Npr1 (encoding NPRA) and receptor function. ANG II significantly decreased NPRA mRNA and protein levels and cGMP accumulation in cultured mesangial cells and attenuated ANP-mediated relaxation of aortic rings ex vivo. The transcription factors, cAMP-response element-binding protein (CREB) and heat-shock factor-4a (HSF-4a) facilitated the ANG II-mediated repressive effects on Npr1 transcription. Tyrosine kinase (TK) inhibitor, genistein and phosphatidylinositol 3-kinase (PI-3K) inhibitor, wortmannin reversed the ANG II-dependent repression of Npr1 transcription and receptor function. ANG II enhanced the activities of Class I histone deacetylases (HDACs 1/2), thereby decreased histone acetylation of H3K9/14ac and H4K8ac. The repressive effect of ANG II on Npr1 transcription and receptor signaling seems to be transduced by TK and PI-3K pathways and modulated by CREB, HSF-4a, HDACs, and modified histones. The current findings suggest that ANG II-mediated repressive mechanisms of Npr1 transcription and receptor function may provide new molecular targets for treatment and prevention of hypertension and cardiovascular diseases.The two vasoactive hormones, angiotensin II  and atrial natriuretic peptide  antagonize the biological actions of each other. ANP acting through natriuretic peptide receptor-A (NPRA) lowers blood pressure and blood volume. We tested hypothesis that ANG II plays critical roles in the transcriptional repression of  Both ANP and BNP are primarily synthesized in atrial myocytes and to a much lesser extent, they are synthesized in ventricular cells and stored in dense granules5. A third peptide, C-type natriuretic peptide (CNP), which is highly homologous to ANP and BNP, is predominantly present in the endothelial cells and the central nervous system6. ANP and BNP exhibit their major effects in diverse organ systems, including kidneys, adrenal glands, heart, vasculature, gonads, and adipose tissues12. The early discovery of three related natriuretic peptides (NPs) hormones, prompted the cloning and characterization of three distinct subtypes of natriuretic peptide receptors (NPRs). These NP receptors included: natriuretic peptide receptor-A (NPRA), receptor-B (NPRB), and receptor-C (NPRC), with binding characteristics of ANP and BNP to NPRA, CNP to NPRB, and all three NPs  to NPRC17. NPRA and NPRB, both of which contain a guanylyl cyclase (GC) domain, are also referred to, respectively, as GC-A/NPRA and GC-B/NPRB. The NPRA has generally been considered to be the primary receptor of ANP and BNP, the reason being that most of the physiological effects of these hormones are triggered by rapidly activating this receptor and the generation of its intracellular second messenger cGMP20. The recent studies have indicated that the ANP/NPRA system also has a central role in insulin resistance, obesity, and metabolic syndromes22.Atrial and brain natriuretic peptides (ANP and BNP) are endogenous cardiac hormones that regulate sodium excretion, water balance, and steroidogenesis, processes that are all largely directed toward reducing blood pressure and blood volume26, endothelin27, and vasopressin28, as well as other stimuli such as osmoregulation30, autoregulation19, and cytokines and growth factors31. The pressure hormone ANG II exhibits the vascular constrictive effects and the retention of sodium and body fluid, thereby, increases the vascular tone and blood pressure33. The two vasoactive hormones ANP (vasodilatory and hypotensive) and ANG II (vasoconstrictive and hypertensive) antagonize each other at all levels, including biochemical, molecular, and physiological effects37. Previous studies have demonstrated that ANP antagonizes ANG II-induced contraction of vascular smooth muscle cells (VSMCs) and mesangial cells (MCs), agonist-induced Ca2+ accumulation, and activation of mitogen-activated protein kinases (MAPKs) and protein kinase C (PKC) in different tissues and cell types44. On the contrary, ANG II also antagonizes ANP-induced GC activity and intracellular accumulation of cGMP45. ANG II has been shown to hydrolyze ANP-induced cGMP levels probably by stimulating the calcium-activated cGMP-dependent phosphodiesterase, thereby antagonizing the ANP-induced inhibitory effects on MAPKs, PKC, and tyrosine kinase (TK) activity24. Interestingly, a recent study using Npr3 (coding of NPRC) gene-knockout mice has indicated that NPRC, a non-guanylyl cyclase containing NP receptor prevents the progression of ANG II-dependent atrial fibrillation and remodeling46. This raises the possibility that the factors other than ANP/NPRA/cGMP, might also play a pivotal role in the cardiovascular protective effects involving NPRC, which needs to be further studied.The expression and activation of NPRA is regulated by various hormonal agents, including angiotensin II (ANG II)Npr1 gene has demonstrated the presence of ANG II-dependent negative regulatory cis-elements upstream of the transcription start site (TSS)25. However, the molecular mechanism of ANG II-dependent transcriptional repression and function of NPRA has not been elucidated. Also, the 5′-flanking region of Npr1 promoter contains potential binding motifs for a variety of transcription factors, including Nkx-2.5, AP-4, GATA-1/2, cAMP-response element-binding protein (CREB), and heat-shock factor-4a (HSF-4a)47. However, the functional significance of these transcription factors in relation to Npr1 gene expression has not been determined. In the present study, we examined the functional consequences of ANG II-stimulated cis-acting response elements, their binding sites, and mechanisms regulating the Npr1 promoter activity. We demonstrated the role of CREB and HSF-4a in ANG II-mediated repression of Npr1 transcription, expression, and physiological functions using cultured male mouse mesangial cells (MMCs) in vitro and intact male mouse aortic rings ex vivo.Promoter analysis of the Npr1 promoter, 5′ deletion constructs were sequentially analyzed for luciferase activity. The cells were transfected with deletion constructs and treated with 10 nM ANG II. As compared with untreated control cells, the constructs ΔA4 (−1182/+55 bp) and ΔA9 (−941/+55 bp) displayed significant decreases  in luciferase activity  construct, with the maximum reduction occurring at a concentration of 10 nM ANG II at 16 h after ANG II treatment  blocker, and PD 123319, an ANG II Type 2 receptor (AT2R) blocker, are shown in Fig. Npr1 promoter deletion constructs ΔR1 and ΔR5 is shown in Fig. 2R blocker PD123319 had no effect on ANG II-mediated repression of Npr1 promoter activity , tyrosine kinase (TK), and phosphatidylinositol 3-kinase (PI-3K), are detailed in Fig. Npr1 .To elucidate the role of transcription factors HSF-4a and CREB, the consensus sequence cgttctt was mutated to ccatgat in ΔR1 construct (−1182/−1127 bp) and the sequence atgccgtca was mutated to atccggtga in ΔR5 construct (−984/−914 bp), as shown by schematic representation in Fig. -4a Fig. . A 55% rity Fig. . On the ity Fig. . The treity Fig. . Howeverity Fig. . In the Electrophoretic mobility shift assay (EMSA) was performed with ∆R1a (−1156 to −1127 bp) and ∆R5a (−959 to −914 bp) probes, respectively, containing HSF-4a and CREB binding sites. The ∆R1a region showed a binding pattern with band I (arrow) corresponding to HSF-4a transcription factor and an additional band II (arrow) was also observed Fig. , lane 3.Npr1 repression, we determined the effect of ANG II on histone deacetylases (HDACs) expression and activity levels of class I HDACs . Treatment with increasing concentrations of ANG II augmented total HDAC activity by more than 3-fold as compared with activity in control cells  relaxed aortic rings that had been contracted with PGF2α. However, overnight treatment of aortic rings with 100 nM ANG II significantly antagonized the ANP response curve . Post-hoc analysis showed significant inhibition at 10 nM (p < 0.001) and 100 nM (p < 0.05) concentrations of ANG II  and AT2R (PD123319) subtypes. The AT1R antagonist, candesartan, but not the AT2R antagonist, PD123319, blocked ANG II-mediated repression of Npr1 in both the ΔR1 and ΔR5 constructs of the Npr1 promoter, suggesting the involvement of AT1 receptor subtype in this repression. Nevertheless, at higher concentrations of AT2R antagonist, PD123319, there was a slight repressive effect on Npr1 promoter activity in both the ANG II responsive regions of the Npr1 gene. Earlier, we observed that Npr1 promoter showed repressive activity in the presence of ANG II56.The results from the deletional analysis of Npr1 promoter. The ANG II-mediated repression of Npr1 promoter ΔR1 construct could be blocked by genistein, a TK inhibitor, suggesting the involvement of TK in the transcriptional repression of Npr1. In the ΔR1 construct, there is a 100% match of the DNA sequence with a putative heat-shock element, which binds to heat-shock factors. It has been reported that HSF-4a represses basal transcription through interaction with transcription factor IIF (TFIIF), which occurs through inhibition of an early step in formation of the preinitiation complex57. Interestingly, ANG II causes an increase in heat-shock factors and that heat-shock protein 90 complex negatively regulates NP receptors59. Moreover, genistein has been shown to inhibit herbimycin A-induced over-expression of inducible heat-shock protein corresponding to 70 kDa60. Our UV cross-linking experiments and gel mobility shift assays demonstrated the formation of DNA-HSF-4a binding complex, which was enhanced in ANG II-treated cells. The novelty in the present study stems from the fact that HSF-4a is activated by ANG II, which negatively regulates Npr1 gene transcription and receptor function. To our knowledge, this is the first report demonstrating the role of ANG II-dependent activation of TK in regulating HSF-4a in transcriptional repression of Npr1.In the current work, we examined the involvement of protein kinases in the signal transduction pathway, mediating the repressive effect of ANG II on Npr1 construct was repressed in the presence of ANG II; this repression could be blocked by genistein or wortmannin, suggesting the involvement of both TK and PI-3K in this pathway. The ΔR5 construct contains a putative cAMP-response element, TGCCGTCA (at −932 bp position), which is recognized by the transcription factor CREB, one of the few reported cis-elements through which ANG II has been shown to regulate gene expression. Interestingly, our results indicate that ANG II was able to activate the recruitment of CREB to Npr1 promoter and exerted the repressive effect on Npr1 gene transcription and function. UV cross-linking and EMSA exhibited enhanced DNA-CREB binding complex in ANG II-treated cells as compared with untreated control cells. Western blot results confirmed the enhanced phosphorylation of CREB in the presence of ANG