Histone H4 acetyl-methyllysine marks accessible chromatin that resists compaction.

Global Levels of Histone Modifications Predict Prognosis in Different Cancers

CTCF-binding sites are frequently found at the boundaries of chromatin domains enriched with specific histone modifications in vertebrate genomes, where they function as barriers that limit the spreading of these modifications. However, their role in domains marked by H3K9me2 or H3K9me3 remains poorly understood. In this study, we analyzed CTCF-binding sites and the distribution of H3K9me2 and H3K9me3 around them in human K562 cells, and investigated the spreading of these modifications following CTCF depletion. Genome-wide analysis revealed that certain CTCF-binding sites are located at the boundaries of chromatin domains enriched with H3K9me2 or H3K9me3. Upon loss of CTCF binding, both modifications spread to neighboring chromatin at over half of the boundaries delimiting highly and distinctly enriched domains. Boundaries where such spreading did not occur were flanked by chromatin in a more active state compared to those where spreading occurred. Gene transcription was significantly reduced in neighboring regions where H3K9me2 or H3K9me3 had spread. Further comprehensive analysis revealed that this transcriptional reduction was specifically associated with the spread of H3K9me2. These findings suggest that CTCF-binding sites demarcate chromatin domains enriched with H3K9me2 or H3K9me3 and act as barriers to restrict the spread of these modifications in mammalian cells. Notably, the spread of H3K9me2 appears to play a specific role in transcriptional repression.

Transcription regulation by histone modifications is a major contributing factor to the structural and functional diversity in biology. These modifications are encrypted as histone codes or histone languages and function to establish and maintain heritable epigenetic codes that define the identity and the fate of the cell. Despite recent advances revealing numerous histone modifications associated with transcription regulation, how such modifications dictate the process of transcription is not fully understood. Here we describe spatial and temporal analyses of the histone modifications that are introduced during estrogen receptor α (ERα)-activated transcription. We demonstrated that aborting RNA polymerase II caused a disruption of the histone modifications that are associated with transcription elongation but had a minimal effect on modifications deposited during transcription initiation. We also found that the histone H3S10 phosphorylation mark is catalyzed by mitogen- and stress-activated protein kinase 1 (MSK1) and is recognized by a 14-3-3ζ/14-3-3ε heterodimer through its interaction with H3K4 trimethyltransferase SMYD3 and the p52 subunit of TFIIH. We showed that H3S10 phosphorylation is a prerequisite for H3K4 trimethylation. In addition, we demonstrated that SET8/PR-Set7/KMT5A is required for ERα-regulated transcription and its catalyzed H4K20 monomethylation is implicated in both transcription initiation and elongation. Our experiments provide a relatively comprehensive analysis of histone modifications associated with ERα-regulated transcription and define the biological meaning of several key components of the histone code that governs ERα-regulated transcription.

CTCF-binding sites are frequently found at the boundaries of chromatin domains enriched with specific histone modifications in vertebrate genomes, where they function as barriers that limit the spreading of these modifications. However, their role in domains marked by H3K9me2 or H3K9me3 remains poorly understood. In this study, we analyzed CTCF-binding sites and the distribution of H3K9me2 and H3K9me3 around them in human K562 cells, and investigated the spreading of these modifications following CTCF depletion. Genome-wide analysis revealed that certain CTCF-binding sites are located at the boundaries of chromatin domains enriched with H3K9me2 or H3K9me3. Upon loss of CTCF binding, both modifications spread to neighboring chromatin at over half of the boundaries delimiting highly and distinctly enriched domains. Boundaries where such spreading did not occur were flanked by chromatin in a more active state compared to those where spreading occurred. Gene transcription was significantly reduced in neighboring regions where H3K9me2 or H3K9me3 had spread. Further comprehensive analysis revealed that this transcriptional reduction was specifically associated with the spread of H3K9me2. These findings suggest that CTCF-binding sites demarcate chromatin domains enriched with H3K9me2 or H3K9me3 and act as barriers to restrict the spread of these modifications in mammalian cells. Notably, the spread of H3K9me2 appears to play a primary role in transcriptional repression.

H3K18ac Primes Mesendodermal Differentiation upon Nodal Signaling.

CCCTC-binding factor (CTCF) binding sites are hotspots of genome instability. Although many factors have been associated with CTCF binding site fragility, no study has integrated all fragility-related factors to understand the mechanism(s) of how they work together. Using an unbiased, genome-wide approach, we found that DNA double-strand breaks (DSBs) are enriched at strong, but not weak, CTCF binding sites in five human cell types. Energetically favorable alternative DNA secondary structures underlie strong CTCF binding sites. These structures coincided with the location of topoisomerase II (TOP2) cleavage complex, suggesting that DNA secondary structure acts as a recognition sequence for TOP2 binding and cleavage at CTCF binding sites. Furthermore, CTCF knockdown significantly increased DSBs at strong CTCF binding sites and at CTCF sites that are located at topologically associated domain (TAD) boundaries. TAD boundary-associated CTCF sites that lost CTCF upon knockdown displayed increased DSBs when compared to the gained sites, and those lost sites are overrepresented with G-quadruplexes, suggesting that the structures act as boundary insulators in the absence of CTCF, and contribute to increased DSBs. These results model how alternative DNA secondary structures facilitate recruitment of TOP2 to CTCF binding sites, providing mechanistic insight into DNA fragility at CTCF binding sites.

Trimethylation of lysine 9 in histone H3 (H3K9me3) enrichment is a characteristic of pericentric heterochromatin. The hypothesis of a stepwise mechanism to establish and maintain this mark during DNA replication suggests that newly synthesized histone H3 goes through an intermediate methylation state to become a substrate for the histone methyltransferase Suppressor of variegation 39 (Suv39H1/H2). How this intermediate methylation state is achieved and how it is targeted to the correct place at the right time is not yet known. Here, we show that the histone H3K9 methyltransferase SetDB1 associates with the specific heterochromatin protein 1alpha (HP1alpha)-chromatin assembly factor 1 (CAF1) chaperone complex. This complex monomethylates K9 on non-nucleosomal histone H3. Therefore, the heterochromatic HP1alpha-CAF1-SetDB1 complex probably provides H3K9me1 for subsequent trimethylation by Suv39H1/H2 in pericentric regions. The connection of CAF1 with DNA replication, HP1alpha with heterochromatin formation and SetDB1 for H3K9me1 suggests a highly coordinated mechanism to ensure the propagation of H3K9me3 in pericentric heterochromatin during DNA replication.

15-Lipoxygenase-1 (15-LOX-1) contributes significantly to inflammation regulation and terminal cell differentiation. 15-LOX-1 is transcriptionally silenced in cancer cells, and its transcriptional reactivation (e.g. via histone deacetylase inhibitors (HDACIs)) is essential for restoring terminal cell differentiation to cancer cells. STAT-6 acetylation via the histone acetyltransferase KAT3B has been proposed to be necessary for 15-LOX-1 transcriptional activation. However, the exact mechanism underlying 15-LOX-1 transcriptional reactivation in cancer cells is still undefined, especially in regard to the contribution of 15-LOX-1 promoter histone modifications. We therefore examined the relative mechanistic contributions of 15-LOX-1 promoter histone modifications and STAT-6 to 15-LOX-1 transcriptional reactivation by HDACIs in colon cancer cells. We found that: 1) histone H3 and H4 acetylation in the 15-LOX-1 promoter through KAT3B was critical to 15-LOX-1 transcriptional activation; 2) 15-LOX-1 transcription was activated independently from STAT-6; and 3) dimethyl-histone H3 lysine 9 (H3K9me2) demethylation in the 15-LOX-1 promoter via the histone lysine demethylase KDM3A was an early and specific histone modification and was necessary for activation of transcription. These findings demonstrate that histone modification in the 15-LOX-1 promoter is important to 15-LOX-1 transcriptional silencing in colon cancer cells and that HDACIs can activate gene transcription via KDM3A demethylation of H3K9me2.

BackgroundSpecific histone modifications play important roles in chromatin functions; i.e., activation or repression of gene transcription. This participation must occur as a dynamic process. Nevertheless, most of the histone modification maps reported to date provide only static pictures that link certain modifications with active or silenced states. This study, however, focuses on the global histone modification variation that occurs in response to the transcriptional reprogramming produced by a physiological perturbation in yeast.ResultsWe did a genome-wide chromatin immunoprecipitation analysis for eight specific histone modifications before and after saline stress. The most striking change was rapid acetylation loss in lysines 9 and 14 of H3 and in lysine 8 of H4, associated with gene repression. The genes activated by saline stress increased the acetylation levels at these same sites, but this acetylation process was quantitatively minor if compared to that of the deacetylation of repressed genes. The changes in the tri-methylation of lysines 4, 36 and 79 of H3 and the di-methylation of lysine 79 of H3 were slighter than those of acetylation. Furthermore, we produced new genome-wide maps for seven histone modifications, and we analyzed, for the first time in S. cerevisiae, the genome-wide profile of acetylation of lysine 8 of H4.ConclusionsThis research reveals that the short-term changes observed in the post-stress methylation of histones are much more moderate than those of acetylation, and that the dynamics of the acetylation state of histones during activation or repression of transcription is a much quicker process than methylation.Electronic supplementary materialThe online version of this article (doi:10.1186/1471-2164-15-247) contains supplementary material, which is available to authorized users.

