Gene network transitions in embryos and other fate-changing contexts involve combinations of transcription factors. A subset of fate-changing transcription factors act as pioneers; they scan and target nucleosomal DNA and initiate cooperative events that can open the local chromatin. However, a gap has remained in understanding how molecular interactions with the nucleosome contribute to the chromatin-opening phenomenon. Here we identified a short α-helical region, conserved among FOXA pioneer factors, that interacts with core histones and contributes to chromatin opening in vitro. The same domain is involved in chromatin opening in early mouse embryos for normal development. Thus, local opening of chromatin by interactions between pioneer factors and core histones promotes genetic programming.
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Genomic data have been deposited in the Gene Expression Omnibus database under accession number GSE134465.
Kundaje, A. et al. Integrative analysis of 111 reference human epigenomes. Nature 518, 317–330 (2015).
Pérez-Lluch, S. et al. Absence of canonical marks of active chromatin in developmentally regulated genes. Nat. Genet. 47, 1158–1167 (2015).
Ho, J. W. K. et al. Comparative analysis of metazoan chromatin organization. Nature 512, 449–452 (2014).
Ernst, J. et al. Mapping and analysis of chromatin state dynamics in nine human cell types. Nature 473, 43–49 (2011).
Gaffney, D. J. et al. Controls of nucleosome positioning in the human genome. PLoS Genet. 8, e1003036 (2012).
Lidor Nili, E. et al. p53 binds preferentially to genomic regions with high DNA-encoded nucleosome occupancy. Genome Res. 20, 1361–1368 (2010).
Tillo, D. et al. High nucleosome occupancy is encoded at human regulatory sequences. PLoS ONE 5, e9129 (2010).
Cirillo, L. et al. Opening of compacted chromatin by early developmental transcription factors HNF3 (FOXA) and GATA-4. Mol. Cell 9, 279–289 (2002).
Iwafuchi-Doi, M. & Zaret, K. S. Pioneer transcription factors in cell reprogramming. Genes Dev. 28, 2679–2692 (2014).
Soufi, A. et al. Pioneer transcription factors target partial DNA motifs on nucleosomes to initiate reprogramming. Cell 161, 555–568 (2015).
Mayran, A. et al. Pioneer factor Pax7 deploys a stable enhancer repertoire for specification of cell fate. Nat. Genet. 50, 259–269 (2018).
Carroll, J. S. et al. Chromosome-wide mapping of estrogen receptor binding reveals long-range regulation requiring the forkhead protein FoxA1. Cell 122, 33–43 (2005).
Golson, M. L. & Kaestner, K. H. Fox transcription factors: from development to disease. Development 143, 4558–4570 (2016).
Ang, S. L. et al. The formation and maintenance of the definitive endoderm lineage in the mouse: involvement of HNF3/forkhead proteins. Development 119, 1301–1315 (1993).
Sasaki, H. & Hogan, B. L. Differential expression of multiple fork head related genes during gastrulation and pattern formation in the mouse embryo. Development 118, 47–59 (1993).
Monaghan, A. P., Kaestner, K. H., Grau, E. & Schütz, G. Postimplantation expression patterns indicate a role for the mouse forkhead/HNF-3 α, β and γ genes in determination of the definitive endoderm, chordamesoderm and neuroectoderm. Development 119, 567–578 (1993).
Ang, S.-L. & Rossant, J. HNF-3β is essential for node and notochord formation in mouse development. Cell 78, 561–574 (1994).
Weinstein, D. C. et al. The winged-helix transcription factor HNF-3β is required for notochord development in the mouse embryo. Cell 78, 575–588 (1994).
Hsu, H.-T. et al. Recruitment of RNA polymerase II by the pioneer transcription factor PHA-4. Science 348, 1372–1376 (2015).
Mango, S. E., Lambie, E. J. & Kimble, J. The pha-4 gene is required to generate the pharyngeal primordium of Caenorhabditis elegans. Development 120, 3019–3031 (1994).