II. It has previously been suggested that ANG II promotes the phosphorylation of CREB at Ser133 through an ERK1/2-dependent mechanism63. CREB activity has also been shown to be regulated by PI-3K/Akt signaling in Jurkat T leukemia cells treated with tumor necrosis factor-related apoptosis-inducing ligand64 and tyrosine kinase B/PI-3K/Akt pathway in SH-SY5Y cells treated with brain-derived neurotrophic factor65. Although CREB is most often described as a positive transcription factor, several reports have shown that it can also inhibit the transcriptional activity of several gene promoters such as those of c-fos and somatostatin67. It is noteworthy to mention that in our preliminary studies, the transfection of MMCs with only either HSF-4a or CREB expression plasmids alone without ANG II, did not exhibit any discernible effect on the negative repression of luciferase activity and Npr1 transcription (data not shown). It is implicated that the hormonal signal of ANG II is required for the activation of HSF-4a and CREB to exert a repressive effect on Npr1 promoter activity and its gene transcription.In the present study, promoter activity of the ΔR5 69. Evidence suggests that ANG II treatment induces epigenetic modifications, including changes in HDACs expression and activity, which is associated with ANG II–induced tissue hypertrophy and fibrosis71. Earlier, it was shown that ANG II treatment of intact E12.5 mouse metanephroi grown ex vivo increased HDAC1 and decreased total acetylated histone H3 protein levels72. Recently, it has been shown that in ApoE gene-knockout mice with abdominal aortic aneurysms, infusion of ANG II for 4 weeks increased expression of class I HDAC1, 2, and 3, as well as expression of class II HDAC 4 and 7; it also decreased acetylation levels of H3-K1873. On the other hand, selective inhibition of class I HDACs led to potent suppression of ANG II-mediated cardiac fibrosis and hypertrophy by targeting cardiac fibroblasts and bone-marrow-derived fibromyocytes74. These findings are consistent with our observation that ANG II treatment repressed acetylation of H3-K9/14 and H4-K8 and enhanced HDAC activity. Our results support the notion depicting a model that ANG II facilitated the recruitment of transcription factors, CREB and HSF-4a to Npr1 promoter via AT1R, resulting in the activation of TK and PI-3K signaling pathways, which exerted the repressive effects on Npr1 gene transcription and function . An in-vitro Site-Directed Mutagenesis kit was purchased from Stratagene . A direct enzyme-linked immunosorbent assay (ELISA) kit for cGMP assay was purchased from Enzo Life Sciences . Candesartan and PD 123319 were generously given to us by Dr. L. Gabriel Navar . Genistein (cat. no. CAS446-72-0), wortmannin (cat. no. CAS19545-26-7), and H-89 dihydrochloride (cat. no. CAS127243-85-0) were purchased from Sigma-Aldrich Co. . MGCD0103 (cat. no S1122) was obtained from Selleckchem . HSF-4a transcription factor plasmid was given to us by Dr. Nahid Mivechi . CREB transcription factor plasmid was a gift from Dr. Jane Reusch .We purchased pGL3-basic vector, pRL-TK, pGL3-control plasmids, and a dual luciferase assay system from Promega . A plasmid isolation kit was bought from Qiagen . Sequence-specific oligonucleotides were purchased from Midland Certified Reagent Company . Cell culture media, fetal calf serum (FCS), ITS , and lipofectamine-2000 were purchased from Invitrogen . ANG II (Ang II: cat. no. H1705) was purchased from Bachem America . An RNeasy mini-kit for total RNA isolation, RT75. Adult mice (30 g) were euthanized by deep anesthesia with isoflurane inhalation. Thoracic part of aorta was isolated and rings were prepared as earlier reported76. Animals were used under the protocol approved by the Institutional Animal Care and Use Committee (IACUC) at the Tulane University Health Sciences Center and were conducted in compliance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animal.Mice used in the present studies were C57/Bl6 wild-type and produced at Tulane Vivarium. Mice were housed under 12:12 h light-dark cycle at 25 °C and fed regular chow (Purina Laboratory and tap water ad libitum as previously describedNpr1 gene promoter region upstream of the promoterless firefly luciferase gene in the pGL3-basic vector as previously described25. All the positions in the following promoter constructs are relative to TSS and were generated by polymerase chain reaction (PCR) using the pNPRA-luc1  as a template and DNA polymerase (elongase): the ∆A1 (−1349 to +55 bp), ∆A2 (−1278 to +55 bp), ∆A3 (−1226 to +55 bp), ∆A4 (−1182 to +55 bp), ∆A5 (−1128 to +55 bp), ∆A6 (−1075 to +55 bp), ∆A7 (−1026 to +55 bp), ∆A8 (−984 to +55 bp), ∆A9 (−941 to +55 bp), and ∆A10 (−882 to +55 bp). The PCR primers used are listed in Supplementary Table All promoter-luciferase reporter constructs were made by cloning the DNA fragments of various lengths of mouse Npr1 promoter region was done at the upstream of SV-40 promoter firefly luciferase gene in the pGL3-promoter vector as previously reported25. The ∆R1 (−1182 to −1127 bp), ∆R2 (−1128 to −1072 bp), ∆R3 (−1071 to −1028 bp), ∆R4 (−1026 to 986 bp), and ∆R5 (−984 to −914 bp) constructs were generated by PCR using the pNPRA-luc1 (−1982 to +55 bp) as a template and DNA polymerase (elongase). All the forward primers  contained a MluI restriction site; whereas the reverse primer contained a BglII restriction site at the 5′ ends. The PCR primers used are listed in Supplementary Table The cloning of the smaller DNA fragments from −1182 to −914 bp of 35. Cultures were maintained at 37 °C in a 5% CO2/95% O2 humidified atmosphere. For all experiments, cells were used between 4 to 12 passages. To study the effect of ANG II, cells were seeded in 24-well plates at 80% to 90% confluence. The cells were washed twice with serum-free assay medium containing 0.1% bovine serum albumin (BSA) and treated with 10 nM ANG II in fresh assay medium in the absence or presence of 100 nM dihydrochloride, 100 nM wortmannin, 100 nM genistein, or 1 µM MGCD0103. The cells were harvested at the indicated time intervals and lysed essentially as described earlier77.MMCs were cultured in Dulbecco modified Eagle’s medium (DMEM) supplemented with 10% FCS and ITS as previously described25. The medium was changed after 24 h. The cells were harvested after 48 h by using passive lysis buffer (Promega). Luciferase activity was measured by TD 20/20 luminometer  with 20 µl cell extract using a dual luciferase reporter assay system. In the transfection experiments, a pGL3-control vector containing both the SV40 promoter and enhancers was used as a positive control; the empty pGL3-basic vector was used as a negative control. The assays were performed in triplicate in 6–8 independent experiments. Results were normalized for the transfection efficiency as relative light units per renilla luciferase activity.MMCs were seeded in 12-well plates at a density producing ~80% confluence. After 24 h, the cells were transfected using lipofectamine-2000 reagent according to the manufacturer’s instructions, with 1 µg of test plasmid and 0.3 µg of pRL-TK carrying the renilla luciferase gene downstream of the thymidine kinase promoter, which was used as internal transfection control as earlier described77. The protein concentration of the lysate was measured with a Bradford protein detection kit . Western blot assay was done as previously described77. The cytoplasmic fraction (50–70 µg) or nuclear extract (40–60 µg) was mixed with sample loading buffer and electrophoresed for 2 h, then transferred to a nylon membrane. The membrane was blocked with 1x Tris-buffered saline-Tween 20  containing 5% fat-free milk for 1 h, then incubated overnight in TBST containing 5% fat-free milk at 4 °C with primary antibody (1:250 dilution). The membrane was treated with corresponding secondary anti-rabbit or anti-mouse horseradish-peroxidase (HRP)-conjugated antibodies. Protein bands were developed using a SuperSignal West Femto Chemiluminescent kit and visualized using a FluorChem detection system from Proteinsimple . The intensity of protein bands was quantified by AlphaView software . The antibodies used in Western blot assay are listed in Supplementary Table Whole cell lysate and nuclear extract were prepared as described earlier19. The cell suspension was subjected to five cycles of freeze and thaw and then centrifuged at 10,000 × g for 15 min. The supernatant thus collected was used for cGMP assays using a direct cGMP complete Elisa kit according to the manufacturer’s instructions .Twenty-four hours after plating, MMCs were made serum-free for 12 h and treated with ANG II (10 nM) for another 24 h. Cells were stimulated with ANP at 37 °C for 15 min in the presence of 0.2 mM 3-isobutyl-1-methylxanthine (IBMX), washed three times with phosphate-buffered saline (PBS), and scraped into 0.5 N HCl, as previously described2 First Strand cDNA kit from Qiagen. Primers for amplification of Npr1 and β-actin were from Qiagen. PCR amplification was done in triplicate in a 25-µl reaction volume using RT2 Real-Time SYBR Green/ROX PCR Master Mix and PCR conditions as previously described78. Control experiments were done with RNA samples but without reverse transcriptase. The specific primers for ß-actin gene were included in the PCR reaction mixture as an internal control.Confluent MMCs were treated with or without ANG II (10 nM) in assay medium. After harvesting the cells, total RNA was extracted and 1 µg of total RNA was reverse transcribed, using a RTin-vitro Site-Directed Mutagenesis kit (Stratagene). The consensus sequence of HSF-4a and CREB along with the mutant sequence is listed in Supplementary Table Npr1 gene promoter, pNPRA-Luc125, was used as a template for generating mutations. The double-stranded DNA template pNPRA-Luc1 was alkaline denatured, annealed with the mutagenic oligonucleotide and selection oligonucleotide in annealing buffer, incubated at 75 °C for 5 min, and allowed to cool slowly to 37 °C. The mutant strand was synthesized with T4 DNA polymerase and T4 DNA ligase in the presence of synthesis buffer and incubated at 37 °C for 90 min. The DNA thus synthesized was transformed in XL-10 Gold super competent cells (Stratagene) and plated in LB amp agar plates containing gene editor antibiotic mix. The probable clone was confirmed by sequencing.The HSF-4a and CREB transcription factor mutants, ΔR1 mutant (−1182/−1127 bp) for HSF-4a and ΔR5 mutant (−984/−914 bp) for CREB, were constructed by using the -type and mutant oligonucleotides corresponding to HSF-4a and CREB transcription factor binding sites were commercially synthesized and labeled at the 5′-end by phosphorylation of the 5′ hydroxyl ends with [γ32-P]ATP, using T4 polynucleotide kinase enzyme as previously described79. The radiolabeled sense oligonucleotide was annealed with the antisense