Gene expression in estrogen receptor (ER)-positive breast cancer cells is predominantly driven by ER-estradiol (E2) signaling. However, the effects of ER-E2 signaling on the overall chromatin architecture and accessibility to transcription regulators are not well understood. Chromatin architecture is a key determinant to gene expression patterns and therefore a comprehensive notation is essential for the interpretation of the ER cistrome. To investigate the effects of E2 signaling on chromatin architecture, we employed Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq), a high-throughput method to map chromatin accessibility via a hyperactive transposase that inserts sequencing tags into open regions. ATAC-seq was performed on MCF-7 cells, an ERα+ breast cancer cell line, treated with E2 or vehicle control for 1 or 3 hours. Results from the ATAC-seq were compared to RNA-seq data of 3-hours E2-treated MCF-7 cells to determine which regions of E2-regulated open or closed chromatin coincide with increased or decreased respective gene expression. Interestingly, we observed greater chromatin accessibility changes in genes repressed by E2 compared to genes induced by E2. Integration of ATAC-seq and RNA-seq data revealed distinct categories of E2-induced chromatin remodeling and downstream gene expression including increased chromatin accessibility, but repression of the corresponding gene, as well as decreased chromatin accessibility, but induction of the corresponding gene. Validation of the chromatin configurations predicted by ATAC-seq of select corresponding genes is currently being performed by ChIP-qPCR against specific histone modifications, such as H3K27Ac, a marker of open chromatin. To mechanistically dissect the distinct modes of E2 chromatin remodelling, we performed motif enrichment analysis of chromatin regions affected by E2. This analysis showed expected enrichment of estrogen response element (ERE), and pioneer factor FOXA1 and GATA3 response elements in E2-inducible genes with enhanced chromatin accessibility after E2-treatment. Regions in which the corresponding mRNA showed decreased transcript levels but displayed open chromatin in their promoter or enhancer region were enriched for the PBX3 motif and the ERE motif, respectively. Our results reveal previously unrecognized modes of E2-mediated restructuring of chromatin architecture and its correlation with transcription dynamics. This study provides grounds for future work to study fundamental relationships between chromatin accessibility and gene regulation in breast cancer under E2 and anti-estrogen treated conditions as well as in metastatic breast cancers with ERα mutations. Citation Format: Taylor M. Parker, Duojiao Chen, Poornima Bhat-Nakshatri, Xiaona Chu, Yunlong Liu, Yue Wang, Harikrishna Nakshatri. Non-linear relationship between estradiol-regulated changes in chromatin architecture and gene expression [abstract]. In: Proceedings of the American Association for Cancer Research Annual Meeting 2019; 2019 Mar 29-Apr 3; Atlanta, GA. Philadelphia (PA): AACR; Cancer Res 2019;79(13 Suppl):Abstract nr 1006.

Gene expression in estrogen receptor (ER)-positive breast cancer cells is predominantly driven by ER-estradiol (E2) signaling. However, the effects of ER-E2 signaling on the overall chromatin architecture and accessibility to transcription regulators are not well understood. Chromatin architecture is a key determinant to gene expression patterns and therefore a comprehensive notation is essential for the interpretation of the ER cistrome. To investigate the effects of E2 signaling on chromatin architecture, we employed Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq), a high-throughput method to map chromatin accessibility via a hyperactive transposase that inserts sequencing tags into open regions. ATAC-seq was performed on MCF-7 cells, an ERα+ breast cancer cell line, treated with E2 or vehicle control for 1 or 3 hours. Results from the ATAC-seq were compared to RNA-seq data of 3-hours E2-treated MCF-7 cells to determine which regions of E2-regulated open or closed chromatin coincide with increased or decreased respective gene expression. Interestingly, we observed greater chromatin accessibility changes in genes repressed by E2 compared to genes induced by E2. Integration of ATAC-seq and RNA-seq data revealed distinct categories of E2-induced chromatin remodeling and downstream gene expression including increased chromatin accessibility, but repression of the corresponding gene, as well as decreased chromatin accessibility, but induction of the corresponding gene. Validation of the chromatin configurations predicted by ATAC-seq of select corresponding genes is currently being performed by ChIP-qPCR against specific histone modifications, such as H3K27Ac, a marker of open chromatin. To mechanistically dissect the distinct modes of E2 chromatin remodelling, we performed motif enrichment analysis of chromatin regions affected by E2. This analysis showed expected enrichment of estrogen response element (ERE), and pioneer factor FOXA1 and GATA3 response elements in E2-inducible genes with enhanced chromatin accessibility after E2-treatment. Regions in which the corresponding mRNA showed decreased transcript levels but displayed open chromatin in their promoter or enhancer region were enriched for the PBX3 motif and the ERE motif, respectively. Our results reveal previously unrecognized modes of E2-mediated restructuring of chromatin architecture and its correlation with transcription dynamics. This study provides grounds for future work to study fundamental relationships between chromatin accessibility and gene regulation in breast cancer under E2 and anti-estrogen treated conditions as well as in metastatic breast cancers with ERα mutations. Citation Format: Taylor M. Parker, Duojiao Chen, Poornima Bhat-Nakshatri, Xiaona Chu, Yunlong Liu, Yue Wang, Harikrishna Nakshatri. Non-linear relationship between estradiol-regulated changes in chromatin architecture and gene expression [abstract]. In: Proceedings of the American Association for Cancer Research Annual Meeting 2019; 2019 Mar 29-Apr 3; Atlanta, GA. Philadelphia (PA): AACR; Cancer Res 2019;79(13 Suppl):Abstract nr 1006.

Low H3K27me3 deposition at CYP82E4 determines the nicotinic conversion rate in Nicotiana tabacum

