which led to the conclusion that significant amounts of histones are present on active rRNA genes not impeding Pol I transcription in yeast  of the JmjC family were found to fulfill dual roles in rDNA transcription control. KDM2A and KDM2B induce silencing, whereas KDM4B removes silencing marks. KDM2A de-methylates H3K36 at the rDNA promoter  are generally associated with transcriptional silencing. HDAC1 and HDAC2 interact with NoRC and facilitate the silencing of rDNA  seem to have inverse roles on rRNA transcription. The former acts as a negative regulator, whereas the latter promotes transcription in mammalian cells .Promoter elements structurally similar to rRNA gene promoters and transcribed by Pol I were identified in the IGS previously. One of these transcripts derived from upstream of the rDNA promoter is involved in rDNA silencing by binding to NoRC and targeting the complex to the promoter  preferentially associate with nucleoli. Interestingly, although located predominately in the perinucleolar heterochromatin, these sequences display genomic architecture similar to euchromatin and are transcribed , nucleoplasmin-like protein led to centromere re-location from nucleoli and was thus suggested to be implicated in nucleolar tethering in concert with CTCF  domains;  and their adjacent chromatin. A complex meshwork made of intermediate filaments, the nuclear lamina, mediates between the chromatin and the nuclear membrane, and consists of a well-defined set of interacting factors that are enriched at the nuclear periphery. In vertebrates, the nuclear periphery seems to be primarily repressive for transcription with the exception of the active chromatin surrounding the NPs  and in preparation improvements [expansion microscopy; (Chen et al. Another very interesting focus is the heterogeneity of the rDNA array, both in terms of genetic variability and in epiallelic composition. Such studies have already revived the interest of pathologists in rDNA and nucleoli and will certainly continue to do so, particularly in oncology. Nucleoli recently became therapeutic targets for small anti-cancer drugs specifically inhibiting the Pol I machinery (Drygin et al. Recent progress made in deciphering the spatio-temporal functional architecture of the nucleolus will enhance our understanding of fundamental cellular processes such as metabolic adaptation to environmental conditions, senescence, and aging leading to the improved diagnosis and treatment of diseases.


Neurospora crassa has just two H3K36 KMTs, ASH1 and SET-2, and were able to explore the function and distribution of each enzyme independently. While H3K36me deposited by SET-2 marks active genes, inactive genes are modified by ASH1 and its activity is critical for their repression. ASH1-marked chromatin can be further modified by methylation of H3K27, and ASH1 catalytic activity modulates the accumulation of H3K27me2/3 both positively and negatively. These findings provide new insight into ASH1 function, H3K27me2/3 establishment, and repression in facultative heterochromatin.Methylation of histone H3 at lysine 36 (H3K36me), a widely-distributed chromatin mark, largely results from association of the lysine methyltransferase (KMT) SET-2 with RNA polymerase II (RNAPII), but most eukaryotes also have additional H3K36me KMTs that act independently of RNAPII. These include the orthologs of ASH1, which are conserved in animals, plants, and fungi but whose function and control are poorly understood. We found that  Not all genes in a cell’s DNA are active all the time. There are several ways to control this activity. One is by altering how the DNA is packaged into cells. DNA strands are wrapped around proteins called histones to form nucleosomes. Nucleosomes can then be packed together tightly, to restrict access to the DNA at genes that are not active, or loosely to allow access to the DNA of active genes.Chemical marks, such as methyl groups, can be attached to particular sites on histones to influence how they pack together. One important site for such marks is known as position 36 on histone H3, or H3K36 for short. Correctly adding methyl groups to this site is critical for normal development, and when this process goes wrong it can lead to diseases like cancer. An enzyme called SET-2 oversees the methylation of H3K36 in fungi, plants and animals. However, many species have several other enzymes that can also add methyl groups to H3K36, and their roles are less clear.Neurospora crassa contains just two enzymes that can add methyl groups to H3K36: SET-2, and another enzyme called ASH1. By performing experiments that inactivated SET-2 and ASH1 in this fungus, Bicocca et al. found that each enzyme works on a different set of genes. Genes in regions marked by SET-2 were accessible for the cell to use, while genes marked by ASH1 were inaccessible. ASH1 also affects whether a methyl group is added to another site on histone H3. This mark is important for controlling the activity