unlabeled oligonucleotide in a 1:1 molar concentration by adding an equal volume of 2 x annealing buffer and incubating for 5 min in boiling water, then slowly cooling. The annealed oligonucleotides were purified on a Sephadex G-50 column. EMSA was done using ATP-labeled and annealed oligonucleotides as previously described80. For gel retardation, reaction mixture was prepared by adding 5–10 μg of nuclear extracts to 40,000–50,000 cpm of probe in 1x binding buffer containing 10 mM Tris (pH 7.5), 1 mM MgCl2, 0.05 mM EDTA, 0.05 mM DTT, 50 mM NaCl, 4% (v/v) glycerol, and 1 µg of nonspecific DNA. The reaction mixture was incubated for 30 min on ice. The DNA-protein complex was resolved from the free-labeled DNA by electrophoresis in a 4% native polyacrylamide gel at 100 V for 2 h and after drying gel was autoradiographed. For competition assays, 100-fold excess molar concentrations of unlabeled probe were added in the reaction mixture as a competitor.The wildNpr1 promoter was 32P-end-labeled and annealed for the EMSA reaction as described80. Binding reactions were set up as those for EMSA. After incubation on ice for 30 min, the reaction mixture was UV-cross-linked by pipetting onto parafilm and irradiated in a UV-cross linker. Samples were resolved by electrophoresis on 10% denaturing polyacrylamide gel and exposed to X-ray film for autoradiography.To determine the molecular mass of nuclear binding proteins to the consensus sites, we did a UV cross-linking experiment. Oligonucleotide spanning the consensus site in the 78. In brief, cells were harvested and suspended in 1x pre-lysis buffer, kept on ice for 10 min, and centrifuged at 10,000 × g for 1 min at 4 °C. The supernatant was removed; the cell pellet was resuspended in 3 volumes of lysis buffer, incubated on ice for 30 min, and centrifuged at 14,000 × g for 5 min at 4 °C. Balance-dithiothreitol (DTT) buffer (0.3 volumes) was added to the supernatant, which was then stored at −80 °C. The protein concentration of the eluted histone was estimated using a Bradford protein detection kit , using BSA as a standard.Total histone was extracted from ANG II-treated and untreated MMCs, using a total histone extraction kit  as earlier reported78. HDAC enzyme activity was calculated by measuring the amount of HDAC-deacetylated product, which was directly proportional to HDAC enzyme activity. Absorbance was read at 450 nm. Results were calculated using a standard curve according to the manufacturer’s instructions and expressed as ng/min/mg protein.Total HDAC activity was measured in nuclear extracts prepared from ANG II-treated cells using a colorimetric ELISA assay kit from Active Motif  essentially as previously described76. Immediately after thoracotomy, the thoracic aorta was removed and placed in cold Dulbecco’s PBS and cleaned by removing the surrounding fat and connective tissue. For experiments, the aorta was cut into 3 to 4 mm rings. After 2 h incubation in DMEM with 0.1% BSA, the aortic rings were treated with ANG II for 20 h. Aortic rings were then homogenized by sonication in lysis buffer, centrifuged, and supernatant was collected and stored at −80 °C for Western blot assay. For qRT-PCR, ANG II-treated and control rings were homogenized with 1.5-ml lysis buffer and RNA was extracted by an RNeasy mini-kit, following the manufacturer’s protocol .The male mice (C57/Bl6) were euthanized by deep anesthesia with isoflurane inhalation. Thoracic aorta was isolated and aortic rings were prepared using the previously described protocol with a minor modification81. Vessels were preconstricted with prostaglandin F2α (PGF2α) and treated with increasing concentrations of ANP. Data are expressed as percent relaxation from PGF2α contraction. For baseline control experiment, aortic rings were incubated overnight in either control media or in media containing ANG II. The next day, rings were exposed to increasing concentrations of ANG II in the wire myograph.Aortas were isolated and excised from C57/Bl6 male mice, cut into 2 mm rings, and incubated in DMEM containing 0.1% BSA. After 2 h, 100 nM ANG II was added to the aortic vessels. After 6 h, another dose of ANG II was added, and incubation was continued. Vessels were mounted on a wire myograph and experiments were completed as previously describedt test. Data was also analyzed using Two-Way ANOVA. Repeated measures by both ANG II and ANP treatments and Sidak’s multiple comparisons test were used. The differences were considered significant with the probability value < 0.05.Statistical analysis was done using GraphPad prism software . The results are expressed as mean ± SE. Statistical significance was evaluated by one-way ANOVA and Student Supplemental Information.


Unexpectedly, the molecular pathogenesis of expanded CAG/CTG diseases does not include changes in 3D chromatin conformation. DMPK and HTT loci from DM1 and HD–derived cells. We find that allele sizes ranging from 15 to 1700 repeats displayed similar chromatin interaction profiles. This was true for both loci and for alleles with different DNA methylation levels and CTCF binding. Moreover, the ectopic insertion of an expanded CAG repeat tract did not change the conformation of the surrounding chromatin. We conclude that CAG/CTG repeat expansions are not enough to alter chromatin conformation in cis. Therefore, it is unlikely that changes in chromatin interactions drive repeat instability or changes in gene expression in these disorders.Expanded CAG/CTG repeats underlie 13 neurological disorders, including myotonic dystrophy type 1 (DM1) and Huntington’s disease (HD). Upon expansion, disease loci acquire heterochromatic characteristics, which may provoke changes to chromatin conformation and thereby affect both gene expression and repeat instability. Here, we tested this hypothesis by performing 4C sequencing at the  The genome is organized into hierarchical chromatin contact domains  binding at sites flanking the expanded GAA repeats  and Huntington’s disease (HD). They are part of a larger group of diseases caused by the expansion of short tandem repeats (STRs)   293–derived cells. We confirmed the alteration of chromatin interactions in a FXS patient cell line with ~935 CGG repeats. However, we found no evidence of changes in chromatin interactions caused by expanded CAG/CTG repeats in HD or DM1 patient cells. This suggests that 3D conformational changes are unlikely to underlie the alterations in transcriptional output or genetic instability of disease-associated expanded CAG/CTG repeat loci. This was consistent in different cell types, genetic backgrounds, and in the presence of specific heterochromatic marks. Therefore, we conclude that changes in higher-order chromatin conformation are unlikely to contribute to the pathogenesis of expanded CAG/CTG repeat disorders.Two critical unknowns in this model are whether changes in higher-order chromatin structure are confined to CGG and GAA repeats or if this is general to daSTRs and whether changes in chromatin structure cause alterations in gene expression and repeat instability. The latter question is especially appealing in the context of the expanded CTG repeats in the 3′ untranslated region of the FMR1 locus upon CGG repeat expansion with 4C-seq . We also determined the chromatin conformation of an unrelated locus, ACTA1 on chromosome 1, to control for a potential effect of FXS on genome-wide chromatin conformation. We obtained three replicates of each 4C viewpoint and found that replicates from the same cell line showed good correlation in 4C fragments with at least 20 mapped reads . We identified interaction peaks and broader high-interacting regions that contacted the 4C viewpoints at frequencies higher than expected given their linear distance away from the viewpoints. These significant interactions were determined using two 4C-seq data analysis packages: FourCSeq . In addition, at the ACTA1 locus, we found that the interaction profiles were similar between the two cell lines within a 2-Mb region around the viewpoint , GM03620 (18 and 70 CAGs), and GM14044 (19 and 750 CAGs), referred to as HD-A, HD-B, and HD-C, respectively. We compared them to two cell lines from unaffected individuals . Their family relationships and their repeat sizes are shown in To assess whether chromatin interactions change upon CAG repeat expansion at the HTT locus, we used two 4C viewpoints within the HTT gene body—1 kb (HTT_d1) and 85 kb (HTT_d85) downstream of the CAG repeats in unaffected and HD patient cells (table S2). We also assessed the chromatin conformation of the unrelated ACTA1 gene. Replicates from the same cell lines showed good correlation in fragments with more than 20 mapped reads . Compared to the UN-B and UN-C cells, we found that the chromatin interaction profiles were similar within a 2-Mb region around the HTT viewpoints in all three HD patient LCLs  experiments with the same unaffected and HD patient cells. Differential gene expression analysis between the HD and unaffected LCLs identified 1183 significantly up-regulated and 1307 significantly down-regulated genes . Within a 2-Mb region centered around HTT, we identified 4 genes of 25 that showed significant differential expression between the HD patient and the unaffected cell lines (fig. S5A). This proportion was not statistically different from the proportion of differentially expressed genes in a 2-Mb region around ACTA1 . Given that the chromatin interactions did not significantly change between the HD and unaffected cell lines in the HTT region, our data suggest that changes in gene expression in cis occur independently of the chromatin conformation of expanded HTT alleles in HD patient cells.To determine the relationship, if any, of chromatin conformation at the HTT locus is not known to be associated with changes in local histone modifications and chromatin accessibility, which might cause changes in chromatin conformation. To determine whether an expanded CTG repeat locus with local heterochromatic chromatin marks is associated with altered 3D chromatin conformation, we used the unaffected cell lines UN-B and UN-C and two DM1 patient–derived LCLs . The DM1-A cell line harbored one expanded DMPK allele with 1700 CTGs and the DM1-B cell line, 1000 CTGs . Similar to our observation in HD individuals, changes in gene expression in cis occurred independently of the 3D conformation of the DMPK locus in DM1 patient cells.We confirmed by RNA-seq that transcriptional misregulation in DM1 patient cells occurs genome-wide as well as within a 2-Mb region centered around DMPK, only 4 of 131 (3%) CTCF binding peaks showed significant differential occupancy in DM1-A patient cells compared to the UN-B cell line (fig. S8). There were no binding sites with significant differences in CTCF occupancy out of 135 peaks in that same region when comparing the UN-B and