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| <?xml version="1.0" encoding="UTF-8"?><!DOCTYPE article PUBLIC "-//NLM//DTD JATS (Z39.96) Journal Archiving and Interchange DTD v1.1d1 20130915//EN" "JATS-archivearticle1.dtd"><article article-type="research-article" dtd-version="1.1d1" xmlns:xlink="http://www.w3.org/1999/xlink"><front><journal-meta><journal-id journal-id-type="nlm-ta">elife</journal-id><journal-id journal-id-type="hwp">eLife</journal-id><journal-id journal-id-type="publisher-id">eLife</journal-id><journal-title-group><journal-title>eLife</journal-title></journal-title-group><issn publication-format="electronic">2050-084X</issn><publisher><publisher-name>eLife Sciences Publications, Ltd</publisher-name></publisher></journal-meta><article-meta><article-id pub-id-type="publisher-id">02557</article-id><article-id pub-id-type="doi">10.7554/eLife.02557</article-id><article-categories><subj-group subj-group-type="display-channel"><subject>Research article</subject></subj-group><subj-group subj-group-type="heading"><subject>Genes and chromosomes</subject></subj-group></article-categories><title-group><article-title>Temporal dynamics and developmental memory of 3D chromatin architecture at <italic>Hox</italic> gene loci</article-title></title-group><contrib-group><contrib contrib-type="author" id="author-11455"><name><surname>Noordermeer</surname><given-names>Daan</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="con1"/><xref ref-type="fn" rid="conf1"/><xref ref-type="other" rid="dataro1"/></contrib><contrib contrib-type="author" id="author-11456"><name><surname>Leleu</surname><given-names>Marion</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="con2"/><xref ref-type="fn" rid="conf1"/><xref ref-type="other" rid="dataro1"/></contrib><contrib contrib-type="author" id="author-11457"><name><surname>Schorderet</surname><given-names>Patrick</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="aff" rid="aff2"/><xref ref-type="fn" rid="con3"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-11458"><name><surname>Joye</surname><given-names>Elisabeth</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="con4"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-11459"><name><surname>Chabaud</surname><given-names>Fabienne</given-names></name><xref ref-type="aff" rid="aff3"/><xref ref-type="fn" rid="con5"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" corresp="yes" id="author-11383"><name><surname>Duboule</surname><given-names>Denis</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="aff" rid="aff3"/><xref ref-type="corresp" rid="cor1">*</xref><xref ref-type="other" rid="par-1"/><xref ref-type="other" rid="par-2"/><xref ref-type="fn" rid="con6"/><xref ref-type="fn" rid="conf1"/><xref ref-type="other" rid="dataro1"/></contrib><aff id="aff1"><institution content-type="dept">School of Life Sciences</institution>, <institution>Ecole Polytechnique Fédérale de Lausanne</institution>, <addr-line><named-content content-type="city">Lausanne</named-content></addr-line>, <country>Switzerland</country></aff><aff id="aff2"><institution content-type="dept">Department of Molecular Biology</institution>, <institution>Harvard University</institution>, <addr-line><named-content content-type="city">Boston</named-content></addr-line>, <country>United States</country></aff><aff id="aff3"><institution content-type="dept">Department of Genetics and Evolution</institution>, <institution>University of Geneva</institution>, <addr-line><named-content content-type="city">Geneva</named-content></addr-line>, <country>Switzerland</country></aff></contrib-group><contrib-group content-type="section"><contrib contrib-type="editor"><name><surname>Krumlauf</surname><given-names>Robb</given-names></name><role>Reviewing editor</role><aff><institution>Stowers Institute for Medical Research</institution>, <country>United States</country></aff></contrib></contrib-group><author-notes><corresp id="cor1"><label>*</label>For correspondence: <email>denis.duboule@epfl.ch</email></corresp></author-notes><pub-date date-type="pub" publication-format="electronic"><day>29</day><month>04</month><year>2014</year></pub-date><pub-date pub-type="collection"><year>2014</year></pub-date><volume>3</volume><elocation-id>e02557</elocation-id><history><date date-type="received"><day>16</day><month>02</month><year>2014</year></date><date date-type="accepted"><day>07</day><month>04</month><year>2014</year></date></history><permissions><copyright-statement>© 2014, Noordermeer et al</copyright-statement><copyright-year>2014</copyright-year><copyright-holder>Noordermeer et al</copyright-holder><license xlink:href="http://creativecommons.org/licenses/by/3.0/"><license-p>This article is distributed under the terms of the <ext-link ext-link-type="uri" xlink:href="http://creativecommons.org/licenses/by/3.0/">Creative Commons Attribution License</ext-link>, which permits unrestricted use and redistribution provided that the original author and source are credited.