of genes that are critical for development.A type of fungus called ASH1 is found in many other organisms, including humans. The results presented by Bicocca et al. could therefore be built upon to understand the more complicated systems for regulating H3K36 methylation in other species. From there, we can investigate how to intervene when things go wrong during developmental disorders and cancer. Methylation of histone H3 at lysine 36 (H3K36me) is largely associated with euchromatic regions of eukaryotic genomes . It servS. cerevisiae and S. pombe, where H3K36me is nonessential and performed by a single RNAPII-associated KMT  , and theated KMT . The simated KMT .Neurospora crassa as an experimental bridge between yeasts and higher organisms, and use it to address unresolved questions concerning H3K36me. As in S. cerevisiae and S. pombe, H3K36me is not essential in N. crassa but unlike the case in the yeasts, we found that H3K36 methylation results from a division of labor between the RNAPII-associated SET-2 enzyme which can catalyze mono-, di-, and tri-methylation  mutation displayed severely compromised growth but only minor reductions in global H3K36me2 and –me3  double mutants showed additive loss of the H3K36me2 observed in the single mutants and loss of the residual H3K36me3 signal  strains, respectively. Overall, we found that H3K36me2 and –me3 is associated with gene-rich DNA and excluded from constitutive heterochromatin, which is marked by DNA methylation and H3K9me3 (−4)  . In the H3K36me2 , and theern blot , H3K36meH3K36me2 .ash1 and set-2 mutants have substantial, but distinct, changes in gene expression relative to WT. The set-2 deletion showed a relatively symmetrical distribution of gene expression changes with 916 genes up-regulated and 1222 genes down-regulated  background, while ASH1-unmarked genes showed no pattern of altered regulation  . Though K27me2/3 , we alsoK27me2/3 . Next, wK27me2/3 . Using aH3K36me2 . AltogetDrosophila Ash1 has previously been reported to inhibit PRC2-mediated repression by preventing H3K27me2/3 accumulation . Similarash1(Y888F) background. Inactivation of ASH1 showed a striking effect on H3K27me2/3, resulting in reduction or complete loss of the mark across roughly one-third of ASH1/PRC2-comarked genes  (ash1(Y888F) strain are delineated by regions normally marked with ASH1-catalyzed H3K36me2  , whereaskground) . ImportaH3K36me2 . Thus, AN. crassa, which we showed consists of two H3K36 KMTs, SET-2 and ASH1. Unlike SET-2 , which can catalyze mono-, di-, and tri-methylation, and a group of specialized KMTs that largely catalyze mono/di-methylation . Study oke SET-2 , ASH1 apFusarium fugikuroi and Plasmodium falciparum have also been reported to have H3K36me3 activity  is essential, and other units of the complex – EAF-3, SIN3, and NPF – are components of various other chromatin modifying complexes , which includes the H3K36me3 reader, EAF-3 . RPD3S aomplexes . Notablyomplexes , furtherN. crassa has no apparent PRC1 components. Even in higher organisms, the mechanism of repression mediated by PRC2 and H3K27me2/3 is far from clear, necessitating additional studies. Interestingly, we observed in Neurospora that loss of ASH1-dependent H3K36 methylation was associated with both losses and gains of H3K27me2/3. Early work with Drosophila gave evidence that ASH1 opposes the action of PRC2 function , NCU07152 , hH4 .All Neurospora strains used in this study are listed in Key resources table. Strains were grown, crossed, and maintained according to standard procedures . KnockouImmunoblotting was performed as previously described . BrieflyN. crassa OR74A (NC12 genome) (ChIP was performed as previously described . qPCR wa genome)  using Bo genome) . ChIP-se genome)  and SAMt genome)  on the o genome) . Sequenc genome) . The folRNA was isolated , DNase t2ratios from two BAM files; 3) computeMatrix was used to prepare data for heatmaps or profiles; 4) plotHeatmap was used to create heatmaps for score distributions; 5) plotProfile was used to create meta-plots of score distributions. Tools available from SAMtools  and are accessible through GEO Series accession number GSE118495 and, as part of a previously reported series GSE82222 , consistent with the 'conventional' view that H3K36me2/3 directly antagonizes H3K27 methylation by PRC2. The authors conclude that ASH1 catalyzed H3K36me2 can affect H3K27me2/3 accumulation both in a positive and in a negative manner.The study by Bicocca et al. investigates the role of H3K36 methylation and its interplay with H3K27 methylation in Drosophila , in worms  and in experiments that used purified PRC2 for HMTase reactions on nucleosomes in vitro . On the other hand, the authors also present data that stand in direct contrast to this regulatory relationship. Specifically, the ChIP analyses suggest that ASH1-mediated H3K36me2 deposition can – somehow – act in a positive way to promote H3K27 methylation by PRC2. The authors discuss these findings that are