Article Figures and data Abstract Editor's evaluation Introduction Results Discussion Materials and methods Appendix 1 Data availability References Decision letter Author response Article and author information Metrics Abstract Transcription factors play a determining role in lineage commitment and cell differentiation. Interferon regulatory factor 8 (IRF8) is a lineage determining transcription factor in hematopoiesis and master regulator of dendritic cells (DC), an important immune cell for immunity and tolerance. IRF8 is prominently upregulated in DC development by autoactivation and controls both DC differentiation and function. However, it is unclear how Irf8 autoactivation is controlled and eventually limited. Here, we identified a novel long non-coding RNA transcribed from the +32 kb enhancer downstream of Irf8 transcription start site and expressed specifically in mouse plasmacytoid DC (pDC), referred to as lncIrf8. The lncIrf8 locus interacts with the lrf8 promoter and shows differential epigenetic signatures in pDC versus classical DC type 1 (cDC1). Interestingly, a sequence element of the lncIrf8 promoter, but not lncIrf8 itself, is crucial for mouse pDC and cDC1 differentiation, and this sequence element confers feedback inhibition of Irf8 expression. Taken together, in DC development Irf8 autoactivation is first initiated by flanking enhancers and then second controlled by feedback inhibition through the lncIrf8 promoter element in the +32 kb enhancer. Our work reveals a previously unrecognized negative feedback loop of Irf8 that orchestrates its own expression and thereby controls DC differentiation. Editor's evaluation Authors provide valuable evidence identifying a lncRNA transcribed specifically in the pDC subtype from the +32Kb promoter region which is also the region for the enhancer for Irf8 specifically in the cDC1 subtype. With convincing methodology, they provide in-depth analysis about the possible role of lncIrf8, and its promoter region and cross-talk with Irf8 promoter to identify that it is not the lncIRF8 itself but its promoter region that is crucial for pDC and cDC1 differentiation conferring feedback inhibition of Irf8 transcription. The work will be of interest to immunologists working on immune cell development. https://doi.org/10.7554/eLife.83342.sa0 Decision letter Reviews on Sciety eLife's review process Introduction Lineage-determining transcription factors (TF) are master regulators of gene programs that frequently initiate self-reinforcing loops by autoactivation. TF autoactivation is important for cells to pass restriction points during development (referred to as points of no return) and to enforce cellular identity. Molecular circuitries of autoactivation have been studied for several TF, such as GATA-binding factor 1 (GATA1), PU.1 (encoded by Spi1), CCAAT enhancer-binding protein α and ε (C/EBPα and ε; Graf and Enver, 2009; Loughran et al., 2020; Nishimura et al., 2000; Okuno et al., 2005; Theilgaard-Mönch et al., 2022). A further example is interferon regulatory factor 8 (IRF8), which shows autoactivation in cooperation with basic leucine zipper ATF-like transcription factor 3 (BATF3; Anderson et al., 2021; Grajales-Reyes et al., 2015). An important principle in nature is negative feedback control to avoid signal overshooting and toxicity. Negative feedback control applies also to lineage-determining TF; however, there is a paucity on our knowledge of the molecular mechanisms involved. IRF8 is a hematopoietic TF positioned at the center of the regulatory gene network for dendritic cell (DC) development (Anderson et al., 2021; Belz and Nutt, 2012; Chauvistré and Seré, 2020; Kim et al., 2020; Lin et al., 2015; Nutt and Chopin, 2020; Tamura et al., 2015; Verlander et al., 2022). IRF8 is a member of the interferon regulatory factor (IRF) family of TF. Initially members of this TF family were found to mediate the induction of interferon induced genes, but are now known to serve diverse functions in regulating the immune system (Honda and Taniguchi, 2006; Tamura et al., 2008). Irf8 knockout mice show abnormal development of classical DC type 1 (cDC1) and plasmacytoid DC (pDC) (Durai et al., 2019; Schiavoni et al., 2002; Sichien et al., 2016; Tsujimura et al., 2003). Irf8 is prominently upregulated during DC development by autoactivation (Grajales-Reyes et al., 2015; Lin et al., 2015), yet how Irf8 autoactivation is controlled and eventually limited, and the epigenetic mechanisms involved is largely unknown. Irf8 expression in hematopoietic cells is induced and maintained by enhancers located at –50 kb,+32 kb,+41 kb and +56 kb relative to Irf8 transcription start site (TSS) (Anderson et al., 2021; Bagadia et al., 2019; Durai et al., 2019; Grajales-Reyes et al., 2015; Murakami et al., 2021; Schönheit et al., 2013). Enhancers are cis-regulatory sequences with multiple TF binding sites that cooperatively bind TF and thereby activate transcription, as demonstrated by many studies including our work (Davidson et al., 1986; Long et al., 2016; Wildeman et al., 1986; Zenke et al., 1986). Enhancers regulate complex gene networks and can also produce non-coding RNA, referred to as enhancer RNA (eRNA). eRNA serve as an indicator for enhancer activity and some eRNA have an activity on their own and act in cis or trans to regulate cell fate decisions (Sartorelli and Lauberth, 2020; Statello et al., 2021). Enhancer-associated long non-coding RNA (lncRNA) represent a class of lncRNA transcribed from active enhancers. Thus, eRNA and enhancer-associated lncRNA provide opportunities to detect enhancer activity and to investigate enhancer function. DC are highly specialized immune cells that play a critical role in regulating innate and adaptive immune responses (Cabeza-Cabrerizo et al., 2021). DC develop from hematopoietic stem cells (HSC) via successive steps of lineage commitment and differentiation. More specifically, HSC develop into multipotent progenitors (MPP) that are committed to DC restricted common DC progenitors (CDP) and differentiate into classic DC (cDC) type 1 and type 2 (cDC1 and cDC2, respectively) and pDC (Anderson et al., 2021; Cabeza-Cabrerizo et al., 2021; Ginhoux et al., 2022; Nutt and Chopin, 2020; Rodrigues and Tussiwand, 2020). pDC were recently also shown to develop from lymphoid progenitors (Dress et al., 2019; Rodrigues et al., 2018; Rodrigues and Tussiwand, 2020). Differential expression of Irf8 regulates DC and monocyte specification in a dose-dependent manner (Cytlak et al., 2020; Murakami et al., 2021). Irf8 expression starts at the CDP stage, and is high in pDC and cDC1, which is attributed to the autoactivation of Irf8 during DC subsets specification (Grajales-Reyes et al., 2015; Lin et al., 2015). Interestingly, IRF8 can act as a transcriptional activator or repressor in hematopoiesis by interacting with different partner TF and binding to specific DNA sequences (Tamura et al., 2015). As an activator, IRF8 binds to its own promoter in DC differentiation, which is considered as the autoactivation capacity of Irf8 (Grajales-Reyes et al., 2015; Lin et al., 2015). For instance, IRF8 interacts with partner TF, such as PU.1, to initiate Irf8 autoactivation at the CDP stage (Grajales-Reyes et al., 2015). Inversely, IRF8 inhibits C/EBPα activity in neutrophil differentiation (Kurotaki et al., 2014). IRF8 also represses C/EBPβ to generate and maintain DC lineage-specific enhancer landscapes (Bornstein et al., 2014). In addition, IRF8 is important for the Myc-Mycl transition in DC differentiation (Anderson lll et al., 2021). IRF8 represses Myc expression in progenitors, while IRF8 at high levels interacts with PU.1 and drives Mycl expression (Anderson lll et al., 2021). All this emphasizes the central position of IRF8 in coordinating the gene network that regulates DC differentiation and function. During DC differentiation, the Irf8 gene locus shows high epigenetic dynamics, including histone modifications and TF binding identified by ChIP-seq (Chauvistré and Seré, 2020; Durai et al., 2019; Grajales-Reyes et al., 2015; Lin et al., 2015), chromatin accessibility measured by ATAC-seq (Kurotaki et al., 2019; Li et al., 2019), and three-dimensional chromatin structure remodeling determined by chromosome conformation capture (3 C) (Kurotaki et al., 2022; Schönheit et al., 2013). All this emphasizes the impact of epigenetic regulators on Irf8 gene activity in DC differentiation. Notably, Irf8 is flanked by multiple enhancers at –50 kb,+32 kb,+41 kb, and +56 kb that regulate Irf8 expression in hematopoietic cells (Anderson et al., 2021; Murakami et al., 2021). These four enhancers were found to be driven by PU.1, BATF3, E proteins and Runt-related transcription factor (RUNX)-core binding factor beta (CBFβ) (RUNX-CBFβ), respectively (Bagadia et al., 2019; Durai et al., 2019; Grajales-Reyes et al., 2015; Murakami et al., 2021; Schönheit et al., 2013). Chromatin conformation, particularly enhancer promoter interactions, provides a platform for TF-driven gene regulation and serves as a driving force for cell-fate determinations (Misteli and Finn, 2021; Oudelaar and Higgs, 2021; Stadhouders et al., 2019). Schönheit et al. demonstrated Irf8 promoter interactions with its upstream enhancers by quantitative 3 C (Schönheit et al., 2013). In this study PU.1 was found to regulate chromatin remodeling between the –50 kb enhancer and the Irf8 promoter in myeloid differentiation. In a recent study Kurotaki et al., 2022 determined the higher-order chromatin structure in DC progenitors, cDC1 and cDC2 on a genome-wide scale by Hi-C. In this study, reorganization of chromatin conformation at DC-specific gene loci was observed during cDC differentiation, and IRF8 was found to promote chromatin activation in DC progenitors leading to cDC lineage-specific gene expression. However, high resolution maps of the physical chromatin interactions of the Irf8 promoter with upstream and downstream enhancers in the full complement of DC subsets, including pDC, are required for understanding Irf8 regulation during DC differentiation. Frequently, chromatin data, including ATAC-seq and/or ChIP-seq data, are used to identify regulatory elements of gene transcription. Here we embarked on a different approach and searched for lncRNA, which by themselves might have regulatory functions or are indicative of enhancer activity. We identified a novel lncRNA transcribed from the Irf8 +32 kb enhancer, which is specifically expressed in pDC, referred to as lncIrf8. We found that the lncIrf8 promoter element but not lncIrf8 itself impacts pDC and cDC1 development. Thus, lncIrf8 acts as an indicator for the Irf8 +32 kb enhancer activity. Importantly, our study revealed a previously unrecognized negative feedback loop of Irf8 in DC differentiation. Irf8 first activates its expression by autoactivation via the +32 kb enhancer and second limits its own expression through the lncIrf8 promoter element in the +32 kb enhancer. Results lncIrf8 marks a pDC-specific Irf8 enhancer element Irf8 expression in DC development is subject to complex epigenetic regulation. Here, we used an integrated approach with RNA-seq, ATAC-seq, ChIP-seq and Capture-C to track the dynamics of gene expression, histone modification and chromatin conformation in the sequel MPP, CDP, pDC, cDC1, and cDC2 (Figure 1, Figure 1—figure supplements 1 and 2). Figure 1 with 3 supplements see all Download asset Open asset Irf8 epigenetic signatures and promoter-enhancer interaction maps during DC differentiation. (A) Gene expression and epigenetic signatures of Irf8 downstream region in MPP, CDP, pDC, all cDC, cDC1, and cDC2 are visualized by IGV browser. Gene expression was measured by RNA-seq, chromatin accessibility by ATAC-seq, H3K27ac and IRF8 binding by ChIP-seq. Positions of Irf8 3’ end, Irf8 enhancers, pDC specific lncIrf8 and cDC1 specific Tcons_00190258 lncRNA are indicated. For RNA-seq - and +strands are shown. Scale bar: 5 kb. (B) Physical interactions of Irf8 promoter with flanking sequences in MPP, CDP, pDC, cDC1, and cDC2 by nuclear-titrated (NuTi) Capture-C (turquoise), and CTCF binding by ChIP-seq in DC (Garber et al., 2012). Mean numbers of unique interactions normalized to a 300 kb region around the Irf8 promoter viewpoint (green triangle) and scaled by a factor of 1,000,000 are shown (n=2). The orientations of CTCF binding are indicated with blue and red arrows. Tcons_00190258 refers to the cDC1-specific lncRNA shown in (A). Scale bar: 100 kb. (C) Comparations of the chromatin interactions with Irf8 promoter in pDC, cDC1 and cDC2. Differential tracks were created by subtraction of the mean normalized tracks of (B). Pairwise comparisons are shown and color coded. Turquoise and orange tracks represent specific interactions with the Irf8 promoter in the indicated cell types. Scale bar: 100 kb. Purple bars and lines indicate the position of flanking enhancers relative to Irf8 TSS. The purple bars from left to right represent –50 kb, –34 kb, –26 kb, –16 kb,+27 kb,+32 kb,+38 kb,+41 kb,+47 kb,+56 kb and +62 kb enhancer, respectively (panels B and C). Irf8 +32 kb enhancer is highlighted by purple box. We performed de novo transcript assembly of the RNA-seq data and detected two previously unknown transcripts without coding potential downstream of Irf8: a pDC specific lncRNA (Tcons_00190250) in the following referred to as lncIrf8 and a cDC1 specific lncRNA (Tcons_00190258; Figure 1A and Figure 1—figure supplement 1). lncIrf8 and Tcons_00190258 show the same expression pattern in pDC and