Cirillo, L. A. et al. Binding of the winged-helix transcription factor HNF3 to a linker histone site on the nucleosome. EMBO J. 17, 244–254 (1998).
Sekiya, T. & Zaret, K. S. Repression by Groucho/TLE/Grg proteins: genomic site recruitment generates compacted chromatin in vitro and impairs activator binding in vivo. Mol. Cell 28, 291–303 (2007).
Simpson, R. T., Thoma, F. & Brubaker, J. M. Chromatin reconstituted from tandemly repeated cloned DNA fragments and core histones: a model system for study of higher order structure. Cell 42, 799–808 (1985).
McPherson, C. E., Shim, E. Y., Friedman, D. S. & Zaret, K. S. An active tissue-specific enhancer and bound transcription factors existing in a precisely positioned nucleosomal array. Cell 75, 387–398 (1993).
Chaya, D., Hayamizu, T., Bustin, M. & Zaret, K. S. Transcription factor FoxA (HNF3) on a nucleosome at an enhancer complex in liver chromatin. J. Biol. Chem. 276, 44385–44389 (2001).
Fletcher, T. M. & Hansen, J. C. The nucleosomal array: structure/function relationships. Crit. Rev. Eukaryot. Gene Expr. 6, 149–188 (1996).
Hill, D. A. & Imbalzano, A. N. Human SWI/SNF nucleosome remodeling activity is partially inhibited by linker histone H1. Biochemistry 39, 11649–11656 (2000).
Horn, P. J. et al. Phosphorylation of linker histones regulates ATP-dependent chromatin remodeling enzymes. Nat. Struct. Biol. 9, 263–267 (2002).
Ramachandran, A., Omar, M., Cheslock, P. & Schnitzler, G. R. Linker histone H1 modulates nucleosome remodeling by human SWI/SNF. J. Biol. Chem. 278, 48590–48601 (2003).
Lai, E. et al. HNF-3A, a hepatocyte-enriched transcription factor of novel structure is regulated transcriptionally. Genes Dev. 4, 1427–1436 (1990).
Qian, X. & Costa, R. H. Analysis of hepatocyte nuclear factor-3 beta protein domains required for transcriptional activation and nuclear targeting. Nucleic Acids Res. 23, 1184–1191 (1995).
Lai, E., Clark, K. L., Burley, S. K. & Darnell, J. E. Jr. Hepatocyte nuclear factor 3/fork head or “winged helix” proteins: a family of transcription factors of diverse biologic function. Proc. Natl Acad. Sci. USA 90, 10421–10423 (1993).
Pani, L. et al. Hepatocyte nuclear factor 3β contains two transcriptional activation domains, one of which is novel and conserved with the Drosophila fork head protein. Mol. Cell. Biol. 12, 3723–3732 (1992).
Barozzi, I. et al. Coregulation of transcription factor binding and nucleosome occupancy through DNA features of mammalian enhancers. Mol. Cell 54, 844–857 (2014).
Ballaré, C. et al. Nucleosome-driven transcription factor binding and gene regulation. Mol. Cell 49, 67–79 (2013).
Iwafuchi-Doi, M. et al. The pioneer transcription factor FoxA maintains an accessible nucleosome configuration at enhancers for tissue-specific gene activation. Mol. Cell 62, 79–91 (2016).
Clark, K. L., Halay, E. D., Lai, E. & Burley, S. K. Co-crystal structure of the HNF3/fork head DNA recognition motif resembles histone H5. Nature 364, 412–420 (1993).
Zaret, K. Early liver differentiation: genetic potentiation and multilevel growth control. Curr. Opin. Genet. Dev. 8, 526–531 (1998).
Chen, S. X. et al. Quantification of factors influencing fluorescent protein expression using RMCE to generate an allelic series in the ROSA26 locus in mice. Dis. Model. Mech. 4, 537–547 (2011).