</license-p></license></permissions><self-uri content-type="pdf" xlink:href="elife02557.pdf"/><abstract><object-id pub-id-type="doi">10.7554/eLife.02557.001</object-id><p><italic>Hox</italic> genes are essential regulators of embryonic development. Their step-wise transcriptional activation follows their genomic topology and the various states of activation are subsequently memorized into domains of progressively overlapping gene products. We have analyzed the 3D chromatin organization of <italic>Hox</italic> clusters during their early activation in vivo, using high-resolution circular chromosome conformation capture. Initially, <italic>Hox</italic> clusters are organized as single chromatin compartments containing all genes and bivalent chromatin marks. Transcriptional activation is associated with a dynamic bi-modal 3D organization, whereby the genes switch autonomously from an inactive to an active compartment. These local 3D dynamics occur within a framework of constitutive interactions within the surrounding Topological Associated Domains, indicating that this regulation process is mostly cluster intrinsic. The step-wise progression in time is fixed at various body levels and thus can account for the chromatin architectures previously described at a later stage for different anterior to posterior levels.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.001">http://dx.doi.org/10.7554/eLife.02557.001</ext-link></p></abstract><abstract abstract-type="executive-summary"><object-id pub-id-type="doi">10.7554/eLife.02557.002</object-id><title>eLife digest</title><p>Most animals are symmetrical about an imaginary line that runs from the head to the tail. A family of genes called the <italic>Hox</italic> family ensures that the cells in an animal embryo develop into the correct body parts along this head-to-tail axis. <italic>Hox</italic> genes—which are found in animals as diverse as flies and humans—are often clustered on the chromosomes, and their order within a cluster affects when and where each <italic>Hox</italic> gene is ‘switched on’.</p><p>In mammals, <italic>Hox</italic> genes at one end of a cluster are switched on first and along almost the entire length of the embryo. <italic>Hox</italic> genes near the other end of the cluster are expressed later and only towards the hind end of the animal. And <italic>Hox</italic> genes at the furthest end of the cluster are expressed last and in the very tip of the developing tail. The time when a <italic>Hox</italic> gene is expressed depends largely on its relative position within the gene cluster. However, it is not clear how the ordering of the genes within a cluster is translated into a schedule whereby the genes are sequentially switched on during development.</p><p>Much of the DNA in a chromosome is wrapped around proteins to form a structure called chromatin; chromatin is normally tightly packed, but ‘unpacking’ it allows the genes to be accessed and switched on. Now, Noordermeer et al. have used a technique called ‘circular chromosome conformation capture’ to follow how the packing of the chromosomes that carry the <italic>Hox</italic> gene clusters changes during embryonic development. Harvesting cells from mouse embryos of different ages, and cross-linking the DNA to the proteins, allowed those genes that are packed in the chromatin to be distinguished from those that have been unpacked and activated.</p><p>When the embryo is still just a ball of almost identical cells, all the <italic>Hox</italic> genes are switched off and packed into inactive chromatin. However, Noordermeer et al. found that, as the embryo develops and when each <italic>Hox</italic> gene is switched on in turn, the relevant region of DNA is also unpacked and moved into more active chromatin. This mechanism likely prevents <italic>Hox</italic> genes that direct the development of the hind end of the mouse from being switched on too early, and hence it avoids body parts being misidentified and developing incorrectly. Further, the patterns of active chromatin vs inactive chromatin can be fixed at each section along head-to-tail axis, such that it will be memorized in all daughter cells produced subsequently from each particular body section.