cDC1, respectively, in BM and spleen (Figure 1—figure supplement 3), as revealed by reanalyzing scRNA-seq and bulk RNA-seq data (Pang et al., 2022; Rodrigues et al., 2018). lncIrf8 is transcribed within an enhancer region located 32 kb downstream of the Irf8 TSS labeled by H3K27ac and H3K4me1 and occupied by DC differentiation-associated TF, such as IRF8 and PU.1 (Figure 1A and Figure 1—figure supplement 1). This region is largely devoid of H3K9me3, a chromatin modification frequently associated with heterochromatin, indicating an open chromatin configureuration in DC (Figure 1—figure supplement 1). In addition, sequences of this region have been implicated in DC development and referred to as +32 kb enhancer (Durai et al., 2019). Thus, we proceeded to study lncIrf8 in detail. ATAC-seq analysis revealed further details of the lncIrf8 region in CDP, pDC, cDC1 and cDC2 (Figure 1A, Figure 2A and Figure 1—figure supplement 1). In cDC1 the prominent ATAC-seq and IRF8 peaks mark the cDC1 specific +32 kb enhancer (Durai et al., 2019). In pDC the ATAC-seq peak is smaller and shifted further towards downstream but aligns well with the valley in the prominent H3K27ac peak. This ATAC-seq peak marks the lncIrf8 promoter and aligns with p300 (Durai et al., 2019) and H3K4me3 (Figure 2A and Figure 1—figure supplement 1). All this indicates that this chromatin region is open and transcriptionally active in pDC, enabling lncIrf8 transcription. Figure 2 with 4 supplements see all Download asset Open asset IncIrf8 promoter KO compromises pDC and cDC1 development in vitro. (A) Genomic anatomy of lncIrf8 locus determined by 3’ and 5’ RACE PCR. Blue box, exon 2 and 3 (48 bp and 468 bp, respectively). The 1010 bp intron and polyA tail are shown. Data of RNA-seq, ATAC-seq, ChIP-seq of H3K27ac (enhancer mark) and H3K4me3 (active promoter mark, near TSS) are visualized by IGV browser for the indicated cell populations (pDC, all cDC, cDC1 and cDC2). Grey box, lncIrf8 promoter KO region; open box, cDC1 specific +32 kb enhancer by Durai et al., 2019. Irf8 +32 kb enhancer based on the H3K27ac enhancer mark is indicated with a purple line. Scale bar: 1 kb. (B) Gene expression of lncIrf8 and Irf8 in lncIrf8 promoter KO and control at day 0, 5, and 7 of Flt3L directed DC differentiation. Gene expression was determined by RT-qPCR and normalized to GAPDH. n=4. (C) Representative flow cytometry analysis of Flt3L directed DC differentiation of lncIrf8 promoter KO HoxB8 MPP and control (Lutz et al., 2022; Xu et al., 2022). pDC, all cDC, cDC1, and cDC2 were gated as in Figure 2—figure supplement 1E and are shown. Bar diagrams depict quantification of pDC, cDC1 and cDC2 normalized to living single cells on DC differentiation day 0, 3, 5, 7, and 9. n=6–7. (D) Representative phase-contrast microscopy images of lncIrf8 promoter KO HoxB8 MPP and control on day 7 of Flt3L directed DC differentiation. Scale bar: 200 μm. (E) Representative flow cytometry analysis of spontaneous DC differentiation of lncIrf8 promoter KO HoxB8 MPP and control with growth factors but without E2 (Lutz et al., 2022; Xu et al., 2022) at day 8. Gr1+ monocytes and CD11c+ DC are shown. Quantification of Gr1+ monocytes of living single cells on day 3, 6, 8, and 10 of spontaneous DC differentiation. n=6, lncIrf8 promoter KO; n=4, control. Empty gRNA vector or non-targeting gRNA vector HoxB8 MPP were used as controls. Data represent mean ± SD of at least three independent experiments with different HoxB8 MPP clones of lncIrf8 promoter KO and control without deletion. *p<0.05, **p<0.01, ***p<0.001, multiple t-tests. Data that have no difference (p>0.05) are not labeled. Next, we determined the chromatin conformation of the Irf8 locus and the lncIrf8 region. We generated interaction profiles by nuclear-titrated (NuTi) Capture-C in MPP, CDP, pDC, cDC1, and cDC2 (Figure 1—figure supplement 2A and B) using Irf8 promoter as viewpoint. The Irf8 promoter shows multiple interactions with regions spanning up to ~100 kb upstream and downstream of Irf8 (Figure 1B and Figure 1—figure supplement 1). In pDC, the Irf8 promoter interactions are stronger with the upstream sequences than with downstream sequences (Figure 1C and Figure 1—figure supplement 2C). In cDC1 Irf8 promoter interactions are more confined to the regions downstream of Irf8 compared to MPP, CDP and pDC (Figure 1C and Figure 1—figure supplement 2C). This suggests that upstream and downstream sequences of Irf8 gene are involved in differentially regulating Irf8 expression and controlling the development of pDC and cDC1, respectively. The CCCTC-binding factor (CTCF) is important for regulation of chromatin conformation through loop extrusion (Sanborn et al., 2015) and we therefore visualized CTCF binding sites in the Irf8 locus in DC (Garber et al., 2012). Interestingly, most of the Irf8 flanking enhancers (Durai et al., 2019; Grajales-Reyes et al., 2015; Murakami et al., 2021; Schönheit et al., 2013) are located within convergent CTCF binding sites upstream and downstream of the Irf8 gene (Figure 1B and Figure 1—figure supplement 1). There are also multiple interactions within this region without convergent CTCF binding sites, suggesting interactions with Irf8 promoter in a CTCF independent manner, such as by TF binding, active histone modifications and gene transcription (Figure 1B and Figure 1—figure supplement 1; Owens et al., 2022). Surprisingly, in pDC H3K27ac at the lncIrf8 promoter is high, but this locus shows less interactions with the Irf8 promoter in pDC compared to CDP, cDC1 and cDC2 (Figure 1C and Figure 1—figure supplement 2C). In addition, in pDC the IRF8 protein occupancy at the lncIrf8 promoter is low and much higher in cDC (Figure 1A and Figure 1—figure supplement 1; Durai et al., 2019; Grajales-Reyes et al., 2015). These observations warrant further studies and we thus proceeded to investigate the lncIrf8 locus in detail. lncIrf8 promoter KO compromises pDC and cDC1 development First, we annotated lncIrf8. Our de-novo transcript assembly of RNA-seq data revealed different isoforms of lncIrf8, with the most prominent isoform comprising exon 2 and 3 (Figure 1A, Figure 2A and Figure 1—figure supplement 1). Additionally, 3' end and 5' end RACE PCR confirmed the anatomy of this lncIrf8 isoform: two exons, one intron, and a polyA tail (Figure 2A). As expected lncIrf8 is not conserved across species (data not shown), which is in line with the general characteristics of lncRNA. Then second, we deleted 300 bp in the lncIrf8 promoter by CRISPR/Cas9 editing in conditionally immortalized HoxB8 MPP (Figure 2A and Figure 2—figure supplement 1A–D). The lncIrf8 promoter is located in the Irf8 +32 kb enhancer region and is in close proximity to the cDC1 specific +32 kb enhancer (Durai et al., 2019; Figure 2A and Figure 2—figure supplement 2). The 300 bp deletion comprises the H3K4me3 promoter mark and is confined to open chromatin identified by ATAC-seq and positioned in the valley of the H3K27ac mark (Figure 2A). Additionally, it contains binding sites for IRF8, PU.1, and BATF3 TF, which are important for DC development (Figure 2A and Figure 2—figure supplement 2B). Generation of a precise deletion requires clonal cell populations, which is hardly achieved in somatic cells due to their limited lifespan. Therefore, we developed a Mx-Cas9-GFP system of conditionally immortalized HoxB8 MPP, which upon differentiation faithfully recapitulate DC development (Figure 2—figure supplement 1A, B; Xu et al., 2022). HoxB8 MPP were obtained from bone marrow of Mx-Cas9-GFP mice by infection with the estrogen (E2) inducible These MPP an and potential and into all DC subsets in and in et al., 2022). of gRNA lncIrf8 promoter in Mx-Cas9-GFP HoxB8 MPP and induction of by interferon generated lncIrf8 promoter KO of with were further studied and to DC differentiation (Figure Figure 2—figure supplement and Figure 2—figure supplement lncIrf8 promoter KO lncIrf8 expression during DC differentiation compared to control without deletion (Figure 2B). Surprisingly, Irf8 expression was also which points to a cross-talk of the lncIrf8 promoter element with the Irf8 lncIrf8 promoter KO also impacts DC subsets, CD11c+ pDC and cDC subsets cDC1 and cDC2 were (Figure and Figure 2—figure supplement of pDC and cDC1 were while cDC2 were (Figure 2C). lncIrf8 promoter KO cDC2 and some cells and were more compared to control without which multiple DC subsets (Figure and Figure 2—figure supplement lncIrf8 promoter KO also the differentiation of progenitors upon E2 from (Figure 1—figure supplement 2A and Figure 2—figure supplement lncIrf8 promoter KO a in cells compared to control (Figure 2—figure supplement lncIrf8 promoter KO higher of Gr1+ monocytes (Figure Figure 2—figure supplement and of all DC subsets CD11c+ pDC, cDC1, and cDC2 (Figure 2—figure supplement compared to control without deletion. The lncIrf8 promoter element is in close proximity to the cDC1 specific +32 kb enhancer previously in mice by Durai et al., and thus we generated the cDC1 specific +32 kb enhancer KO following the same as for the lncIrf8 promoter of clones with of cDC1 +32 kb enhancer were to DC differentiation and further (Figure 2—figure supplement to lncIrf8 promoter cDC1 specific +32 kb enhancer KO lncIrf8 expression and also Irf8 expression during DC differentiation (Figure 2—figure supplement The cDC1 +32 kb enhancer KO also the of cDC1 in Flt3L directed DC differentiation, while cDC2 were (Figure 2—figure supplement of pDC were also at day however, this was not These observations are in line with studies in mice that cDC1 +32 kb enhancer KO cDC1 differentiation and left pDC and cDC2 largely (Durai et al., 2019; Murakami et al., 2021). In addition, cDC1 +32 kb enhancer KO higher of Gr1+ monocytes upon spontaneous DC differentiation by of E2 (Figure 2—figure supplement and thus a as the lncIrf8 promoter KO upon spontaneous DC differentiation (Figure the novel of the lncIrf8 promoter we proceeded to investigate the impact of the lncIrf8 promoter KO on DC differentiation in in We lncIrf8 promoter KO and control HoxB8 MPP into mice (Figure supplement DC in bone marrow and spleen were by flow cytometry on day 7 and cell (Figure 3, Figure supplement 1A, In bone lncIrf8 promoter KO cells into Gr1+ and of all DC subsets were observed on day 7 for lncIrf8 promoter KO cells compared to control (Figure In of cell populations from lncIrf8 promoter KO and control were to bone including of all DC subsets for lncIrf8 promoter KO (Figure HoxB8 cells were largely at day cell (Figure and Figure Figure 3 with 1 supplement see all Download asset Open asset lncIrf8 promoter KO comprises pDC and cDC1 development in upon cell (A) Representative flow cytometry analysis of lncIrf8 promoter KO and control HoxB8 MPP in BM at day 7 cell details see Figure supplement 1A, cell populations were gated from cells and Gr1+ pDC, cDC1 and cDC2 are shown. Quantification of Gr1+ CD11c+ pDC, cDC1, and cDC2 of living single cells in BM on day 7 and cell Representative flow cytometry analysis of lncIrf8 promoter KO and control HoxB8 MPP in spleen at day 7 cell was as in (A). Quantification of Gr1+ CD11c+ pDC, cDC1 and cDC2 on day 7 and cell Data represent mean ± SD from 3 to 4 *p<0.05, **p<0.01, ***p<0.001, multiple t-tests. Data that have no difference (p>0.05) are not labeled. Thus, lncIrf8 promoter KO pDC and cDC1 development both in and in lncIrf8 acts as an indicator of Irf8 +32 kb enhancer activity in pDC of lncIrf8 promoter and thus of lncIrf8 expression pDC and cDC1 development in and in The lncIrf8 promoter is located within Irf8 +32 kb enhancer (Durai et al., 2019) and thus it was important to lncIrf8 itself a role in regulating pDC and cDC1 development. this we lncIrf8 in MPP and lncIrf8 in lncIrf8 promoter KO MPP and its impact on DC development (Figure 4 and Figure supplement 1). Figure 4 with 1 supplement see all Download asset Open asset lncIrf8 pDC and cDC1 development and B) of lncIrf8 in HoxB8 MPP and of and A polyA signal for transcription was at the 3’ end of lncIrf8 and (C) Gene expression of lncIrf8 and Irf8 in and HoxB8 MPP on day 0, 3, 5, and 7 of Flt3L directed DC differentiation Gene expression was by RT-qPCR and normalized to GAPDH. (D) Representative flow cytometry of DC subsets at day 5 and of Flt3L directed DC differentiation of and HoxB8 Quantification of pDC and cDC1 of living single cells on Flt3L directed DC differentiation day 0, 3, 5, 7, and is shown. for pDC and cDC1 was as in Figure 2—figure supplement (E) of DC subsets of (D) at day 0, 3, 5, 7, and of DC differentiation. high and low Data represent mean ± SD of at least three independent experiments with different HoxB8 MPP clones of and *p<0.05, **p<0.01, ***p<0.001, multiple t-tests. Data that have no difference (p>0.05) are not labeled. lncIrf8 was into a polyA An signal et al., 2015) was at the 3’ end of lncIrf8 to avoid transcripts than lncIrf8 Figure The vector was used as control. lncIrf8 clones were generated by (Figure supplement and to DC differentiation. As expected lncIrf8 expression was in cells compared to while there were no in Irf8 expression between the two during DC differentiation (Figure there were no in the of pDC, cDC1, and cDC2 between cells and controls (Figure and Figure supplement indicating that lncIrf8 no on Irf8 expression and DC differentiation. further this we performed a lncIrf8 in lncIrf8 promoter KO lncIrf8 was in lncIrf8 promoter KO MPP by vector and cells were to DC differentiation (Figure supplement lncIrf8 RNA was expressed and cells in response to Flt3L