DM1-B cell lines (fig. S8). Therefore, we concluded that there were no large-scale changes in CTCF occupancy in the DM1 patient LCLs either genome-wide or within a 2-Mb region around the expanded CTG repeats, which is consistent with the absence of changes in chromatin conformation observed by 4C-seq in these cells.To determine the extent of CTCF occupancy alterations in the DM1 patient cells, we performed ChIP sequencing (ChIP-seq) (fig. S8). At the genome-wide level, comparing DM1-A to UN-B cells, we found 1951 sites where CTCF binding changed significantly [false discovery rate (FDR) < 5%]. DM1-B cells harbored 833 sites with significantly altered CTCF occupancy genome-wide compared to UN-B cells (FDR < 5%). However, in a 2-Mb region centered around DM1 and HD are both dominantly inherited disorders wherein affected individuals are heterozygous for the expanded allele. Thus, one potential caveat in our data was that the presence of a normal-length allele could mask changes in the 3D chromatin interactions made by the expanded allele. To evaluate this possibility, we took advantage of the presence of at least one parental cell line for the DM1-A and HD-B individuals in our dataset  and 3A a15] or 270 CAGs [GFP(CAG)270] in isogenic cell lines. We obtained two clonal populations of HEK293 T-Rex Flp-In cells that contain a single, stably integrated construct containing CAG repeats within the intron of a green fluorescent protein (GFP) mini-gene controlled by a doxycycline-inducible promoter promoter . We perfpromoter . Thus, w15 and GFP(CAG)270 cells with doxycycline for 5 days and subsequently performed 4C-seq using the same GFP viewpoint. We observed that 2 of 17 regions of differential interactions overlapped fully or in part with high-interacting regions over a 2-Mb region (Some studies suggest that transcription can help define chromatin domain boundaries (b region . By contHTT and DMPK) and one ectopic locus in two different cell types. Our findings are supported by allele-specific analysis of 4C-seq chromatin interactions. Furthermore, increased CpG methylation and CTCF binding alterations at four sites in a 2-Mb region around DMPK did not affect the chromosome conformation of this locus in the DM1-A patient cells. This is especially relevant because CTCF is a key architectural protein involved in the demarcation of TAD and sub-TAD boundaries 270 and GFP(CAG)15 lines were previously characterized , and hygromycin (150 μg ml−1). Transcription of the GFP mini-gene was activated by culturing GFP(CAG)n cells with doxycycline at a final concentration of 2 μg ml−1 for 5 days.All LCLs were obtained from the Coriell Institute for Medical Research Cell Repository. They were grown at 37°C and 5% COHTT were produced with primers oVIN-1333 and oVIN-1334 (table S5). PCR products containing the CTG repeats from DMPK were amplified with primers oVIN-1252 and oVIN-1251 (table S5). For normal-length alleles, several PCRs were set up with MangoTaq (Bioline), and the products were gel-extracted and Sanger-sequenced with the same primers used for the amplification. For expanded alleles, small-pool PCRs were performed on the basis of a previously described protocol . PCR products with the CAG repeats from Bisulfite sequencing was performed according to a previously described method  with the following parameters: -q 20 --length 20 –paired. Reads were aligned using QuasR , with 10 cycles of 30 s “on” and 30 s “off.” Correct DNA fragmentation was verified by agarose gel electrophoresis. Immunoprecipitation was performed with 4 × 106 cells using a CTCF antibody (Diagenode) and the Diagenode IP-Star Compact Automated System robot (Diagenode). Results were analyzed using the StepOnePlus qPCR by Applied Biosystems (Thermo Fisher Scientific) with Applied Biosystems SYBR Green PCR Master Mix (Thermo Fisher Scientific). The primer sequences used for qPCR are listed in table S5. CTCF ChIP-seq libraries were prepared in triplicate using the Illumina TruSeq ChIP Sample Preparation Kit. We performed 1 × 50-bp single-end sequencing with Illumina HiSeq 4000.ChIP was performed according to the Diagenode Auto iDeal ChIP-qPCR Kit (Diagenode) standard protocol. Samples of 6 × 10P < 0.05 and FDR < 5%). First, we computed count information for each of the peaks in the consensus set. Bedgraph files  were created to visualize the tracks.CTCF ChIP-seq reads were demultiplexed with the bcl2fastq2 Illumina software (v 2.20.0). Data analysis was performed according to the ENCODE Transcription Factor ChIP-seq processing pipeline (phase 3). Demultiplexed reads were aligned to the GRCh38 human reference genome (GCA_000001405.15) using bowtie2 (v 2.3.4.3) , 150 mM NaCl, 5 mM EDTA, 0.5% NP-40, 1% Triton X-100, and 1× protease inhibitors]. The first digestion was performed with 200 U of Dpn II (New England Biolabs) and incubating for 4 hours at 37°C, then 200 U of Dpn II and overnight incubation at 37°C, and lastly 200 U of Dpn II and incubating 4 hours at 37°C. Dpn II digestion efficiency was assessed by agarose gel electrophoresis, and Dpn II was subsequently heat inactivated. The first ligation was performed at 16°C overnight with 50 U of T4 DNA ligase (Thermo Fisher Scientific) in 7 ml. Ligated samples were decross-linked with 30-μl Proteinase K (10 mg/ml) at 65°C overnight followed by ribonuclease A (RNAse A) treatment and phenol-chloroform purification. The second digestion was performed with 50 U of Bfa I (New England Biolabs) at 37°C overnight followed by Bfa I heat inactivation. The second ligation was performed at 16°C overnight with 100 U of T4 DNA ligase in 14 ml. The resulting 4C template samples were precipitated and purified with the QIAquick PCR purification Kit. 4C libraries were generated by amplifying 1-μg total of purified 4C template using the Expand Long Template PCR System (Roche) with 4C viewpoint primers with Illumina sequencing adapters that permitted multiplexing (in 50-μl PCRs), pooling reactions, and purifying the PCR products with AMPure XP beads to exclude products less than 130 bp. The 4C viewpoint primer sequences are listed in table S5. Single-end sequencing of pooled 4C libraries was performed on Illumina HiSeq 2500.4C library preparation was performed on the basis of a previously described protocol (www.gfeatbrowser.com). Significant chromatin interactions were identified with two 4C-seq data analysis packages: FourCSeq (v 1.18.0) . The UN-B cell line was used as the reference condition for all comparisons except for Demultiplexing, trimming, and mapping were performed using the BBCF HTSstation  , and its concentration was measured by NanoDrop . RNA quality was assessed on a Fragment Analyzer (Agilent Technologies), and all of the RNA samples had an RNA quality number between 8.7 and 10. RNA-seq libraries were prepared using 500 ng of total RNA and the Illumina TruSeq Stranded Total RNA Library Prep Kit with Ribo-Zero Gold (Illumina) according to the manufacturer’s protocol. Multiplexed samples were pooled in equimolar amounts, and 2 × 100-bp paired-end sequencing was performed on Illumina HiSeq 2500.P values were adjusted using the Benjamini-Hochberg multiple testing correction. Significantly differentially expressed genes were defined as those with adjusted P values less than 0.05 and log2 fold change greater than 0.5. Functional gene ontology enrichment analyses of significantly differentially expressed genes in HD and DM1 LCLs were performed using the DAVID tool (v 6.8) (RNA-seq reads were demultiplexed using the bcl2fastq Illumina software (v 2.20.0). Purity-filtered reads were trimmed with Cutadapt (v 1.3) , 150 mM NaCl, 5 mM EDTA, 0.5% NP-40, 1% Triton X-100, and 1× protease inhibitors). Cross-linked samples were digested at 37°C overnight with 400 U of Nla III (New England Biolabs) followed by Nla III heat inactivation. Samples were ligated at room temperature for 2 hours with 20 U of T4 DNA ligase (Thermo Fisher Scientific) in 500 μl. Ligated samples were decross-linked with 5-μl Proteinase K (10 mg/ml) at 65°C overnight, followed by RNAse A treatment and phenol-chloroform purification. Samples then digested overnight at 37°C with 50 U of Nsp I (New England Biolabs) followed by Nsp I inactivation. The second ligation was performed overnight at 16°C with 100 U of T4 DNA ligase in 14 ml. The resulting TLA circularized templates were purified with the QIAquick PCR Purification Kit. TLA libraries were generated by amplifying 800 ng of purified TLA template with TLA viewpoint primers (table S5). Paired-end sequencing (2 × 150 bp) of pooled TLA libraries was performed on Illumina HiSeq 4000. Demultiplexed reads were mapped using a custom TLA analysis pipeline using the Burrows-Wheeler Aligner mapping software (v 0.7.17) (Targeted locus amplification (TLA) was performed on the basis of a previously described protocol (


The identification of upstream transcription factors regulating the expression of a gene is generally not an easy process.  To facilitate this task, we constructed an expression cDNA library named Transcription Factor Expression Library (TFEL), which is composed of nearly all the transcription factors in the mouse genome. Genome-wide screening using this library (TFEL scan method) enables us to easily identify transcription factors controlling any given promoter or enhancer of interest in a chromosomal context-dependent manner. Thus, TFEL scan method is a powerful approach to explore transcriptional regulatory networks. TFEL, transcription factor expression library; VEGF, vascular endothelial growth factor; Fasn, fatty acid synthase; SREBP, sterol regulatory element-binding protein; HIF, hypoxia-inducible factor; LXR, liver X receptor; KLF, Kruppel-like factor.1. Generally, downstream targets of a transcription factor can be identified in a convincing way through transcriptome analyses after overexpression or knockdown of the transcription factor, or through investigating the binding interaction between the transcription factor and genomic DNA using chromatin immunoprecipitation (ChIP)-sequencing method. However, no assured methods of identifying upstream transcription factors regulating a specific promoter or enhancer exists, since the relatively poor amount of transcription factors expressed in protein pools or cDNA libraries makes purification difficult, while binding factor prediction using computer analysis based on binding motif databases suffers from high false-positive and false-negative rates in general2. Therefore, we constructed a comprehensive expression library of transcription factors in the mouse genome, the Transcription Factor Expression Library (TFEL), and developed a new method of identifying upstream transcription factors through an expression cloning technique using the TFEL (TFEL scan method). Using this TFEL scan method, we can easily identify the specific transcription factor