Arai, R., Wriggers, W., Nishikawa, Y., Nagamune, T. & Fujisawa, T. Conformations of variably linked chimeric proteins evaluated by synchrotron X-ray small-angle scattering. Proteins 57, 829–838 (2004).
Yoshiba, S. & Hamada, H. Roles of cilia, fluid flow, and Ca2+ signaling in breaking of left–right symmetry. Trends Genet. 30, 10–17 (2014).
Kanai-Azuma, M. et al. Depletion of definitive gut endoderm in Sox17-null mutant mice. Development 129, 2367–2379 (2002).
Wat, M. J. et al. Mouse model reveals the role of SOX7 in the development of congenital diaphragmatic hernia associated with recurrent deletions of 8p23.1. Hum. Mol. Genet. 21, 4115–4125 (2012).
Shindo, T. et al. Kruppel-like zinc-finger transcription factor KLF5/BTEB2 is a target for angiotensin II signaling and an essential regulator of cardiovascular remodeling. Nat. Med. 8, 856–863 (2002).
Ema, M. et al. Kruppel-like factor 5 is essential for blastocyst development and the normal self-renewal of mouse ESCs. Cell Stem Cell 3, 555–567 (2008).
Kuo, C. T. et al. GATA4 transcription factor is required for ventral morphogenesis and heart tube formation. Genes Dev. 11, 1048–1060 (1997).
Molkentin, J. D., Lin, Q., Duncan, S. A. & Olson, E. N. Requirement of the transcription factor GATA4 for heart tube formation and ventral morphogenesis. Genes Dev. 11, 1061–1072 (1997).
Forlani, S., Lawson, K. A. & Deschamps, J. Acquisition of Hox codes during gastrulation and axial elongation in the mouse embryo. Development 130, 3807–3819 (2003).
Candia, A. F. et al. Mox-1 and Mox-2 define a novel homeobox gene subfamily and are differentially expressed during early mesodermal patterning in mouse embryos. Development 116, 1123–1136 (1992).
Bettenhausen, B., Hrabĕ de Angelis, M., Simon, D., Guénet, J. L. & Gossler, A. Transient and restricted expression during mouse embryogenesis of Dll1, a murine gene closely related to Drosophila Delta. Development 121, 2407–2418 (1995).
Hochgreb, T. et al. A caudorostral wave of RALDH2 conveys anteroposterior information to the cardiac field. Development 130, 5363–5374 (2003).
Abu-Abed, S. et al. The retinoic acid-metabolizing enzyme, CYP26A1, is essential for normal hindbrain patterning, vertebral identity, and development of posterior structures. Genes Dev. 15, 226–240 (2001).
Sakai, Y. et al. The retinoic acid-inactivating enzyme CYP26 is essential for establishing an uneven distribution of retinoic acid along the anterio-posterior axis within the mouse embryo. Genes Dev. 15, 213–225 (2001).
Sanford, L. P. et al. TGFβ2 knockout mice have multiple developmental defects that are non-overlapping with other TGFβ knockout phenotypes. Development 124, 2659–2670 (1997).
Buenrostro, J. D., Wu, B., Chang, H. Y. & Greenleaf, W. J. ATAC-seq: a method for assaying chromatin accessibility genome-wide. Curr. Protoc. Mol. Biol. 109, 21.29.1–21.29.9 (2015).
Li, Z., Schug, J., Tuteja, G., White, P. & Kaestner, K. H. The nucleosome map of the mammalian liver. Nat. Struct. Mol. Biol. 18, 742–746 (2011).
Koumangoye, R. B. et al. SOX4 interacts with EZH2 and HDAC3 to suppress microRNA-31 in invasive esophageal cancer cells. Mol. Cancer 14, 24 (2015).
Lee, H., Habas, R. & Abate-Shen, C. MSX1 cooperates with histone H1b for inhibition of transcription and myogenesis. Science 304, 1675–1678 (2004).