We have recently shown that at late times in an SV40 infection a nucleosome containing modified histones is positioned over the enhancer region consistent with repression of early transcription and activation of late transcription. However, this nucleosome appears to slide toward the late region during virion formation consistent with a role in the activation of early transcription and repression of late transcription in a subsequent infection. Because the transcription factor SP1 plays a major role in regulating SV40 early and late transcription and its binding should be sensitive to nucleosome positioning, we measured the ability of HIS‐tagged SP1 to bind to various forms of SV40 chromatin in vitro and compared this to the presence of SP1 in SV40 chromatin derived in vivo. We found that HIS‐tagged SP1 bound well to chromatin from virions and very poorly to other forms of SV40 chromatin in vitro, while cellular SP1 was associated with minichromosomes isolated from infected cells and was not found in chromatin from virions. In order to determine whether the location of nucleosomes and associated histone modifications plays a role in the activation and continuation of early transcription during the initiation of an infection, we have mapped the location of RNAPII and nucleosomes containing a number of histone modifications in chromatin from virions and minichromosomes isolated 30 minutes and 2 hours post‐infection using ChIP‐Seq. During the first two hours of infection we found that RNAPII first associates with the miRNA site in the SV40 genome (at 30 minutes post‐infection) and then shifts to the early regulatory region around the enhancer (2 hours post‐infection). We next compared the location of RNAPII at the two time points to the location of nucleosomes carrying one of nine different specific histone modifications in the chromatin from virions, and minichromosomes isolated at these times. Chromatin from virions was used because it is the substrate for the activation of transcription, while chromatin from the minichromosomes would indicate any changes occurring as a consequence of transcriptional activation or extension. Based upon this rationale, we identified acetylated H3 as the most likely histone modification in SV40 predicting subsequent binding by RNAPII. We also identified acetylated H4 and the shifting of nucleosomes away from the RNAPII binding site as changes which result from activation of transcription. Together these results indicate that positioned nucleosomes carrying certain histone modifications may control accessibility of transcription factors and also serve as marks for either the binding of RNAPII during initiation of transcription or subsequent re‐initiation during active transcription.Support or Funding InformationNIH 1R03AI127969‐01NIH 1R03AI142011‐01