or transcription factor complex trans-acting against any given cis-element of interest.Transcriptional regulation is widely involved in many biological processes of relevance such as cellular development, tissue differentiation, reprogramming and apoptosis as well as nutrient metabolism; in many cases, such processes involve complicated regulatory networksmouse transcription factor DNABook™ was purchased from DNAFORM, Tokyo, Japan3. Each spot on the sheets of DNABook was punched, and plasmid DNA was eluted in 10μl distilled water. Competent DH5α E. coli was transformed with 1μl of the eluted DNA solution, and plasmids were cloned in a standard manner.The Among 1,588 clones in the DNABook, 1,350 clones were on pFLCI vector, 204 clones on modified BluescriptI(+), 23 clones on pFLCIII, and 11 clones were on pFLCII.nyahagi-tky@umin.ac.jp)The clones on pFLCI were digested by restriction enzymes EcoRI and BamHI, clones on pFLCII by XhoI and BamHI, pFLCIII by Bsu36I and BamHI, and clones on modified BluescriptI(+) were cut by SacI and XhoI on a 37 °C heat block. The restricted DNA fragments were separated on agarose gel  by electrophoresis, and verified if they are correspondent with the predicted band pattern calculated from the database (GenBank / EMBL / DDBJ) sequence. Unfortunately, information about which clones are included in the DNABook cannot be made public due to a confidentiality agreement. For more details, please contact the corresponding author via email . To facilitate this process, we first modified pcDNA3.1 vector and inserted the PCR-amplified kanamycin-resistant gene from pENTR4 vector (Invitrogen) into the original pcDNA3.1 at the SmaI-SalI sites by replacement of neomycin-resistant gene to make the vector resistant to kanamycin. By this modification, we could ligate an insert DNA fragment to the pcDNA3.1 vector directly without agarose gel isolation of the insert from vector backbone fragment and undigested plasmid after restriction enzyme digestion of DNABook clones, which are ampicillin-resistant.Because DNABook clones were on as many as 23 versions of various vectors : 2), multiple strategies based on the calculation of restriction sites were used to construct TFEL clones.For pFLCI clones, the combination of EcoRI and BamHI digestion was first tried to cut insert fragments out of vectors when inserts were not predicted to be cut by the two restriction enzymes, and excised fragments were inserted into pcDNA3.1(-) vector at EcoRI and BamHI sites. For pFLCI clones whose inserts were cut by EcoRI or BamHI, SfiI digestion and ligation to pcDNA3.1(-)-SfiI (XhoI-EcoRI part of pcDNA3.1(-) multiple cloning site was replaced with synthesized SfiI sequence) was next attempted. For clones to which neither strategy was applicable, either EcoRI-ApaI (into pcDNA3.1(+)), EcoRI-KpnI (into pcDNA3.1(-)), or SacI-BamHI (into pcDNA3.1(+)-SacI-mod whose SacI site is modified to be unique by introducing mutation to the other SacI site outside of the multiple cloning site) strategy was tried. For the other 73 clones to which these strategies were not applicable, SfiI digested multiple fragments were inserted into pcDNA3.1(-)-SfiI at one time. When another restriction enzyme such as ApaI, KpnI, NotI, or ScaI could be used to cut the vector backbone without cutting the insert, this treatment was done at the same time to reduce the possibility of a vector fragment being cloned into pcDNA3.1.For modified BluescriptI(+) clones, the SfiI (into pcDNA3.1(-)-SfiI) or XhoI-SacI (into pcDNA3.1(-)-SacI-mod) strategy was used. For 47 plasmids on ZA vector whose orientation of the insert is opposite to that of others, corresponding “(+)” version of pcDNA3.1 vectors were used. 7 clones on ZX vector had corrupted 5’-side SfiI and XhoI sites, therefore KpnI-SfiI, KpnI-BglII, or BssHII strategy was used instead.For pFLCII clones, the XhoI-BamHI strategy was used when applicable. Others were excised with SacI and cloned into SacI site of pcDNA3.1(-)-SacI-mod, after which the inserted direction was checked, and clones constructed in the correct direction was selected.pFLCIII clones were ligated using I-CeuI and PI-SceI.Vegfa-luc plasmid (-1210 to +246) was generated from a PCR-amplified fragment inserted into pGL2-basic vector  at SmaI site. The PCR primers used were 5’-AAGATGAACCGTAAGCCTAGGCT-3’ and 5’-AACCGTTGGCACGATTTAAGA-3’ and amplification was performed according to a 3-step method. The ratFasn-luc plasmid (-397 to +28) was prepared as described previously4. The mouseSrebf1c-luc plasmids (-2.2k to +40 and -249 to -144) were constructed as described elsewhere6.The mouseTo screen transcription factors regulating a target promoter, TFEL clones were co-transfected with the specific promoter-luc plasmid into HEK293 cells using SuperFect Transfection Reagent . The screenings were performed by co-transfecting 10–20 clones with each luciferase plasmid per one well. Transfection was performed using 0.5μg of plasmid DNA pooled equally from 10 or 20 TFEL clones per well. The luciferase activity in transfectants was measured on a luminometer (BERTHOLD) with a standard assay kit . First screen was performed with 10 or 20-clone pools per well, and significantly shifted pools were further tested with one clone at a time.All the data from a luminometer were output as Excel sheets, and directly input to graphs. All the graphs were drawn in Microsoft Excel 2016 (16.0.5083.1000).7. We retrieved all the clones in the DNABook usingEscherichia coli transformation. After checking the band patterns using several restriction enzymes and sequencing if necessary, each excised insert DNA fragment was transferred to a pcDNA3.1 expression vector. In this manner, we constructed a pcDNA3.1 expression library composed of 1,588 clones of genome-wide and nonredundant mouse transcription factors.To construct the TFEL expression plasmid library, we used clones from the RIKEN FANTOM libraries . The clo8).8 shows three cases in which the upstream transcription factors influencing well -known promoters were examined: A, vascular endothelial growth factor (gene name:Vegfa), B, fatty acid synthase (gene name:Fasn), and C, sterol regulatory element-binding protein (SREBP)-1c (gene name:Srebf1c). In these cases, well-known determinant transcription factors  were identified as expected. These screenings were performed by co-transfecting 10–20 clones with each luciferase plasmid per one well. Next, we evaluated the validity of this expression cloning method (TFEL scan method) using the TFEL 8.FigurThese results demonstrated that the identification of upstream transcription factors using this expression screening method works well.6. As shown in9, KLF4 was picked up as a candidate from the first screening performed by co-transfecting 20 clones with an SREBP-1c promoter luciferase plasmid in the presence of an LXR ligand T0901317. Further screening of the clones one by one revealed that KLF4 and KLF15 interacted with and suppressed LXR on the SREBP-1c promoter  interacting with a specific transcription factor on a specific gene promoter. To this end, we screened interacting partner(s) for LXR on the SREBP-1c promoterIn the present study, we clearly demonstrated that our new method of TFEL scan is a powerful approach to explore transcriptional regulatory networks by identifying transcription factor(s) involved in the regulation.Vegfa-luc,Fasn-luc andSrebf1c-luc), we could easily identify well-known determinant transcription factors  as expected (8). In addition, we recently succeeded in identifying the binding transcription factor for an important SNP at the diabetes-associatedTCF7L2 gene locus through our TFEL scan method10. It is suggested thatTCF7L2 is the single largest effect of a common SNP on type 2 diabetes risk in European populations11.First, we showed that in three cases , cannot be identified by mere overexpression without ligands (data not shown). Therefore, it may depend on transcription factors whether they can be detected without ligands/additional activators or not.8). Therefore, the TFEL scan method was proved to be similarly effective even in these more complex cases.Next, we additionally succeeded in identifying transcription factors forming a complex such as LXR and KLF4 8. There1. This cooperation is often mediated by direct physical contact between two or more transcription factors, forming homodimers, heterodimers, or larger transcriptional complexes12. In fact, it has been estimated that approximately 75% of all metazoan transcription factors heterodimerize with other factors13. Thus, the importance of transcription factor combinations has been highlighted more and more, and the mapping of the combinatorial interactions among transcription factors has been attempted using a mammalian two-hybrid system12. Our successful results identifying KLFs as interacting partners of LXR onSrebf1c promoter clearly showed that the expression cloning method using the TFEL (TFEL scan method) may also be useful for exploring interacting partner(s) among transcription factors on a specific DNA fragment of interest. In contrast, binding site prediction based on binding motif databases suffers from high false-positive and false-negative rates in general9. In particular, the sequence-based approach has no power to predict transcription factor complexes formed by protein-protein interaction. Conversely, the search for protein-protein interactions among transcription factors alone, for example, using a mammalian two-hybrid system, may not be sufficient to elucidate the regulatory complexes in certain situations including the present case of LXR-KLFs complex, because complexes like LXR-KLFs are also dependent on DNA binding and therefore locus-specific. This relatively weak interaction supported by the DNA backbone enables gene-specific regulations and may possibly give more diversity to transcriptional networks. Thus, our strategy of screening for the transcription factor complex in a chromosomal context-dependent manner using the TFEL scan method can be a very effective and powerful approach for exploring sophisticated transcriptional networks in detail, because this screening system is performed under more physiological conditions than traditional methods.It is well known that the transcriptional output of a gene is due to the joint activity of many transcription factors, the binding and activation of which are highly interdependentOur initial approach using a mixed pool assay handling 20 clones at a time failed to detect KLF15 in the first screening described above. Interference among the co-transfected transcription factors in the pools was thought to have affected the results and veiled the KLF15-LXR interaction, suggesting that a single clone assay is preferable, especially for screening for combinatorial interactions. In relation to this point, interactions with endogenous transcription factors should also affect the results to varying degrees, and screenings in different cell lines should provide more information.7. For example, among the 17 KLF family members, 4 KLFs  are missing. We are planning to fix this problem and to complete the construction of a comprehensive expression library in the near future.Our TFEL library is based on the Mouse Transcription factor DNABook™, and a small portion of transcription factors are missing, as they previously reportedIn summary, the present study clearly demonstrates that our expression cloning method using the TFEL (TFEL scan method) enables us to efficiently identify regulatory transcription factors as well as to elucidate combinatorial interactions among transcription factors in a chromosomal context-dependent manner. Thus, the TFEL scan method is a powerful approach to explore transcriptional regulatory networks.https://doi.org/10.6084/m9.figshare.132373198Figshare: Underlying data ofThis project contains the following underlying data:- Fig2A_090109_BxVEGF1.3k.xlsRepresentative cases of the analysis of upstream transcription factors using TFEL against specific promoters: (A) vascular endothelial growth factor (gene name: Vegfa).