</p><p>Future challenges will be to uncover the trigger behind the step-wise transition of every <italic>Hox</italic> gene from inactive chromatin to active chromatin, and to crack the underlying ‘clock’ that controls the timing of this process.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.002">http://dx.doi.org/10.7554/eLife.02557.002</ext-link></p></abstract><kwd-group kwd-group-type="author-keywords"><title>Author keywords</title><kwd>chromatin domains</kwd><kwd>gene regulation</kwd><kwd>topological domains</kwd><kwd>Hox gene regulation</kwd></kwd-group><kwd-group kwd-group-type="research-organism"><title>Research organism</title><kwd>mouse</kwd></kwd-group><funding-group><award-group id="par-1"><funding-source><institution-wrap><institution>Swiss National Research Foundation</institution></institution-wrap></funding-source><award-id>310030B_138662</award-id><principal-award-recipient><name><surname>Duboule</surname><given-names>Denis</given-names></name></principal-award-recipient></award-group><award-group id="par-2"><funding-source><institution-wrap><institution-id institution-id-type="FundRef">http://dx.doi.org/10.13039/501100000781</institution-id><institution>European Research Council</institution></institution-wrap></funding-source><award-id>232790</award-id><principal-award-recipient><name><surname>Duboule</surname><given-names>Denis</given-names></name></principal-award-recipient></award-group><funding-statement>The funder had no role in study design, data collection and interpretation, or the decision to submit the work for publication.</funding-statement></funding-group><custom-meta-group><custom-meta><meta-name>elife-xml-version</meta-name><meta-value>2</meta-value></custom-meta><custom-meta specific-use="meta-only"><meta-name>Author impact statement</meta-name><meta-value><italic>Hox</italic> genes are activated sequentially and, at the same time, undergo a transition from an inactive to an active chromatin compartment, most likely to prevent posterior genes being activated too early.</meta-value></custom-meta></custom-meta-group></article-meta></front><body><sec id="s1" sec-type="intro"><title>Introduction</title><p>Mammalian <italic>Hox</italic> genes encode proteins that are essential for patterning along the rostral-to-caudal body axis of the developing embryo (<xref ref-type="bibr" rid="bib15">Duboule and Morata, 1994</xref>; <xref ref-type="bibr" rid="bib28">Krumlauf, 1994</xref>). Mouse and human <italic>Hox</italic> genes are organized in four genomic clusters (<italic>HoxA</italic> to <italic>HoxD</italic>), where the relative position of the genes strongly impacts upon their patterns of expression. This structure-function relationship was initially described in <italic>Drosophila</italic> (<xref ref-type="bibr" rid="bib29">Lewis, 1978</xref>) and further extended to vertebrates (<xref ref-type="bibr" rid="bib22">Gaunt et al., 1988</xref>; <xref ref-type="bibr" rid="bib14">Duboule and Dolle, 1989</xref>; <xref ref-type="bibr" rid="bib25">Graham et al., 1989</xref>), where an additional correspondence exists between gene position and the timing of transcriptional activation (‘temporal colinearity’, <xref ref-type="bibr" rid="bib26">Izpisua-Belmonte et al., 1991</xref>; <xref ref-type="bibr" rid="bib9">Deschamps and van Nes, 2005</xref>).</p><p>In murine embryos, transcription of <italic>Hox</italic> genes can be divided in several phases and is first detected at around embryonic day 7 (E7) at the most posterior aspect of the primitive streak region (<xref ref-type="bibr" rid="bib10">Deschamps and Wijgerde, 1993</xref>; <xref ref-type="bibr" rid="bib19">Forlani et al., 2003</xref>). Over time, <italic>Hox</italic> genes are sequentially activated following their chromosomal order and transcripts encoded by the last <italic>Hox</italic> group 13 genes can be detected at around E9, that is two days after the onset of activation (<xref ref-type="bibr" rid="bib8">Deschamps et al., 1999</xref>; <xref ref-type="bibr" rid="bib27">Kmita and Duboule, 2003</xref>; <xref ref-type="bibr" rid="bib9">Deschamps and van Nes, 2005</xref>). This transcriptional progression (the ‘<italic>Hox</italic> clock’, <xref ref-type="bibr" rid="bib12">Duboule, 1994</xref>) thus extends over several days. In the pre-somitic mesoderm (PSM), this sequential activation needs to be coordinated with the time-sequenced production of body segments (the ‘segmentation clock’, <xref