Article Figures and data Abstract Editor's evaluation Introduction Results Discussion Materials and methods Appendix 1 Data availability References Decision letter Author response Article and author information Metrics Abstract Transcription factors play a determining role in lineage commitment and cell differentiation. Interferon regulatory factor 8 (IRF8) is a lineage determining transcription factor in hematopoiesis and master regulator of dendritic cells (DC), an important immune cell for immunity and tolerance. IRF8 is prominently upregulated in DC development by autoactivation and controls both DC differentiation and function. However, it is unclear how Irf8 autoactivation is controlled and eventually limited. Here, we identified a novel long non-coding RNA transcribed from the +32 kb enhancer downstream of Irf8 transcription start site and expressed specifically in mouse plasmacytoid DC (pDC), referred to as lncIrf8. The lncIrf8 locus interacts with the lrf8 promoter and shows differential epigenetic signatures in pDC versus classical DC type 1 (cDC1). Interestingly, a sequence element of the lncIrf8 promoter, but not lncIrf8 itself, is crucial for mouse pDC and cDC1 differentiation, and this sequence element confers feedback inhibition of Irf8 expression. Taken together, in DC development Irf8 autoactivation is first initiated by flanking enhancers and then second controlled by feedback inhibition through the lncIrf8 promoter element in the +32 kb enhancer. Our work reveals a previously unrecognized negative feedback loop of Irf8 that orchestrates its own expression and thereby controls DC differentiation. Editor's evaluation Authors provide valuable evidence identifying a lncRNA transcribed specifically in the pDC subtype from the +32Kb promoter region which is also the region for the enhancer for Irf8 specifically in the cDC1 subtype. With convincing methodology, they provide in-depth analysis about the possible role of lncIrf8, and its promoter region and cross-talk with Irf8 promoter to identify that it is not the lncIRF8 itself but its promoter region that is crucial for pDC and cDC1 differentiation conferring feedback inhibition of Irf8 transcription. The work will be of interest to immunologists working on immune cell development. https://doi.org/10.7554/eLife.83342.sa0 Decision letter Reviews on Sciety eLife's review process Introduction Lineage-determining transcription factors (TF) are master regulators of gene programs that frequently initiate self-reinforcing loops by autoactivation. TF autoactivation is important for cells to pass restriction points during development (referred to as points of no return) and to enforce cellular identity. Molecular circuitries of autoactivation have been studied for several TF, such as GATA-binding factor 1 (GATA1), PU.1 (encoded by Spi1), CCAAT enhancer-binding protein α and ε (C/EBPα and ε; Graf and Enver, 2009; Loughran et al., 2020; Nishimura et al., 2000; Okuno et al., 2005; Theilgaard-Mönch et al., 2022). A further example is interferon regulatory factor 8 (IRF8), which shows autoactivation in cooperation with basic leucine zipper ATF-like transcription factor 3 (BATF3; Anderson et al., 2021; Grajales-Reyes et al., 2015). An important principle in nature is negative feedback control to avoid signal overshooting and toxicity. Negative feedback control applies also to lineage-determining TF; however, there is a paucity on our knowledge of the molecular mechanisms involved. IRF8 is a hematopoietic TF positioned at the center of the regulatory gene network for dendritic cell (DC) development (Anderson et al., 2021; Belz and Nutt, 2012; Chauvistré and Seré, 2020; Kim et al., 2020; Lin et al., 2015; Nutt and Chopin, 2020; Tamura et al., 2015; Verlander et al., 2022). IRF8 is a member of the interferon regulatory factor (IRF) family of TF. Initially members of this TF family were found to mediate the induction of interferon induced genes, but are now known to serve diverse functions in regulating the immune system (Honda and Taniguchi, 2006; Tamura et al., 2008). Irf8 knockout mice show abnormal development of classical DC type 1 (cDC1) and plasmacytoid DC (pDC) (Durai et al., 2019; Schiavoni et al., 2002; Sichien et al., 2016; Tsujimura et al., 2003). Irf8 is prominently upregulated during DC development by autoactivation (Grajales-Reyes et al., 2015; Lin et al., 2015), yet how Irf8 autoactivation is controlled and eventually limited, and the epigenetic mechanisms involved is largely unknown. Irf8 expression in hematopoietic cells is induced and maintained by enhancers located at –50 kb,+32 kb,+41 kb and +56 kb relative to Irf8 transcription start site (TSS) (Anderson et al., 2021; Bagadia et al., 2019; Durai et al., 2019; Grajales-Reyes et al., 2015; Murakami et al., 2021; Schönheit et al., 2013). Enhancers are cis-regulatory sequences with multiple TF binding sites that cooperatively bind TF and thereby activate transcription, as demonstrated by many studies including our work (Davidson et al., 1986; Long et al., 2016; Wildeman et al., 1986; Zenke et al., 1986). Enhancers regulate complex gene networks and can also produce non-coding RNA, referred to as enhancer RNA (eRNA). eRNA serve as an indicator for enhancer activity and some eRNA have an activity on their own and act in cis or trans to regulate cell fate decisions (Sartorelli and Lauberth, 2020; Statello et al., 2021). Enhancer-associated long non-coding RNA (lncRNA) represent a class of lncRNA transcribed from active enhancers. Thus, eRNA and enhancer-associated lncRNA provide opportunities to detect enhancer activity and to investigate enhancer function. DC are highly specialized immune cells that play a critical role in regulating innate and adaptive immune responses (Cabeza-Cabrerizo et al., 2021). DC develop from hematopoietic stem cells (HSC) via successive steps of lineage commitment and differentiation. More specifically, HSC develop into multipotent progenitors (MPP) that are committed to DC restricted common DC progenitors (CDP) and differentiate into classic DC (cDC) type 1 and type 2 (cDC1 and cDC2, respectively) and pDC (Anderson et al., 2021; Cabeza-Cabrerizo et al., 2021; Ginhoux et al., 2022; Nutt and Chopin, 2020; Rodrigues and Tussiwand, 2020). pDC were recently also shown to develop from lymphoid progenitors (Dress et al., 2019; Rodrigues et al., 2018; Rodrigues and Tussiwand, 2020). Differential expression of Irf8 regulates DC and monocyte specification in a dose-dependent manner (Cytlak et al., 2020; Murakami et al., 2021). Irf8 expression starts at the CDP stage, and is high in pDC and cDC1, which is attributed to the autoactivation of Irf8 during DC subsets specification (Grajales-Reyes et al., 2015; Lin et al., 2015). Interestingly, IRF8 can act as a transcriptional activator or repressor in hematopoiesis by interacting with different partner TF and binding to specific DNA sequences (Tamura et al., 2015). As an activator, IRF8 binds to its own promoter in DC differentiation, which is considered as the autoactivation capacity of Irf8 (Grajales-Reyes et al., 2015; Lin et al., 2015). For instance, IRF8 interacts with partner TF, such as PU.1, to initiate Irf8 autoactivation at the CDP stage (Grajales-Reyes et al., 2015). Inversely, IRF8 inhibits C/EBPα activity in neutrophil differentiation (Kurotaki et al., 2014). IRF8 also represses C/EBPβ to generate and maintain DC lineage-specific enhancer landscapes (Bornstein et al., 2014). In addition, IRF8 is important for the Myc-Mycl transition in DC differentiation (Anderson lll et al., 2021). IRF8 represses Myc expression in progenitors, while IRF8 at high levels interacts with PU.1 and drives Mycl expression (Anderson lll et al., 2021). All this emphasizes the central position of IRF8 in coordinating the gene network that regulates DC differentiation and function. During DC differentiation, the Irf8 gene locus shows high epigenetic dynamics, including histone modifications and TF binding identified by ChIP-seq (Chauvistré and Seré, 2020; Durai et al., 2019; Grajales-Reyes et al., 2015; Lin et al., 2015), chromatin accessibility measured by ATAC-seq (Kurotaki et al., 2019; Li et al., 2019), and three-dimensional chromatin structure remodeling determined by chromosome conformation capture (3 C) (Kurotaki et al., 2022; Schönheit et al., 2013). All this emphasizes the impact of epigenetic regulators on Irf8 gene activity in DC differentiation. Notably, Irf8 is flanked by multiple enhancers at –50 kb,+32 kb,+41 kb, and +56 kb that regulate Irf8 expression in hematopoietic cells (Anderson et al., 2021; Murakami et al., 2021). These four enhancers were found to be driven by PU.1, BATF3, E proteins and Runt-related transcription factor (RUNX)-core binding factor beta (CBFβ) (RUNX-CBFβ), respectively (Bagadia et al., 2019; Durai et al., 2019; Grajales-Reyes et al., 2015; Murakami et al., 2021; Schönheit et al., 2013). Chromatin conformation, particularly enhancer promoter interactions, provides a platform for TF-driven gene regulation and