Wang, J. et al. The Msx1 homeoprotein recruits polycomb to the nuclear periphery during development. Dev. Cell 21, 575–588 (2011).
Rave-Harel, N., Miller, N. L. G., Givens, M. L. & Mellon, P. L. The Groucho-related gene family regulates the gonadotropin-releasing hormone gene through interaction with the homeodomain proteins MSX1 and OCT1. J. Biol. Chem. 280, 30975–30983 (2005).
Watts, J. A. et al. Study of FoxA pioneer factor at silent genes reveals Rfx-repressed enhancer at Cdx2 and a potential indicator of esophageal adenocarcinoma development. PLoS Genet. 7, e1002277 (2011).
Sandoval, G. J. et al. Binding of TMPRSS2-ERG to BAF chromatin remodeling complexes mediates prostate oncogenesis. Mol. Cell 71, 554–566.e7 (2018).
Barisic, D., Stadler, M. B., Iurlaro, M. & Schübeler, D. Mammalian ISWI and SWI/SNF selectively mediate binding of distinct transcription factors. Nature 569, 136–140 (2019).
Hoffman, J. A. & Trotter, K. W. & Ward, J. M. & Archer, T. K. BRG1 governs glucocorticoid receptor interactions with chromatin and pioneer factors across the genome. eLife 7, e35073 (2018).
Liu, J. K., DiPersio, C. M. & Zaret, K. S. Extracellular signals that regulate liver transcription factors during hepatic differentiation in vitro. Mol. Cell. Biol. 11, 773–784 (1991).
Jackson, D. A. et al. Modulation of liver-specific transcription by interactions between hepatocyte nuclear factor 3 and nuclear factor 1 binding DNA in close apposition. Mol. Cell. Biol. 13, 2401–2410 (1993).
Meers, M. P. & Janssens, D. H. & Henikoff, S. Pioneer factor-nucleosome binding events during differentiation are motif encoded. Mol. Cell 75, 562–575.e5 (2019).
Gualdi, R. et al. Hepatic specification of the gut endoderm in vitro: cell signaling and transcriptional control. Genes Dev. 10, 1670–1682 (1996).
Bossard, P. & Zaret, K. S. GATA transcription factors as potentiators of gut endoderm differentiation. Development 125, 4909–4917 (1998).
Cirillo, L. A. & Zaret, K. S. An early developmental transcription factor complex that is more stable on nucleosome core particles than on free DNA. Mol. Cell 4, 961–969 (1999).
Boller, S. et al. Pioneering activity of the C-terminal domain of EBF1 shapes the chromatin landscape for B cell programming. Immunity 44, 527–541 (2016).
Li, R. et al. Dynamic EBF1 occupancy directs sequential epigenetic and transcriptional events in B-cell programming. Genes Dev. 32, 96–111 (2018).
van Oevelen, C. et al. C/EBPα activates pre-existing and de novo macrophage enhancers during induced pre-B cell transdifferentiation and myelopoiesis. Stem Cell Reports 5, 232–247 (2015).
Sardina, J. L. et al. Transcription factors drive Tet2-mediated enhancer demethylation to reprogram cell fate. Cell Stem Cell 23, 905–906 (2018).
Johnson, J. L. et al. Lineage-determining transcription factor TCF-1 initiates the epigenetic identity of T cells. Immunity 48, 243–257.e10 (2018).
Imbeault, M., Helleboid, P. -Y. & Trono, D. KRAB zinc-finger proteins contribute to the evolution of gene regulatory networks. Nature 543, 550–554 (2017).
Sai, L. & Zheng, E. B. & Zhao, L. & Liu, S. Nonreciprocal and conditional cooperativity directs the pioneer activity of pluripotency transcription factors. Cell Rep. 28, 2689–2703.e4 (2019).