not so simple to reconcile in a balanced way. Perhaps, their discussion could also touch on a two main issues mentioned in point 4 below.This study is interesting. At some genomic regions, H3K36me2/3 deposited by ASH1 antagonizes H3K27 methylation by PRC2, consistent with previous findings made in Essential revisions:set-2 ash1 double mutant) in Figure 1F it indeed appears that there is a further reduction of H3K36me3 signal compared to lane 3 (set-2 single mutant). The authors should clarify this point because, as the authors point out in the Introduction, there is no evidence that HMTases of the ASH1/NSD class are at all capable to tri-methylate H3K36. Since the effects are subtle, it would have been useful to show a more quantitative analysis of bulk modification levels, by performing western blots on serial dilutions (e.g. 4:2:1) of the extracts, rather than just one amount of extract per genotype. If the authors have more quantitative WB data, we encourage them to add them to the manuscript.1) Looking at WB in Figure 1E, it seems that there is still a very faint H3K36me3 WB signal in lane 7 and that this signal is comparable to that seen in lanes 3 and 4 of the same blot. So, from this, it does not look like ASH1 would contribute to generating H3K36me3, whereas in lane 4  Figure 2. First, how were the ChIP-seq reads from wt, 3) For the RNA-seq analysis, it would be good to have more quantitative information on the extent of up- and down-regulation. In particular, how many genes are more than 2-fold up- or down-regulated and how many genes are more than 4-fold up- or down-regulated?4) These are two points that the authors may want to discuss.Neurospora? Does Neurospora contain a chromodomain-containing protein in a PRC1-type complex ?First, what is the function of H3K27me2/3 in Neurospora PRC2 is at all inhibited on H3K36me2/3 nucleosomes in vitro.The second point that might be interesting to discuss is that unlike in flies and mammals, in plants, not all forms of PRC2 are inhibited by active marks . Specifically, PRC2 complexes containing the Su(z)12 ortholog EMF2 are inhibited by H3K4me3 but PRC2 containing the Su(z)12 ortholog VRN2 are not inhibited by this mark . So, it remains to be tested whether the HMTase activity of  Essential revisions:1) Looking at WB in Figure 1E, it seems that there is still a very faint H3K36me3 WB signal in lane 7 and that this signal is comparable to that seen in lanes 3 and 4 of the same blot. So, from this, it does not look like ASH1 would contribute to generating H3K36me3, whereas in lane 4 (set-2 ash1 double mutant) in Figure 1F it indeed appears that there is a further reduction of H3K36me3 signal compared to lane 3 (set-2 single mutant). The authors should clarify this point because, as the authors point out in the Introduction, there is no evidence that HMTases of the ASH1/NSD class are at all capable to tri-methylate H3K36. Since the effects are subtle, it would have been useful to show a more quantitative analysis of bulk modification levels, by performing western blots on serial dilutions (e.g. 4:2:1) of the extracts, rather than just one amount of extract per genotype. If the authors have more quantitative WB data, we encourage them to add them to the manuscript.set-2 mutant and the ash1/set-2 double mutant. The contribution of ASH1 is more obvious in Figure 1F (perhaps helped by the lanes being directly adjacent) than in Figure 1E, but we believe the conclusions drawn from both blots are consistent. In each case, the signal remaining after set-2 deletion is very faint, suggesting SET-2 is responsible for nearly all of H3K36me3. Though faint, the intensity of this remaining signal is reduced when combined with the ash1 mutant. The phenotype is robust in its reproducibility, but there is slight variation in the apparent contribution of ASH1. We felt these blots were representative of the phenotype, but took the recommendation of performing serial dilution blots to further demonstrate the phenotype. The new data, now shown as Figure 1F, reinforce our conclusion.We share the concern regarding the best way to demonstrate the subtle change between the Fusarium fujikuroi and Plasmodium falciparum. In addition, C. elegans mes-4 (an ortholog of NSD1) has been reported to have H3K36me3 activity .Regarding evidence of H3K36me3 activity from ASH1/NSD orthologs, we note in the Introduction that no in vitro analysis of ASH1 orthologs has demonstrated H3K36me3 activity. However, as we note in the Discussion section, there is in vivo evidence for it. Janebska et al. (2017) and Jiang et al. (2013) describe H3K36me3 activity from ASH1-orthologs in 2) Figure 2. First, how were the ChIP-seq reads from wt, ash1 and set-2 mutants normalized to take into account the changes in bulk H3K36me2 and -me3 levels? Second, have the authors generated H3K36me2 and -me3 profiles in set-2 ash1 double mutant cells?ChIP-seq tracks are shown with as little manipulation as possible; they are simply normalized