serves as a driving force for cell-fate determinations (Misteli and Finn, 2021; Oudelaar and Higgs, 2021; Stadhouders et al., 2019). Schönheit et al. demonstrated Irf8 promoter interactions with its upstream enhancers by quantitative 3 C (Schönheit et al., 2013). In this study PU.1 was found to regulate chromatin remodeling between the –50 kb enhancer and the Irf8 promoter in myeloid differentiation. In a recent study Kurotaki et al., 2022 determined the higher-order chromatin structure in DC progenitors, cDC1 and cDC2 on a genome-wide scale by Hi-C. In this study, reorganization of chromatin conformation at DC-specific gene loci was observed during cDC differentiation, and IRF8 was found to promote chromatin activation in DC progenitors leading to cDC lineage-specific gene expression. However, high resolution maps of the physical chromatin interactions of the Irf8 promoter with upstream and downstream enhancers in the full complement of DC subsets, including pDC, are required for understanding Irf8 regulation during DC differentiation. Frequently, chromatin data, including ATAC-seq and/or ChIP-seq data, are used to identify regulatory elements of gene transcription. Here we embarked on a different approach and searched for lncRNA, which by themselves might have regulatory functions or are indicative of enhancer activity. We identified a novel lncRNA transcribed from the Irf8 +32 kb enhancer, which is specifically expressed in pDC, referred to as lncIrf8. We found that the lncIrf8 promoter element but not lncIrf8 itself impacts pDC and cDC1 development. Thus, lncIrf8 acts as an indicator for the Irf8 +32 kb enhancer activity. Importantly, our study revealed a previously unrecognized negative feedback loop of Irf8 in DC differentiation. Irf8 first activates its expression by autoactivation via the +32 kb enhancer and second limits its own expression through the lncIrf8 promoter element in the +32 kb enhancer. Results lncIrf8 marks a pDC-specific Irf8 enhancer element Irf8 expression in DC development is subject to complex epigenetic regulation. Here, we used an integrated approach with RNA-seq, ATAC-seq, ChIP-seq and Capture-C to track the dynamics of gene expression, histone modification and chromatin conformation in the sequel MPP, CDP, pDC, cDC1, and cDC2 (Figure 1, Figure 1—figure supplements 1 and 2). Figure 1 with 3 supplements see all Download asset Open asset Irf8 epigenetic signatures and promoter-enhancer interaction maps during DC differentiation. (A) Gene expression and epigenetic signatures of Irf8 downstream region in MPP, CDP, pDC, all cDC, cDC1, and cDC2 are visualized by IGV browser. Gene expression was measured by RNA-seq, chromatin accessibility by ATAC-seq, H3K27ac and IRF8 binding by ChIP-seq. Positions of Irf8 3' end, Irf8 enhancers, pDC specific lncIrf8 and cDC1 specific Tcons_00190258 lncRNA are indicated. For RNA-seq - and +strands are shown. Scale bar: 5 kb. (B) Physical interactions of Irf8 promoter with flanking sequences in MPP, CDP, pDC, cDC1, and cDC2 by nuclear-titrated (NuTi) Capture-C (turquoise), and CTCF binding by ChIP-seq in DC (Garber et al., 2012). Mean numbers of unique interactions normalized to a 300 kb region around the Irf8 promoter viewpoint (green triangle) and scaled by a factor of 1,000,000 are shown (n=2). The orientations of CTCF binding are indicated with blue and red arrows. Tcons_00190258 refers to the cDC1-specific lncRNA shown in (A). Scale bar: 100 kb. (C) Comparations of the chromatin interactions with Irf8 promoter in pDC, cDC1 and cDC2. Differential tracks were created by subtraction of the mean normalized tracks of (B). Pairwise comparisons are shown and color coded. Turquoise and orange tracks represent specific interactions with the Irf8 promoter in the indicated cell types. Scale bar: 100 kb. Purple bars and lines indicate the position of flanking enhancers relative to Irf8 TSS. The purple bars from left to right represent –50 kb, –34 kb, –26 kb, –16 kb,+27 kb,+32 kb,+38 kb,+41 kb,+47 kb,+56 kb and +62 kb enhancer, respectively (panels B and C). Irf8 +32 kb enhancer is highlighted by purple box. We performed de novo transcript assembly of the RNA-seq data and detected two previously unknown transcripts without coding potential downstream of Irf8: a pDC specific lncRNA (Tcons_00190250) in the following referred to as lncIrf8 and a cDC1 specific lncRNA (Tcons_00190258; Figure 1A and Figure 1—figure supplement 1). lncIrf8 and Tcons_00190258 show the same expression pattern in pDC and cDC1, respectively, in BM and spleen (Figure 1—figure supplement 3), as revealed by reanalyzing scRNA-seq and bulk RNA-seq data (Pang et al., 2022; Rodrigues et al., 2018). lncIrf8 is transcribed within an enhancer region located 32 kb downstream of the Irf8 TSS labeled by H3K27ac and H3K4me1 and occupied by DC differentiation-associated TF, such as IRF8 and PU.1 (Figure 1A and Figure 1—figure supplement 1). This region is largely devoid of H3K9me3, a chromatin modification frequently associated with heterochromatin, indicating an open chromatin configureuration in DC (Figure 1—figure supplement 1). In addition, sequences of this region have been implicated in DC development and referred to as +32 kb enhancer (Durai et al., 2019). Thus, we proceeded to study lncIrf8 in detail. ATAC-seq analysis revealed further details of the lncIrf8 region in CDP, pDC, cDC1 and cDC2 (Figure 1A, Figure 2A and Figure 1—figure supplement 1). In cDC1 the prominent ATAC-seq and IRF8 peaks mark the cDC1 specific +32 kb enhancer (Durai et al., 2019). In pDC the ATAC-seq peak is smaller and shifted further towards downstream but aligns well with the valley in the prominent H3K27ac peak. This ATAC-seq peak marks the lncIrf8 promoter and aligns with p300 (Durai et al., 2019) and H3K4me3 (Figure 2A and Figure 1—figure supplement 1). All this indicates that this chromatin region is open and transcriptionally active in pDC, enabling lncIrf8 transcription. Figure 2 with 4 supplements see all Download asset Open asset IncIrf8 promoter KO compromises pDC and cDC1 development in vitro. (A) Genomic anatomy of lncIrf8 locus determined by 3' and 5' RACE PCR. Blue box, exon 2 and 3 (48 bp and 468 bp, respectively). The 1010 bp intron and polyA tail are shown. Data of RNA-seq, ATAC-seq, ChIP-seq of H3K27ac (enhancer mark) and H3K4me3 (active promoter mark, near TSS) are visualized by IGV browser for the indicated cell populations (pDC, all cDC, cDC1 and cDC2). Grey box, lncIrf8 promoter KO region; open box, cDC1 specific +32 kb enhancer by Durai et al., 2019. Irf8 +32 kb enhancer based on the H3K27ac enhancer mark is indicated with a purple line. Scale bar: 1 kb. (B) Gene expression of lncIrf8 and Irf8 in lncIrf8 promoter KO and control at day 0, 5, and 7 of Flt3L directed DC differentiation. Gene expression was determined by RT-qPCR and normalized to GAPDH. n=4. (C) Representative flow cytometry analysis of Flt3L directed DC differentiation of lncIrf8 promoter KO HoxB8 MPP and control (Lutz et al., 2022; Xu et al., 2022). pDC, all cDC, cDC1, and cDC2 were gated as in Figure 2—figure supplement 1E and are shown. Bar diagrams depict quantification of pDC, cDC1 and cDC2 normalized to living single cells on DC differentiation day 0, 3, 5, 7, and 9. n=6–7. (D) Representative phase-contrast microscopy images of lncIrf8 promoter KO HoxB8 MPP and control on day 7 of Flt3L directed DC differentiation. Scale bar: 200 μm. (E) Representative flow cytometry analysis of spontaneous DC differentiation of lncIrf8 promoter KO HoxB8 MPP and control with growth factors but without E2 (Lutz et al., 2022; Xu et al., 2022) at day 8. Gr1+ monocytes and CD11c+ DC are shown. Quantification of Gr1+ monocytes of living single cells on day 3, 6, 8, and 10 of spontaneous DC differentiation. n=6, lncIrf8 promoter KO; n=4, control. Empty gRNA vector or non-targeting gRNA vector HoxB8 MPP were used as controls. Data represent mean ± SD of at least three independent experiments with different HoxB8 MPP clones of lncIrf8 promoter KO and control without deletion. *p<0.05, **p<0.01, ***p<0.001, multiple t-tests. Data that have no difference (p>0.05) are not labeled. Next, we determined the chromatin conformation of the Irf8 locus and the lncIrf8 region. We generated interaction profiles by nuclear-titrated (NuTi) Capture-C in MPP, CDP, pDC, cDC1, and cDC2 (Figure 1—figure supplement 2A and B) using Irf8 promoter as viewpoint. The Irf8 promoter shows multiple interactions with regions spanning up to ~100 kb upstream and downstream of Irf8 (Figure 1B and Figure 1—figure supplement 1). In pDC, the Irf8 promoter interactions are stronger with the upstream sequences than with downstream sequences (Figure 1C and Figure 1—figure supplement 2C). In cDC1 Irf8 promoter interactions are more confined to the regions downstream of Irf8 compared to MPP, CDP and pDC (Figure 1C and Figure 1—figure supplement 2C). This suggests that upstream and downstream sequences of Irf8 gene are involved in differentially regulating Irf8 expression and controlling the