- Fig2B_081218_AxFAS0.4k.xlsRepresentative cases of the analysis of upstream transcription factors using TFEL against specific promoters: (B) fatty acid synthase (gene name: Fasn).- Fig2C_101026_B(10)xSREBP1c2.2k.xlsRepresentative cases of the analysis of upstream transcription factors using TFEL against specific promoters: (C) sterol regulatory element-binding protein-1c (gene name: Srebf1c).- Fig2D_081219_AxSREBP1c-T_1st.xlsRepresentative case identifying the combinatorial interactions on a specific promoter. In this assay, we screened for transcription factor(s) that suppress LXR activity on the SREBP-1c promoter (-249 to -144). 1st screening data.- Fig2D_081225_SREBP1c-T_2nd_A28.xlsRepresentative case identifying the combinatorial interactions on a specific promoter. In this assay, we screened for transcription factor(s) that suppress LXR activity on the SREBP-1c promoter (-249 to -144). 2nd screening data.Creative Commons Attribution 4.0 International license (CC-BY 4.0).Data are available under the terms of the I have no additional comments.Is the work clearly and accurately presented and does it cite the current literature?YesIf applicable, is the statistical analysis and its interpretation appropriate?NoAre all the source data underlying the results available to ensure full reproducibility?YesIs the study design appropriate and is the work technically sound?YesAre the conclusions drawn adequately supported by the results?YesAre sufficient details of methods and analysis provided to allow replication by others?YesReviewer Expertise:Regulation of gene expression, TF-mediated DNA demethylation, Cell reprogrammingI confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard. Identification of upstream TF(s) should be objective. Test cases shown in Figure 2A, B and C should be statistically evaluated. HIF-1a,2a and SREBP-1a,-2 may be evident, however I am not sure whether LXRa-mediated luciferase activity is significantly high or not.In figure 2D, authors selected 4 pools. I think this selection should also be objective by using statistical evaluation. If authors statistically selected pools for further one by one test, the pool with KLF4 has been selected?One of the most important value in this work must be TF expression plasmids that authors have systematically constructed. I think this resource should be available to scientific community.TFs regulate downstream genes by associating with promoters or enhancers. It is worth if the screen system works for TFs associating with enhancers.This paper describes a method to identify transcription factors (TFs) regulating target gene(s). The expression constructs that authors has made are highly useful for scientific community who are interested in TF-mediated gene regulation. The manuscript is written well, but there are several issues to be addressed.Is the work clearly and accurately presented and does it cite the current literature?YesIf applicable, is the statistical analysis and its interpretation appropriate?NoAre all the source data underlying the results available to ensure full reproducibility?YesIs the study design appropriate and is the work technically sound?YesAre the conclusions drawn adequately supported by the results?YesAre sufficient details of methods and analysis provided to allow replication by others?YesReviewer Expertise:Regulation of gene expression, TF-mediated DNA demethylation, Cell reprogrammingI confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard, however I have significant reservations, as outlined above. 1. Identification of upstream TF(s) should be objective. Test cases shown in Figure 2A, B and C should be statistically evaluated. HIF-1a,2a and SREBP-1a,-2 may be evident, however I am not sure whether LXRa-mediated luciferase activity is significantly high or not. We thank the reviewer for the valuable advice. Because the data have been obtained for the screening purpose, each pool has just single value, so robust statistical analyses to show significance are a little difficult to perform. However, as an empirical method, 2SD (standard deviation) criteria is often used. For Figure 2A (Vegfa-luc), only the pools #5 and #29  are above the 2SD line. For Figure 2B (Fasn-luc), the pools #30 and #60 (black bar) are above the 2SD line, but the pool #30 was revealed to contain no positive clones after the 2nd screening of each clone in the pool. For Figure 2C (Srebf1c-luc), the pools #5, #50, #76 (black bar) and #78 are above the 2SD line, but only the pool #76 was revealed to have a positive clone (LXRα) after the 2nd screening. Our actual strategy was to pick up the top 5%  for the 2nd screening. As a result, both approaches (the 2SD criterion and the top 5% criterion) are proved to work sufficiently as a screening method. Based on this discussion, we added the following paragraph to the revised manuscript: From the statistical point of view, we picked up the top (or the bottom for D) 5% pools for the 2nd screening, and as a result, this criterion was similar to the 2SD (standard deviation) criterion in these cases.2. In figure 2D, authors selected 4 pools. I think this selection should also be objective by using statistical evaluation. If authors statistically selected pools for further one by one test, the pool with KLF4 has been selected? Thank you for the question. We selected the top or bottom 5% pools for the 2nd screening, and only the pool #28 containing KLF4 was proved to be a true hit after the 2nd screening.3. One of the most important value in this work must be TF expression plasmids that authors have systematically constructed. I think this resource should be available to scientific community. Thank you for the comment. We are planning to talk with DNAFORM about how to share this expression library in the near future.4. TFs regulate downstream genes by associating with promoters or enhancers. It is worth if the screen system works for TFs associating with enhancers. We thank the reviewer for the valuable comment. Yes, we think that our method is also effective for enhancers to be matched with TFs, as shown in the case of TCF7L2 gene :422-433 (ref. 9)). According to the reviewer’s advice, we added the following sentence to the revised manuscript:TCF7L2 gene. It is noteworthy that the TFEL scan method is also applicable to enhancers as well as promoters as shown in the case of 1). How much of the mouse transcription factors in the expression library is overlapping with the human transcription factors. A brief discussion will be helpful.Human genome is estimated to encode for 1639 transcription factors . How much of the mouse transcription factors in the expression library is overlapping with the human transcription factors. A brief discussion will be helpful. We thank the reviewer for the valuable advice. According to this suggestion, we checked the overlapping among the transcription factor databases such as RIKEN TFdb :787-793 (ref.7)), Human TF :650-665.), and Animal TFDB 3.0 :D33-D38.), using a gene symbol converter available at http://biodb.jp/. The results were as follows: the overlap between RIKEN TFdb and Human TF was 889 genes, app 65% of Human TF and 53% of RIKEN TFdb, and the overlap between RIKEN TFdb and Animal TFDB 3.0 was 958 genes, app 59% of Animal TFDB 3.0 and 57% of RIKEN TFdb. The differences were a little larger than expected and even when we compared the same Animal TFDB human and mouse databases, the overlap was limited to 1,340 genes, which were app. 80% of the 1665 human TF genes in the database. Therefore, app. 300 TF genes (20%) seemed to be different between human and mouse. Thanks to the reviewer’s comment, we could better understand the differences among TF databases, but because this content is beyond the scope of this article, we will keep it within the Comments. Also, we realized during these series of gene symbol checking that KLF 9, 10, 11 had been included in the TFEL library, and corrected the description in the discussion.2. Many of the transcription factors characterized thus far are responsive to stresses or ligands. The results in this manuscript show that such transcription factors can be identified by overexpression. A brief discussion is needed. We thank the reviewer for the valuable comment. According to the reviewer’s advice, we added a new paragraph of discussion to the revised manuscript like this: Many of the transcription factors characterized thus far are responsive to stresses or ligands. The results of the case of LXR shown here demonstrate that such transcription factors can also be identified by overexpression. However, at the same time, we have experienced that some transcription factors activated in a ligand-dependent manner, for example, glucocorticoid receptor (GR), cannot be identified by mere overexpression without ligands (data not shown). Therefore, it may depend on transcription factors whether they can be detected without ligands/additional activators or not.3. How will this expression library be shared? Thank you for the comment. We are planning to talk with DNAFORM about how to share this expression library in the near future.4. The description in the text of experiment in Fig. 2D may be improved. Did authors try to identify factors that restrict/inhibit LXR? Why was this important for the regulation of Srebf1c promoter? Thank you for the question. Actually, there is a long story behind this. To make the long story short, we had found an important element shown to exert a suppressive effect on the SREBP-1c promoter in a fasting state :2373-2386 (ref. 6)). According to the reviewer’s advice, we added this explanation to the revised manuscript.