Yan, C., Chen, H. & Bai, L. Systematic study of nucleosome-displacing factors in budding yeast. Mol. Cell 71, 294–305.e4 (2018).
Henikoff, S. & Ramachandran, S. Pioneers invade the nucleosomal landscape. Mol. Cell 71, 193–194 (2018).
Brahma, S. & Henikoff, S. RSC-associated subnucleosomes define MNase-sensitive promoters in yeast. Mol. Cell 73, 238–249.e3 (2019).
Sekiya, T., Muthurajan, U. M., Luger, K., Tulin, A. V. & Zaret, K. S. Nucleosome-binding affinity as a primary determinant of the nuclear mobility of the pioneer transcription factor FoxA. Genes Dev. 23, 804–809 (2009).
Donaghey, J. et al. Genetic determinants and epigenetic effects of pioneer-factor occupancy. Nat. Genet. 50, 250–258 (2018).
Hurtado, A., Holmes, K. A., Ross-Innes, C. S., Schmidt, D. & Carroll, J. S. FOXA1 is a key determinant of estrogen receptor function and endocrine response. Nat. Genet. 43, 27–33 (2011).
Jozwik, K. M. & Carroll, J. S. Pioneer factors in hormone-dependent cancers. Nat. Rev. Cancer 12, 381–385 (2012).
Lupien, M. et al. FoxA1 translates epigenetic signatures into enhancer-driven lineage-specific transcription. Cell 132, 958–970 (2008).
Zhu, F. et al. The interaction landscape between transcription factors and the nucleosome. Nature 562, 76–81 (2018).
Fernandez Garcia, M. et al. Structural features of transcription factors associating with nucleosome binding. Mol. Cell 75, 921–932.e6 (2019).
Scholtz, J. M. & Baldwin, R. L. The mechanism of alpha-helix formation by peptides. Annu. Rev. Biophys. Biomol. Struct. 21, 95–118 (1992).
Zaret, K. S. & Stevens, K. Expression of a highly unstable and insoluble transcription factor in Escherichia coli: purification and characterization of the fork head homolog HNF3 alpha. Protein Expr. Purif. 6, 821–825 (1995).
Cuesta, I., Zaret, K. S. & Santisteban, P. The forkhead factor FoxE1 binds to the thyroperoxidase promoter during thyroid cell differentiation and modifies compacted chromatin structure. Mol. Cell. Biol. 27, 7302–7314 (2007).
Merzlyak, E. M. et al. Bright monomeric red fluorescent protein with an extended fluorescence lifetime. Nat. Methods 4, 555–557 (2007).
Picelli, S. et al. Full-length RNA-seq from single cells using Smart-seq2. Nat. Protoc. 9, 171–181 (2014).
Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
Love, M. I., Anders, S., Kim, V. & Huber, W. RNA-Seq workflow: gene-level exploratory analysis and differential expression. F1000Res. 4, 1070 (2015).
Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014).
Huang, D. W., Sherman, B. T. & Lempicki, R. A. Bioinformatics enrichment tools: paths toward the comprehensive functional analysis of large gene lists. Nucleic Acids Res. 37, 1–13 (2009).
Zhang, Y. et al. Model-based analysis of ChIP-Seq (MACS). Genome Biol. 9, R137 (2008).
Heinz, S. et al. Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities. Mol. Cell 38, 576–589 (2010).
We thank R. Dunbrack (Fox Chase Cancer Center) for the secondary structure analysis, C. Ducker for the crosslinking studies, S. Hipkens for the mouse embryonic stem cell culture and gene targeting in the Transgenic Mouse/Embryonic Stem Cell Shared Resource, R. McCarthy for data review and the University of Pennsylvania Flow Cytometry and Cell Sorting Facility. M.I. was supported by postdoctoral fellowships from Japan Society for the Promotion of Science Foundation (H26-683), Naito Foundation (RYU10000032), Astellas Foundation for Research on Metabolic Disorders (K0076) and Uehara Memorial Foundation (H24-20124). P.S. was supported by grant nos. SAF2016-75531-R (MICINN/FEDER, UE) and B2017/BMD-3724 (Comunidad de Madrid). The research was supported by NIH grant no. GM36477 to K.S.Z.