development of pDC and cDC1, respectively. The CCCTC-binding factor (CTCF) is important for regulation of chromatin conformation through loop extrusion (Sanborn et al., 2015) and we therefore visualized CTCF binding sites in the Irf8 locus in DC (Garber et al., 2012). Interestingly, most of the Irf8 flanking enhancers (Durai et al., 2019; Grajales-Reyes et al., 2015; Murakami et al., 2021; Schönheit et al., 2013) are located within convergent CTCF binding sites upstream and downstream of the Irf8 gene (Figure 1B and Figure 1—figure supplement 1). There are also multiple interactions within this region without convergent CTCF binding sites, suggesting interactions with Irf8 promoter in a CTCF independent manner, such as by TF binding, active histone modifications and gene transcription (Figure 1B and Figure 1—figure supplement 1; Owens et al., 2022). Surprisingly, in pDC H3K27ac at the lncIrf8 promoter is high, but this locus shows less interactions with the Irf8 promoter in pDC compared to CDP, cDC1 and cDC2 (Figure 1C and Figure 1—figure supplement 2C). In addition, in pDC the IRF8 protein occupancy at the lncIrf8 promoter is low and much higher in cDC (Figure 1A and Figure 1—figure supplement 1; Durai et al., 2019; Grajales-Reyes et al., 2015). These observations warrant further studies and we thus proceeded to investigate the lncIrf8 locus in detail. lncIrf8 promoter KO compromises pDC and cDC1 development First, we annotated lncIrf8. Our de-novo transcript assembly of RNA-seq data revealed different isoforms of lncIrf8, with the most prominent isoform comprising exon 2 and 3 (Figure 1A, Figure 2A and Figure 1—figure supplement 1). Additionally, 3' end and 5' end RACE PCR confirmed the anatomy of this lncIrf8 isoform: two exons, one intron, and a polyA tail (Figure 2A). As expected lncIrf8 is not conserved across species (data not shown), which is in line with the general characteristics of lncRNA. Then second, we deleted 300 bp in the lncIrf8 promoter by CRISPR/Cas9 editing in conditionally immortalized HoxB8 MPP (Figure 2A and Figure 2—figure supplement 1A–D). The lncIrf8 promoter is located in the Irf8 +32 kb enhancer region and is in close proximity to the cDC1 specific +32 kb enhancer (Durai et al., 2019; Figure 2A and Figure 2—figure supplement 2). The 300 bp deletion comprises the H3K4me3 promoter mark and is confined to open chromatin identified by ATAC-seq and positioned in the valley of the H3K27ac mark (Figure 2A). Additionally, it contains binding sites for IRF8, PU.1, and BATF3 TF, which are important for DC development (Figure 2A and Figure 2—figure supplement 2B). Generation of a precise deletion requires clonal cell populations, which is hardly achieved in somatic cells due to their limited lifespan. Therefore, we developed a Mx-Cas9-GFP system of conditionally immortalized HoxB8 MPP, which upon differentiation faithfully recapitulate DC development (Figure 2—figure supplement 1A, B; Xu et al., 2022). HoxB8 MPP were obtained from bone marrow of Mx-Cas9-GFP mice by infection with the estrogen (E2) inducible HoxB8-ER. These MPP an and potential and into all DC subsets in and in et al., 2022). of gRNA lncIrf8 promoter in Mx-Cas9-GFP HoxB8 MPP and induction of by interferon generated lncIrf8 promoter KO of with were further studied and to DC differentiation (Figure Figure 2—figure supplement and Figure 2—figure supplement lncIrf8 promoter KO lncIrf8 expression during DC differentiation compared to control without deletion (Figure 2B). Surprisingly, Irf8 expression was also which points to a cross-talk of the lncIrf8 promoter element with the Irf8 lncIrf8 promoter KO also impacts DC subsets, CD11c+ pDC and cDC subsets cDC1 and cDC2 were (Figure and Figure 2—figure supplement of pDC and cDC1 were while cDC2 were (Figure 2C). lncIrf8 promoter KO cDC2 and some cells and were more compared to control without which multiple DC subsets (Figure and Figure 2—figure supplement lncIrf8 promoter KO also the differentiation of progenitors upon E2 from (Figure 1—figure supplement 2A and Figure 2—figure supplement lncIrf8 promoter KO a in cells compared to control (Figure 2—figure supplement lncIrf8 promoter KO higher of Gr1+ monocytes (Figure Figure 2—figure supplement and of all DC subsets CD11c+ pDC, cDC1, and cDC2 (Figure 2—figure supplement compared to control without deletion. The lncIrf8 promoter element is in close proximity to the cDC1 specific +32 kb enhancer previously in mice by Durai et al., and thus we generated the cDC1 specific +32 kb enhancer KO following the same as for the lncIrf8 promoter of clones with of cDC1 +32 kb enhancer were to DC differentiation and further (Figure 2—figure supplement to lncIrf8 promoter cDC1 specific +32 kb enhancer KO lncIrf8 expression and also Irf8 expression during DC differentiation (Figure 2—figure supplement The cDC1 +32 kb enhancer KO also the of cDC1 in Flt3L directed DC differentiation, while cDC2 were (Figure 2—figure supplement of pDC were also at day however, this was not These observations are in line with studies in mice that cDC1 +32 kb enhancer KO cDC1 differentiation and left pDC and cDC2 largely (Durai et al., 2019; Murakami et al., 2021). In addition, cDC1 +32 kb enhancer KO higher of Gr1+ monocytes upon spontaneous DC differentiation by of E2 (Figure 2—figure supplement and thus a as the lncIrf8 promoter KO upon spontaneous DC differentiation (Figure the novel of the lncIrf8 promoter we proceeded to investigate the impact of the lncIrf8 promoter KO on DC differentiation in in We lncIrf8 promoter KO and control HoxB8 MPP into mice (Figure supplement DC in bone marrow and spleen were by flow cytometry on day 7 and cell (Figure 3, Figure supplement 1A, In bone lncIrf8 promoter KO cells into Gr1+ and of all DC subsets were observed on day 7 for lncIrf8 promoter KO cells compared to control (Figure In of cell populations from lncIrf8 promoter KO and control were to bone including of all DC subsets for lncIrf8 promoter KO (Figure HoxB8 cells were largely at day cell (Figure and Figure Figure 3 with 1 supplement see all Download asset Open asset lncIrf8 promoter KO comprises pDC and cDC1 development in upon cell (A) Representative flow cytometry analysis of lncIrf8 promoter KO and control HoxB8 MPP in BM at day 7 cell details see Figure supplement 1A, cell populations were gated from cells and Gr1+ pDC, cDC1 and cDC2 are shown. Quantification of Gr1+ CD11c+ pDC, cDC1, and cDC2 of living single cells in BM on day 7 and cell Representative flow cytometry analysis of lncIrf8 promoter KO and control HoxB8 MPP in spleen at day 7 cell was as in (A). Quantification of Gr1+ CD11c+ pDC, cDC1 and cDC2 on day 7 and cell Data represent mean ± SD from 3 to 4 *p<0.05, **p<0.01, ***p<0.001, multiple t-tests. Data that have no difference (p>0.05) are not labeled. Thus, lncIrf8 promoter KO pDC and cDC1 development both in and in lncIrf8 acts as an indicator of Irf8 +32 kb enhancer activity in pDC of lncIrf8 promoter and thus of lncIrf8 expression pDC and cDC1 development in and in The lncIrf8 promoter is located within Irf8 +32 kb enhancer (Durai et al., 2019) and thus it was important to lncIrf8 itself a role in regulating pDC and cDC1 development. this we lncIrf8 in MPP and lncIrf8 in lncIrf8 promoter KO MPP and its impact on DC development (Figure 4 and Figure supplement 1). Figure 4 with 1 supplement see all Download asset Open asset lncIrf8 pDC and cDC1 development and B) of lncIrf8 in HoxB8 MPP and of and A polyA signal for transcription was at the 3' end of lncIrf8 and (C) Gene expression of lncIrf8 and Irf8 in and HoxB8 MPP on day 0, 3, 5, and 7 of Flt3L directed DC differentiation Gene expression was by RT-qPCR and normalized to GAPDH. (D) Representative flow cytometry of DC subsets at day 5 and of Flt3L directed DC differentiation of and HoxB8 Quantification of pDC and cDC1 of living single cells on Flt3L directed DC differentiation day 0, 3, 5, 7, and is shown. for pDC and cDC1 was as in Figure 2—figure supplement (E) of DC subsets of (D) at day 0, 3, 5, 7, and of DC differentiation. high and low Data represent mean ± SD of at least three independent experiments with different HoxB8 MPP clones of and *p<0.05, **p<0.01, ***p<0.001, multiple t-tests. Data that have no difference (p>0.05) are not labeled. lncIrf8 was into a polyA An signal et al., 2015) was at the 3' end of lncIrf8 to avoid transcripts than lncIrf8 Figure The vector was used as control. lncIrf8 clones were generated by (Figure supplement and to DC differentiation. As expected lncIrf8 expression was in cells compared to while there were no in Irf8 expression between the two during DC differentiation (Figure there were no in the of pDC, cDC1, and cDC2 between cells and controls (Figure and Figure supplement indicating that lncIrf8 no on Irf8 expression and DC differentiation. further this we performed a lncIrf8 in lncIrf8 promoter KO lncIrf8 was in lncIrf8 promoter KO MPP by vector and cells were to DC differentiation (Figure supplement lncIrf8 RNA was expressed and cells in response to Flt3L