Transcriptional enhancers are major genomic elements that control gene activity in eukaryotes. Recent studies provided deeper insight into the temporal and spatial organization of transcription in the nucleus, the role of non-coding RNAs in the process, and the epigenetic control of gene expression. Thus, multiple molecular details of enhancer functioning were revealed. Here, we describe the recent data and models of molecular organization of enhancer-driven transcription. Gene transcription is precisely organized in time and space. The process requires the participation of hundreds of molecules, which form an extensive interaction network. Substantial progress was achieved recently in our understanding of the molecular processes that take place in the cell nucleus cis-regulatory DNA elements [Eukaryotic mRNA synthesis proceeds in short intense pulses, or bursts, which are periods of gene transcriptional activity followed by periods of gene silence. The intermittent pulsation of gene expression in eukaryotes is an intrinsic property of eukaryotic transcription and chromatin organization and is not caused by any external factor . Bursts elements . The aveelements . As a phenomenon, transcriptional bursting might represent a source of intrinsically random fluctuations in mRNA and protein levels in the cell. Bursts in mRNA level may be compensated for by decreases in the degradation levels of the respective proteins . An incrThe level of gene activity depends on the frequency of transcriptional bursts . Highly Transcriptional bursts coincide with promoter–enhancer chromatin contacts in tissues, indicating that enhancer–promoter chromatin looping acts as a regulatory mechanism . In earlin cis, the burst sizes, but not burst frequencies, of both genes were comparable. Furthermore, the co-transcription of both genes was statistically mutually inhibitory [in cis accounts for the alternation in burst frequency, emphasizing that changes in enhancer–promoter contacts do not alter the burst size, but affect only the burst frequency. The finding supports the model that the enhancer rapidly switches between promoters to allow co-transcriptional events to occur, suggesting that enhancer–promoter loops are formed and released with rapid kinetics. At the special level, such genes can be located in different transcription factories (see below) or be in the same factory. Definitive evidence for this model would require a direct imaging of enhancer–promoter contacts in live cells. In cells where two developmentally distinct LCR-regulated globin genes are co-transcribed hibitory . Thus, pHow does chromatin control the bursting events? At the very beginning of bursting research, re-expression was shown to be more likely to arise than de novo transcription in a cell population, indicating that epigenetic processes and transcription memory are involved in controlling gene expression . It was How may bursting be specially organized? It was noted that when two genes are integrated close together into the genome, they exhibit the same bursts , suggestThus, eukaryotic transcription occurs in short bursts. Enhancers regulate the total gene expression output predominately by modulating the bursting frequency.According to the current views of how transcription is organized in eukaryotes, the enhancers and promoters of co-expressed genes located in different chromosomal positions or even on different chromosomes cluster together during transcription to produce discrete nuclear foci, which contain the phosphorylated RNA polymerase II (Pol II) form and are known as transcription factories ,27,28,29Recent findings shed some light on why many transcription factors (TFs) and co-activators contain the so-called low-complexity domains (LCDs), which are regions with low amino acid diversity and are also known as the intrinsically disordered regions. LCDs are capable of forming liquid-like condensates, which are possible to disrupt using chemicals that perturb condensates, for example, with hexanediols . ProteinAs for enhancer-dependent activation, the above data suggest that activation must be associated with a phase switching, when one of the regulatory elements, for example, the gene promoter, is recruited from one—the liquid phase—to another, the protein phase, where the enhancer resides. Once the elements are in the same phase enriched in TFs, Mediator, Pol II, etc., their higher concentrations may facilitate the Linking/Chaining/Looping between the regulatory elements. A physical contact between the enhancer and promoter is not essential for transcriptional activation; it is enough that they are brought to a distance shorter than 330 nm of each other . On the The above data clarify, to an extent, the nature of enhancer function and the organization of eukaryotic transcription. Yet the question is still open as to whether Pol II moves along the gene during transcription or the gene moves relative to Pol II immobilized in a dynamically forming factory. Studies with 3C methods and super-resolution microscopy showed that the distance between two transcripts of two co-expressed genes co-localized in one transcription factory is lying within 35 nm of the surface of an 87-nm sphere. Given the known gene sizes, the finding is only possible to explain by assuming that two responding genes diffuse to an 87-nm factory to be transcribed by immobilized enzymes and that when two genes are reeled in to the transcription factory, only parts being transcribed at a given moment lie transiently together . The samRecently, evidence was provided in favor of an alternative point of view. Using fluorescence in situ hybridization (FISH) on very long genes, it was demonstrated that such genes form transcriptional loops and that the bases of these loops are not brought together. It was found that loops extend into the nuclear interior where euchromatin is situated and are decorated with ribonucleoprotein complexes containing moving Pol II  Figure .Thus, different scenarios are possible for different genes in different conditions. The enhancer–polymerase complex may move along a chromatin fiber during transcription elongation or, oppositely, it may lie within a transcription factory while the gene body is drawn through the factory and looped out to gradually move away. The mechanisms of enhancer-dependent activation of gene transcription are tightly associated with how Pol II works on the given gene. Two basic models currently explain the activation of gene transcription in higher eukaryotes. One model assumes that Pol II is absent from the promoter and coding region when a gene is repressed (inactive). An activation signal—for example, a hormonal or stress stimulus—serves to recruit a transcriptional activator to the promoter and/or enhancer of the gene. The activator stimulates the recruitment of Pol II and assembly of the Pol II complex. Once bound to the promoter, Pol II immediately starts elongation on the coding region and departs after having completed transcription until the next recruitment round. Recruitment of the Pol II complex is a limiting step of the process in this case. The other mechanism is assumed for the genes where the Pol II complex is efficiently recruited to the promoter even when the gene is inactive. The complex pre-assembled on the promoter of an inactive gene undergoes all of the transcription initiation steps. The C-terminal domain of Pol II is phosphorylated at Ser5 by Cdk7 kinase, and the complex leaves the promoter to start RNA synthesis. After adding 40–60 nucleotides to RNA, the Pol II complex is stalled by the negative elongation factor  and DRB sensitivity inducing factor  and waits for a signal to start productive transcription. An activator recruited to the promoter facilitates the transition of the promoter-associated complex into the elongating state. Elongation kinase positive transcription elongation factor  phosphorylates the subunits NELF-A and NELF-E of NELF and the SPT5 subunit of DSIF, as well as Ser2 in C-terminal domain of Pol II ,45. NELFDrosophila [Thus, two types of promoters are recognized: the promoters that are associated with paused Pol II and are regulated by stalling the Pol II complex  and the stochastic promoters, which are characterized by Pol II complex turnover. A substantial proportion of active genes was found to have stalled Pol II on the promoter in osophila . The mecosophila . The proosophila . The enhDrosophila showed that 94% of the enhancers have stable contacts with promoters before expression is activated during mesoderm development and that Pol II occurs in the paused state on the respective genes [In accordance with the above mechanisms, two basic models were assumed for the mechanism of action of enhancers. One is a permissive model and suggests that enhancer–promoter contacts pre-exist before gene activation  and are only slightly reorganized in the course of cell differentiation (specification), as was demonstrated for TNF-alpha-, thrombopoietin-, and p53-dependent genes ,51,52. Ive genes . Thus, pve genes . Pre-exiShh promoter. The enhancer was spatially closer to the promoter in the limb bud part where Shh was active, and the distance between the elements was < 200 nm [Microscopic studies identified the threshold above which the bringing of a promoter and an enhancer together is not accompanied by gene activation. For example, the transcription of a reporter gene did not start when the gene–enhancer distance was reduced from 700 to 400 nm, but started when the distance was 330 nm in an insulator-pairing assay . Similar< 200 nm .The other model is instructive and suggests that contacts form de novo in response to a stimulus or upon cell differentiation and that their formation is accompanied by simultaneous changes in gene expression ,62,63,64Drosophila. Both of the two scenarios are simultaneously utilized during cell differentiation, which is accompanied by loss of pre-existing contacts, changes in chromatin state, and the acquisition of new contacts. A reorganization of contacts is a highly dynamic process that may take place within a few hours ,65,66. TAs for the direct activation triggers, the role may be played not only by the distance between the enhancer and promoter, but also by their minor dynamic movements that occur within the preformed topology and are accompanied by the establishment of a limited number of new contacts ,67. The In sum, gene regulatory elements are highly mobile in the nuclear space. Molecular forces and specific chromatin proteins organize their movement, the formation of contacts of the elements, the assembly of transcription factories, as well as dissociation of the assemblies . HoweverDrosophila developmental enhancer transcription strongly correlates with the timing and location of enhancer activity [The discovery of mammalian enhancer transcription added another layer of complexity to our deepening understanding of the nature of these elements. New insights continue to upend long-held, common assumptions about enhancers, forcing us to re-examine the mechanisms of their action. The advent of the genomic era taught that the transcription of enhancers is a universal phenomenon characteristic of Metazoa ,72,73,74activity . Yet transcription is not found at all active enhancers. This suggests that eRNA is not required for the manifestation of enhancer activity, at