The authors declare no competing interests.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
a, Schematic of crosslinking of histone octamers used as input and FOXA1. SDS-PAGE analysis of FOXA1 or FOXA1 crosslinked to core histones, stained with Coomassie blue. Crosslinked products of a mobility expected for FOXA1 and core histone together are noted as band A and band B. The full blot gel is presented in the Source Data files. b, Underlined sequences are identified by peptide mass matching while the subset highlighted by green are used for relative peptide quantification in Extended Data Fig. 2. c, Strategy to map candidate interaction sites, explaining how crosslinked peptides gain a much greater mass and become depleted from the m/z spectrum. Source data
Extended Data Fig. 2 Mass spectrometry identification of FOXA1 peptides depleted by crosslinking to core histones.
Relative quantification of FOXA1 peptides in crosslinked bands A and B shown in Extended Data Fig. 1a. The integrated intensities of ions corresponding to peptides YPHAKPPYSYISLITMAIQQAPSK and ASQLEGAPAPGPAASPQTLDHSGATATGGASELK were used for normalizing tabulated intensities of other FOXA1 peptide ions. Those peptide intensities found unaltered in bands A and B (Extended Data Fig. 1a) are shown in the blue panel while those whose intensities changed in the FOXA1:Histone x-linked bands are shown in the red panel, and noted by red arrows, as discussed in the main text. Lower right, quantitation of peptide signals of aa415–443 over control peptide signals of aa313–246 within the same respective spectrum, demonstrating a diminution of aa415–442 in band A due to blockage at K414.
a, A putative α-helical region is conserved in FOXA1 and FOXA2 homologs. Conserved regions II and III are highlighted in teal and the α-helical region around K414 (orange) is highlighted in green. b, FOXA2 wild-type (WT) protein and delta-helix (ΔHx) mutant bound to Sepharose beads in a pulldown assay to assess binding to histone octamers. The bar graph represents mean ± s.e.m. of four replicates. P values are from a one-sided paired t-test, comparing the ratio of the core histones H3/H2A/H2B as a group and H4 to the recovered amount of full length FOXA2 in the same lane (n = 4 experiments; *, P < 0.05 different from WT). Dashed lines indicate partial FOXA2 degradation products from the C-terminus of the 6×-his tagged FOXA2 proteins, which were tagged on the N-terminus. The full gel is presented in the Source Data files. Source data
Extended Data Fig. 4 Deficiency in activation of endogenous FOXA1 liver target genes by FOXA- ΔHx and FOXA1-PP mutant proteins.
a, Schematic of functional assay for FOXA1 wild-type and mutants in H2.35 liver cells. b, Apoa1 and Ttr1 expression analysis by RT-qPCR relative to expression levels in the control. Results shown as mean ± s.e.m. of four biological replicates. P values are from two-sided Student’s t-test. c, Western blot analysis of two biological replicates for endogenous and exogenous FOXA proteins, demonstrating similar amounts of FOXA-ΔHx and FOXA1-PP mutant proteins as FOXA1-WT, and thus indicating an intrinsic deficiency in the mutants’ abilities to restore expression of endogenous liver genes in a Foxa1 knock-down background. Two experiments were repeated independently with similar results. Full blots are presented in the Source Data files. Source data
a, Generation of Foxa2WT-tRFP and Foxa2∆Hx-tRFP knock-in alleles with targeting and exchange cassettes. b, Frequency of genotypes resulting from heterozygous intercrosses of Foxa2WT-tRFP/WT at embryonic stages. c, Box and whisker plots show stage distribution at wild-type, heterozygous, and homozygous of Foxa2WT-tRFP and Foxa2∆Hx-tRFP embryos at E8.5. The bottom and top of the boxes correspond to the 25th and 75th percentiles, and the internal band is the 50th percentile (median). The ends of the whiskers represent 1.5 times the IQR. The points represent outliers. The indicated P-values are obtained by one-sided Wilcoxon rank sum test.