ref-type="bibr" rid="bib42">Pourquie, 2003</xref>), such that newly produced somites acquire distinct genetic identifiers (<xref ref-type="bibr" rid="bib16">Dubrulle et al., 2001</xref>; <xref ref-type="bibr" rid="bib59">Zakany et al., 2001</xref>). Next, the various states of <italic>Hox</italic> gene activity are fine-tuned and memorized, ultimately leading to domains along the rostral to caudal axis where partially overlapping sets of HOX products can be observed (‘spatial colinearity’). As a result, genes located at 3′positions (e.g., groups 3, 4) are transcribed almost along the entire embryonic axis, including the lateral plate mesoderm, paraxial mesoderm and neural tube, whereas the 5′-located group 10 or 11 are active in the posterior trunk and group 13 in the tip of the tail bud only (<xref ref-type="bibr" rid="bib8">Deschamps et al., 1999</xref>; <xref ref-type="bibr" rid="bib27">Kmita and Duboule, 2003</xref>; <xref ref-type="bibr" rid="bib9">Deschamps and van Nes, 2005</xref>).</p><p>While both temporal and spatial colinear processes likely reflect one and the same organizational principle, they are nevertheless implemented with distinctive features. Spatial colinearity could be recapitulated by several single-gene transgenes (e.g., [<xref ref-type="bibr" rid="bib43">Puschel et al., 1991</xref>; <xref ref-type="bibr" rid="bib56">Whiting et al., 1991</xref>]), yet not in all instances (<xref ref-type="bibr" rid="bib50">Tschopp et al., 2012</xref>). Indeed, a systematic analysis of modified <italic>HoxD</italic> clusters <italic>in vivo</italic> revealed that, at a late stage, the sustained transcription of these genes at the correct body level primarily relies upon local regulatory elements (<xref ref-type="bibr" rid="bib53">Tschopp et al., 2009</xref>), which are present in transgenic constructs. In contrast, the precise timing of <italic>Hoxd</italic> gene activation depends on the integrity of the full cluster, a genomic situation observed thus far in all animals developing following a temporal rostral to caudal progressive strategy during their early development (<xref ref-type="bibr" rid="bib12">Duboule, 1994</xref>). The genomic clustering of <italic>Hox</italic> genes is thus considered as an essential feature for temporal colinearity to properly process, whereas it may not be as important for the correct distribution of HOX products along the AP-axis, at least in the late phase of spatial colinearity (<xref ref-type="bibr" rid="bib13">Duboule, 2007</xref>; <xref ref-type="bibr" rid="bib53">Tschopp et al., 2009</xref>; <xref ref-type="bibr" rid="bib35">Noordermeer and Duboule, 2013</xref>).</p><p>Even though the mechanisms underlying temporal and spatial colinearities are becoming increasingly understood, many aspects of how genomic topology is translated into sequential transcriptional activation remain to be clarified. In vertebrates, two conceptual frameworks have been proposed to account for this process, the first relying on bio-molecular mechanisms (e.g., <xref ref-type="bibr" rid="bib12">Duboule, 1994</xref>) and the second involving biophysical forces (<xref ref-type="bibr" rid="bib40">Papageorgiou, 2001</xref>). In embryonic stem (ES) cells, that is cells that reflect best the state of <italic>Hox</italic> genes before their activation, <italic>Hox</italic> clusters are decorated by both repressive (H3K27me3) and activating (H3K4me3) marks (<xref ref-type="bibr" rid="bib5">Bernstein et al., 2006</xref>; <xref ref-type="bibr" rid="bib45">Schuettengruber et al., 2007</xref>; <xref ref-type="bibr" rid="bib47">Soshnikova and Duboule, 2009</xref>; <xref ref-type="bibr" rid="bib35">Noordermeer and Duboule, 2013</xref>). Subsequently, cells that activate these genes in a time sequence resolve this bivalent chromatin state and show two opposing distributions of histone marks over the <italic>HoxD</italic> cluster: transcribed genes carry large domains of H3K4me3 marks, whereas inactive genes are covered by H3K27me3 marks only (<xref ref-type="bibr" rid="bib47">Soshnikova and Duboule, 2009</xref>).