to reads per kilobase per million (RPKM) and averaged over 25bp bins. We do not think the ChIP-seq tracks should be used to interpret the relative intensity of H3K36me catalyzed by SET-2 versus ASH1 and tried not to make claims to suggest they could. Largely, we think each mutant should be considered independently to draw more qualitative conclusions about where the mark is being found. However, the data do provide an opportunity to compare the relative contribution of SET-2 and ASH1 to the profile observed in WT. Consider the 50kb region we highlight in Figure 2A and 2C, where we see that SET-2 contributes little or no H3K36me2 signal, but in the WT track we still observe a H3K36me2 signal that is comparable in intensity to adjacent regions. This suggests ASH1 is both responsible for nearly all the H3K36me2 signal found at this location and the intensity of the ASH1 signal is comparable to that catalyzed by SET-2 in the neighboring region.set-2/ash1 double mutant. As one might expect, ChIP of H3K36me from the set-2/ash1 double mutant yielded very little DNA  and the ChIP-seq tracks we interpreted as non-specific. The set-2/ ash1 double mutant was more valuable as a control in ChIP qRT-PRC experiments, which avoid some of the normalization concerns of ChIP-seq experiments. We have therefore added qRT-PCR results to Figure 2—figure supplement 1 to better demonstrate the relative signal contribution from ASH1 and SET-2, and to show that the signal attributed to either ASH1 and SET-2 is eliminated in the double mutant.We were able to build a set-2, and Δset-2; ASH1(Y888F) strains at three genomic regions with distinct H3K36me profiles. 8:G3 is a constitutive heterochromatin region that lacks H3K36me and is used to assess “background” in the ChIP. hH4 is an actively expressed gene that is marked by SET-2-catalyzed H3K36me but not ASH1-catalyzed H3K36me. NCU07152 in a silent uncharacterized gene that is densely marked by ASH1-catalyzed H3K36me and H3K27me2/3.H3K36me3 ChIP results are shown for WT, Δ3) For the RNA-seq analysis, it would be good to have more quantitative information on the extent of up- and down-regulation. In particular, how many genes are more than 2-fold up- or down-regulated and how many genes are more than 4-fold up- or down-regulated?set-7 data were taken from Klocko et al. (2016).We agree and have included a more quantitative analysis of our RNA-seq data in a table included as Figure 5—figure supplement 2. Δ4) These are two points that the authors may want to discuss.First, what is the function of H3K27me2/3 in Neurospora? Does Neurospora contain a chromodomain-containing protein in a PRC1-type complex ?Neurospora , the mechanism remains unknown. We revised the Discussion section to provide additional background and context for H3K27me2/3 in Neurospora and we have specifically addressed conservation (or lack thereof) of the PRC2 and PRC1 complexes. Briefly, as in some other organisms, no obvious homologs of the PRC1 components are apparent in the Neurospora genome. However, it is clear that the presence of H3K27me2/3 is associated with repression . Ongoing work is directed at discovering the mechanism behind H3K27me2/3-associated repression in Neurospora, and the mechanism of repression by H3K27me2/3 is not completely clear even in organisms sporting obvious PRC1 homologs. Moreover, recent work has called into question seeming established models for the role of PRC1 in repression .Although it is clear that PRC2 is repressive in The second point that might be interesting to discuss is that unlike in flies and mammals, in plants, not all forms of PRC2 are inhibited by active marks . Specifically, PRC2 complexes containing the Su(z)12 ortholog EMF2 are inhibited by H3K4me3 but PRC2 containing the Su(z)12 ortholog VRN2 are not inhibited by this mark . So it remains to be tested whether the HMTase activity of Neurospora PRC2 is at all inhibited on H3K36me2/3 nucleosomes in vitro.Neurospora, PRC2 complexes that consist of different components could potentially explain why only certain H3K27me2/3 genes are derepressed when ASH1 is inactivated. Of possible relevance, earlier work from our lab showed partial loss of H3K27me2/3 associated with deletion of the gene encoding Neurospora p55 , which is thought to be a component of Neurospora PRC2. The mutant specifically loses H3K27me2/3 at telomere-proximal regions. This is in stark contrast to the ASH1 mutant, which retains telomere-proximal regions. In fact, when we categorize H3K27me2/3-marked genes into NPF-dependent and ASH1-dependent we see they each influence a similar number of genes . If NPF and ASH1 were randomly regulating H3K27me2/3 genes we would expect 30% of these genes would be regulated by both NPF and ASH1; however, we see only 17% overlap, suggesting they are actively regulating separate compartments of H3K27me2/3-marked genes.Schmitges et al. (2011) is an interesting paper and may be relevant to the observations we report. Although not known in