least for this subset of active enhancers. Andersson et al. found that 20% to 33% of non-transcribed regulatory regions can activate transcription in an in vitro enhancer assay, suggesting that they may also function as enhancers without eRNA transcription ,84,85.As promoters of protein-coding genes, enhancers expressing eRNA are enriched in transcription initiation complex, CBP/p300, and other TFs, and serine 5 phosphorylated RNA Pol II. Yet, unlike gene promoters, enhancers are depleted of serine 2 phosphorylated Pol II . PromoteDrosophila that IncRNA transcribed from the extragenic enhancers in the Bithorax complex (BX-C) can transcriptionally interfere with gene transcription [A number of mammalian intragenic enhancers are transcribed to yield tissue-specific, long non-coding RNAs (lncRNA) in the direction of gene transcription . Unlike cription . These dcription ,86. Howecription . Active cription . ConsequDrosophila embryogenesis was assessed in vivo in transgenic embryos [Drosophila suggests that there is also a correlation between enhancer strength and endogenous Cap-analysis gene expression (CAGE) levels [The directionality of transcription appears to play a role in the ability to function as an enhancer or promoter in vivo. The relationship between eRNA transcription and enhancer activity during  embryos . Mikhayl embryos . The tes embryos . Evidenc) levels ,99. The cis regulation of distal genes in their natural context. A systematic analysis of the promoters of human coding genes revealed that up to 3% of promoters have enhancer activity and are involved in the regulation of distal genes through establishing long-range contacts [In recent years, massively parallel reporter assay (MPRA) and self-transcribing active regulatory region sequencing (STARR-seq) enabled quantitative and straightforward measurements of enhancer activity in mammals ,100. It contacts ,102. Notcontacts ,104,105.Taken together, these data indicate that transcriptional regulators have a wide range of activities. There are elements that function strictly as enhancers or promoters, while others work as enhancers with weak promoter activity, while some elements show both strong enhancer and promoter activity. This spectrum of activities tightly correlates with the directionality of their transcription, which likely reflects the underlying sequence properties of the element . The aboEnhancer transcription proceeds through the same steps as gene promoter transcripts, including TSS-proximal pausing, capping, and release ,107,108.The functional role of eRNA remains largely unknown. We do know that the production of eRNAs is clearly associated with enhancer activity, which is observed in well-characterized enhancers (α-globin and β-globin loci) and globally in putative enhancers upon stimulation  ,77,79,81How eRNAs contribute to transcriptional activation remains a subject of many ongoing investigations. There are three hypotheses regarding the role of enhancer transcription. The first states that the process of transcription, but not eRNA itself, is important for activating the capacity of enhancers. The second posits that eRNA transcripts are necessary for enhancer activity. The last suggests that eRNA can represent a transcriptional noise due to the promiscuous recruitment of Pol II to open chromatin . The thrAdditionally, RNA-tethering experiments in reporter assays demonstrated a quantitative effect of eRNA transcripts in conferring gene activation, independent of the act of transcription ,119,127.Experiments to discern the role of enhancer transcription or eRNAs are difficult to design. This is because alterations of the signals required for transcription may have negative, inadvertent effects on other aspects of enhancer activity. Recently, a different approach was used to manipulate eRNA transcription . In thisin cis, although there is growing evidence that they can exert the regulatory function in trans [CEeRNA) regulates transcription of the adjacent MyoD gene in cis, whereas the distal regulatory region (DRR) enhancer transcript (DRReRNA) promotes Myogenin transcription in trans [DRReRNA mediates loop formation through cohesin recruitment. A DRReRNA knockdown leads to reduced cohesin occupancy at the Myogenin locus, and this reduction downregulates Myogenin, ultimately preventing muscle cell differentiation. Furthermore, it was speculated that the process of cohesin recruitment mechanistically differs from estrogen receptor-regulated transcription, where the cohesin complex is present on many estrogen receptor-regulated enhancers even before ligand treatment and contributes to gene activation, at least in part, by stabilizing E2/ER-alpha/eRNA-induced enhancer–promoter looping [DRReRNA, ncRNA-a7, KLK3eRNA, and Bloodlinc are polyadenylated [Most eRNAs function in trans ,140,146.in trans . The corin trans ,148,149. looping ,140. As enylated ,140,146.A subset of other eRNAs was also found to interact with cohesin, which was proposed to facilitate communication between an enhancer and its target promoter by stabilizing looping . HoweverThe TF Yin Yang 1 (YY1) was assumed to mediate enhancer loops. Unlike with cohesin, YY1 binding is strongly enriched at promoters and enhancers in mouse and human cell types ,155. IntAll the aforementioned studies report average expression profiles, which have a limited potential to accurately describe what occurs in individual cells and at individual alleles. Fluorescence in situ hybridization (FISH) studies to measure eRNA and mRNA co-expression at individual alleles revealed that co-expressing alleles are rare, and co-expression rarely occurs in a closed enhancer–promoter configuration ,85,156. As presented in this chapter, the transcription of eRNAs may be more important for gene activation than was realized, with up to 50% of all enhancers depending on it for functioning . NumerouAs for the subset of enhancers that do not require eRNA transcription, nature has solved the problem of enhancer–promoter communication in a different way; exactly how is yet to be discovered. Some enhancers evolved to rely on transcription for functioning. For this other subset of enhancers, both the transcription process and some eRNAs by themselves are proven to have functional roles. More evidence is required to fully decipher the biological roles. To understand how eRNAs and TFs act—either jointly or independently—to stimulate transcription requires the targeted deletion of distinct enhancer regions to change the eRNA structure or abolish eRNA expression, while maintaining the binding of TFs. On the other hand, information on the dynamics of enhancer–promoter interactions in the presence or absence of eRNAs is still lacking and will require analyses of living cells at high spatial and temporal resolution. One of the first epigenetic characteristics of enhancers is their depletion of nucleosomes as detectable by nuclease treatment. A significant correlation was revealed between cell-specific DNase I hypersensitive loci and enhancer elements . In addiThe recruitment of multiple TFs is another characteristic feature of enhancers. Two modes of their binding to chromatin were proposed. The first is that the process is positively cooperative, which helps TFs overcome the nucleosomal barrier. The second is that there are certain pioneer factors that bind to nucleosomal DNA, thereby enabling other TFs to bind to the enhancer. The latter occurs via a direct effect on the nucleosome position, the recruitment of chromatin remodelers and modifiers, and the protection of DNA from methylation ,168,169.The nucleosomes that flank the TF binding sites in enhancers carry the specific histone marks H3K4me1 and H3K27ac ,177,178.The exact function of the H3K4me1 mark is still unclear. H3K4 methylation seems to affects the recruitment of some co-activators and co-repressors. There are several hypothetical players in this process. Methylated H3K4 can prevent the recruitment of some co-repressors, including DNA methyltransferase Dnmt3 . Thus, HDrosophila and MLL3/4 in mammals [The methylation of H3K4 is accomplished by methyltransferases Trr1 in  mammals ,186,187. mammals . The dep mammals ,186. H3K mammals . Finally mammals . At the same time, a broader range of methyltransferases may probably be involved in the formation of this modification. The mechanism of their recruitment is still unknown. It is likely that pioneer TFs are responsible for the recruitment to at least a certain group of enhancers . MoreoveDrosophila [During enhancer disengagement, the methylation mark is erased by LSD1 demethylase in ESCs . Howeverosophila . Anotherosophila .In addition to H3K4me1, enhancers could carry H3K4me3 modification ,98. ThisHistone acetylation is a widespread modification of active chromatin. For the enhancer, there is an interplay of methylation and acetylation modifications: the characteristic H3K27ac mark is established on the enhancers that already carry H3K4me1 . DiffereHowever, not all enhancers carry a p300/CBP marker, and some other HATs appear to bind to them. For example, Gcn5/PCAF HAT and its preferential H3K9ac mark were detected on enhancers ,195,196.Acetylated histones are recognized by bromodomains, which are found in various proteins . The acetylated histone–bromodomain interaction can stabilize the protein assemblies on enhancers. Acetylation can also destabilize nucleosomes and contribute to the formation of histone-free chromatin regions . MoreoveDrosophila, H3K27me3 histone demethylase UTX directly binds CBP/p300, which helps to change H3K27me3 into H3K27ac and activate poised enhancers [There is another class of enhancers, referred to as poised enhancers, which were described in ESCs ,178. Sucnhancers .It is as yet unclear whether the H3K4me1 or H3K27ac modifications are sufficient or necessary for enhancer activity. Enhancers show heterogeneous histone modifications and, in addition to the aforementioned marks, they may carry other ones, such as H3K9ac and H3K18ac ,202, H3KFinally, the epigenetic mark of DNA methylation was found on inactive enhancers ,210,211.In sum, enhancers are dynamic entities that carry different epigenetics marks at different developmental stages. Epigenetic marks can participate in the assembly of transcription complexes, open access for TF binding, and prime regulatory elements . HoweverRecent advances in the techniques available to study the molecular processes in the nucleus contributed a lot into our understanding of enhancer action. At the same time, novel models brought novel issues, and several of them should be mentioned. The precise molecular mechanism that various enhancers use to control the transcription burst duration and frequency is still unknown. The molecular forces that drive the formation of sub-nuclear bodies, including transcription factories, are under intense study now. The contributions of different classes of chromatin-associated molecules, including cohesins and eRNAs, to the formation of enhancer–promoter contacts need thorough investigation. Different molecular landscapes and potentially different modes of action of different classes of enhancers and promoters require substantial attention. Finally, the arising concept of a mixing of different types of activities, such as enhancer, promoter, and insulator activities, in one regulatory element is attracting a lot of interest.