</p><p>The same dichotomy in chromatin marks over <italic>Hox</italic> gene clusters was observed in various parts of the E10.5 embryonic trunk, in parallel with the spatial colinear distribution of transcripts (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). The analysis of the 3D chromatin organization at this stage revealed a bi-modal compartmentalization, whereby active genes labeled by H3K4me3 are clustered together and physically separated from the inactive genes, labeled by H3K27me3 that are also found in a defined spatial structure (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). These 3D compartments, whose sizes correlate with the number of active vs inactive genes, may reinforce the proper maintenance of long-term transcriptional states at various AP levels by isolating <italic>Hox</italic> clusters from their surrounding chromatin and reducing interference between the active and inactive chromatin domains. Such distinct bimodal 3D organizations, associated with transcriptional regulation at <italic>Hox</italic> clusters, have been observed in other instances, either in the embryo (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>) or in mouse and human cultured cells (<xref ref-type="bibr" rid="bib20">Fraser et al., 2009</xref>; <xref ref-type="bibr" rid="bib18">Ferraiuolo et al., 2010</xref>; <xref ref-type="bibr" rid="bib55">Wang et al., 2011</xref>; <xref ref-type="bibr" rid="bib44">Rousseau et al., 2014</xref>).</p><p>However, <italic>in-embryo</italic> conformation studies were reported so far only in the context of spatial colinearity, that is by comparing samples from different body levels at the same developmental stage. Consequently, a potential association between these bimodal chromatin structures and the progressive activation of transcription along the <italic>Hox</italic> gene clusters, rather than its maintenance, remained to be assessed. In this study, we describe the 3D organization of <italic>Hox</italic> gene clusters at high resolution during the implementation of temporal colinearity in the PSM and show that their stepwise activation occurs in parallel with their physical transition from a negative to a positive domain. We also show that this process is accompanied by series of long-range contacts with the flanking gene deserts, even though these contacts remain largely invariable throughout temporal colinearity, unlike what was observed during limb development (<xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>). We discuss whether this stepwise transition of genes from one domain to the other may guide temporal colinearity or, in contrast, is a mere consequence of a sequential transcriptional activation.</p></sec><sec id="s2" sec-type="results"><title>Results</title><sec id="s2-1"><title>Inactive <italic>Hox</italic> genes in ES cells are organized into a single 3D compartment</title><p>In order to monitor the 3D organization of <italic>Hox</italic> clusters during their sequential activation, we considered ES cells as a starting point of our time curve. These cells indeed represent early embryonic cells related to blastocyst inner cell mass cells, that is when <italic>Hox</italic> genes are all supposedly silent. We hypothesized that these cells reflect the ground state 3D architecture of the <italic>Hox</italic> clusters, which we assessed by using high-resolution 4C-seq (Circular Chromosome Conformation Capture; <xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>; <xref ref-type="bibr" rid="bib54">van de Werken et al., 2012</xref>) and a variety of viewpoints within all four <italic>Hox</italic> clusters. These various baits generated similar interaction profiles with the majority of sequence reads covering the gene clusters and extending within several kilobases (kb) on either sides, as illustrated by <italic>Hoxd13</italic>, <italic>Hoxd9</italic> and <italic>Hoxd4</italic> (<xref ref-type="fig" rid="fig1">Figure 1A</xref>, <xref ref-type="fig" rid="fig1s1 fig1s2 fig1s3 fig1s4">Figure 1—figure supplements 1–4</xref>). Additional contacts were scored in the flanking gene deserts, though with significantly lower frequencies (see section ‘Temporal colinearity within a constitutive framework of long-range interactions’). The overall size of the strong interaction profiles exactly matched the distribution of bivalent chromatin marks in these cells, with a moderate level of H3K27me3 covering the cluster and rather weak H3K4me3 peaks labeling promoters (<xref ref-type="fig" rid="fig1">Figure 1A</xref>, <xref ref-type="fig" rid="fig1s1 fig1s2 fig1s3 fig1s4">Figure 1—figure supplements 1–4</xref>; <xref ref-type="bibr" rid="bib5">Bernstein et al., 2006</xref>; <xref ref-type="bibr" rid="bib47">Soshnikova and Duboule, 2009</xref>). Therefore, prior to their activation, <italic>Hox</italic> clusters are already organized into 3D chromatin compartments that physically separate the chromatin decorated by bivalent marks from the genomic surroundings, even though some contacts are established at a larger scale, outside the gene cluster itself (see section ‘Temporal colinearity within a constitutive framework of long-range interactions’).