a, Bright field images of Foxa2∆Hx-tRFP/WT and Foxa2∆Hx-tRFP/∆Hx-tRFP (with a gross phenotype) littermate at E12.5 from heterozygous intercrosses. Images are representative of the numbers of embryos indicated in Fig. 3a. b, Representative FACS pattern of FOXA-tRFP-high (P5 gate), -middle (P4 gate), and -negative (P3 gate) cells from Foxa2tRFP/tRFP E7.5 embryos. Foxa2WT/WT and Foxa2tRFP/WT samples were loaded only for setting the gates, but not for the sorting. The FOXA2-tRFP-middle (P4 gate) was set by avoiding autofluorescence and including up to the maximum tRFP intensity of heterozygous (Foxa2tRFP/WT) cells expressed. The FOXA2-tagRFP-high (P5 gate) exhibited a higher tagRFP signal than heterozygous cells. The FACS experiments were repeated more than 40 times independently with similar results.
a, Heatmaps show DESeq adjusted RNA-seq counts for all differentially expressed genes with adjusted p-value < 0.1 (by one-sided Wald test with FDR correction at 10%). The individual replicates of wild-type (Foxa2WT-tRFP /WT-tRFP) and ∆Hx (Foxa2∆Hx-tRFP/∆Hx-tRFP) in FOXA2-tRFP-high and -middle cells were presented. n = 3 biologically independent RNA-seq datasets per group. b, RNA-seq tracks of each biological replicate of FOXA2-tRFP-mid cells from E7.5 Foxa2WT/WT (green) and Foxa2∆Hx-tRFP/∆Hx-tRFP (gray) embryos at down-regulated gene loci in Foxa2∆Hx-tRFP/∆Hx-tRFP.
Extended Data Fig. 8 Deletion of α-helical region of FOXA2 alters the accessible chromatin sites in E7.5 embryos.
a, de novo motif enrichment analysis at differential open chromatin sites (ATAC-seq peaks) between FOXA2[WT]-tRFP-high cells and FOXA2[WT]-tRFP-middle cells. P-value (by one-sided Monte Carlo simulation with FDR controlled at 5%) and % targets are indicated in parentheses. n = 2 biologically independent ATAC-seq datasets per group. b, de novo motif enrichment analysis at differential open chromatin sites (ATAC-seq peaks) of “wild-type (Foxa2WT-tRFP /WT-tRFP)-specific”, “∆Hx (Foxa2∆Hx-tRFP/∆Hx-tRFP)-specific”, and “wild-type and ∆Hx common” open chromatin sites in E7.5 FOXA2-tRFP-high cells and FOXA2-tRFP-middle cells. P-value (by one-sided Monte Carlo simulation with FDR controlled at 5%) and % targets are indicated in parentheses. n = 2 biologically independent ATAC-seq datasets per group.
Extended Data Fig. 9 Deletion of α-helical region of FOXA2 alters gene expression and accessible chromatin landscapes in E7.5 embryos.
a, GO term enrichment analysis of downregulated and upregulated genes in FOXA2-tRFP-high and FOXA2-tRFP-middle cells. P-value (by one-sided EASE/Fisher’s exact test) and gene count are indicated in parentheses. n = 3 biologically independent RNA-seq datasets per group. b, The distribution of WT-specific, ∆Hx-specific, and WT-∆Hx common open chromatin sites at non-overlapped genomic features.
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Iwafuchi, M., Cuesta, I., Donahue, G. et al. Gene network transitions in embryos depend upon interactions between a pioneer transcription factor and core histones. Nat Genet (2020). https://doi.org/10.1038/s41588-020-0591-8