<fig-group><fig id="fig1" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.003</object-id><label>Figure 1.</label><caption><title><italic>Hox</italic> clusters in ES cells are organized as 3D compartments.</title><p>(<bold>A</bold>) Quantitative local 4C-seq signal for the <italic>Hoxd13</italic> (top), <italic>Hoxd9</italic> (middle) and <italic>Hoxd4</italic> (bottom) viewpoints in ES cells. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The boundaries of the inactive <italic>Hox</italic> gene compartments are indicated by dashed lines. The locations of <italic>Hox</italic> genes (red) and of other transcripts (black) are shown below. (<bold>B</bold>) Quantitative local 4C-seq signal for the <italic>Hoxd13</italic> (left) and <italic>Hoxb9</italic> (right) viewpoints, either in ES (orange) or in E10.5 forebrain (green) cells. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. Ratios between the 4C-seq signals in ES cells and E10.5 forebrain are indicated between the profiles, with signal in one color indicating that the viewpoint interacts more with this fragment in the sample represented by this color. Regions of increased interactions outside the 3D <italic>Hox</italic> gene compartments in ES cells are highlighted in orange. (<bold>C</bold>) Distribution of ratios inside and outside the inactive 3D <italic>Hox</italic> gene compartments in both ES and E10.5 forebrain cells. Fragments are classified either as positive in ES cells (orange), or positive in E10.5 forebrain cells (green). The number of fragments is indicated below. Significance between distribution inside and outside 3D compartments was calculated using a G-test of independence. (<bold>D</bold>) Model of 3D compartmentalization of the inactive <italic>HoxD</italic> and <italic>HoxB</italic> clusters in both ES cells and E10.5 forebrain cells. The increased contacts with the surrounding chromatin in ES cells are illustrated by invading grey lines.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.003">http://dx.doi.org/10.7554/eLife.02557.003</ext-link></p></caption><graphic xlink:href="elife02557f001"/></fig><fig id="fig1s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.004</object-id><label>Figure 1—figure supplement 1.</label><caption><title>3D compartments in the <italic>HoxD</italic> cluster are less discrete in ES cells than in embryonic brain cells.</title><p>Comparison of quantitative local 4C-seq signals for replicate samples with the indicated viewpoints, either in ES (orange) or E10.5 forebrain (green) cells. All six comparisons between two replicates in each condition are given. Viewpoints are indicated with arrowheads and regions excluded around the viewpoints are indicated with light grey boxes. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The ratios between 4C-seq signals are indicated between the corresponding profiles. The locations of <italic>Hoxd</italic> genes (red) and other transcripts (black) are shown below. Only regions covered by the random 4C-seq libraries are shown.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.004">http://dx.doi.org/10.7554/eLife.02557.004</ext-link></p></caption><graphic xlink:href="elife02557fs001"/></fig><fig id="fig1s2" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.005</object-id><label>Figure 1—figure supplement 2.</label><caption><title>3D compartments in the <italic>HoxD</italic> and <italic>HoxB</italic> cluster are less discrete in ES cells than in embryonic brain cells.</title><p>Comparison of quantitative local 4C-seq signals for replicate samples with the indicated viewpoints, either in ES (orange) or E10.5 forebrain (green) cells. All six comparisons between two replicates in each condition are given. Viewpoints are indicated with arrowheads and regions excluded around the viewpoints are indicated with light grey boxes. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The ratios between 4C-seq signals are indicated between the corresponding profiles. The locations of <italic>Hoxd</italic> and <italic>Hoxb</italic> genes (red) and other transcripts (black) are shown below. Only regions covered by the random 4C-seq libraries are shown.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.005">http://dx.doi.org/10.7554/eLife.02557.005</ext-link></p></caption><graphic xlink:href="elife02557fs002"/></fig><fig id="fig1s3" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.006</object-id><label>Figure 1—figure supplement 3.</label><caption><title>3D compartments in the <italic>HoxB</italic> cluster are less discrete in ES cells than in embryonic brain cells.</title><p>Comparison of quantitative local 4C-seq signals for replicate samples with the indicated viewpoints, either in ES (orange) or E10.5 forebrain (green) cells. All six comparisons between two replicates in each condition are given. Viewpoints are indicated with arrowheads and regions excluded around the viewpoints are indicated with light grey boxes. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The ratios between 4C-seq signals are indicated between the corresponding profiles. The locations of <italic>Hoxb</italic> genes (red) and other transcripts (black) are shown below. Only regions covered by the random 4C-seq libraries are shown.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.006">http://dx.doi.org/10.7554/eLife.02557.006</ext-link></p></caption><graphic xlink:href="elife02557fs003"/></fig><fig id="fig1s4" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.007</object-id><label>Figure 1—figure supplement 4.</label><caption><title>3D compartments in the <italic>HoxC</italic> and <italic>HoxA</italic> cluster are less discrete in ES cells than in embryonic brain cells.</title><p>Comparison of quantitative local 4C-seq signals with the indicated viewpoints in ES (orange) or E10.5 forebrain (green) cells. Viewpoints are indicated with arrowheads and regions excluded around the viewpoints are indicated with light grey boxes. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The ratios between 4C-seq signals are indicated between the corresponding profiles. The locations of <italic>Hoxc</italic> and <italic>Hoxa</italic> genes (red) and other transcripts (black) are shown below. Only regions covered by the random 4C-seq libraries are shown.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.007">http://dx.doi.org/10.7554/eLife.02557.007</ext-link></p></caption><graphic xlink:href="elife02557fs004"/></fig><fig id="fig1s5" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.008</object-id><label>Figure 1—figure supplement 5.</label><caption><title>Distribution of ratios inside and outside the inactive 3D <italic>Hox</italic> gene compartments in both ES and E10.5 forebrain cells.</title><p>The comparison between replicate samples one (as used in the main text) is indicated on the left, the comparison between combined replicate samples is indicated at the center left, the comparison between ES cell replicates is indicated at the center right and the comparison between E10.5 forebrain replicates is indicated on the right. Fragments are classified as positive either in ES (orange) or in E10.5 forebrain (green) cells within the region covered by the random 4C-seq libraries. The number of fragments is indicated below. Significance between distribution inside and outside the 3D compartments was calculated using a G-test of independence.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.008">http://dx.doi.org/10.7554/eLife.02557.008</ext-link></p></caption><graphic xlink:href="elife02557fs005"/></fig><fig id="fig1s6" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.009</object-id><label>Figure 1—figure supplement 6.</label><caption><title>Different discretion of 3D compartments is not due to overall increased background signal.</title><p>(<bold>A</bold>) Distribution of 4C-seq signal on chromosome 2 from viewpoints in the <italic>HoxD</italic> cluster. On the right, a schematic representation of chromosome 2 is given, with color codes for the three categories that have been quantified in ES cells and E10.5 forebrain indicated below. Comparison of distributions between ES cells and E10.5 forebrain show that TAD signal in ES cells are considerably increased, but that more distal signal is reduced. Elevated signal in the TADs in ES cells is therefore not a representation of generally increased background signal. (<bold>B</bold>) Distribution of 4C-seq signal on chromosome 11 from viewpoints in the <italic>HoxB</italic> cluster. Similar effects are observed as for viewpoints in the <italic>HoxD</italic> cluster.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.009">http://dx.doi.org/10.7554/eLife.02557.009</ext-link></p></caption><graphic xlink:href="elife02557fs006"/></fig><fig id="fig1s7" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.010</object-id><label>Figure 1—figure supplement 7.</label><caption><title>Increased <italic>Hox</italic> background transcription in ES cells.</title><p>(<bold>A</bold>) Expression levels of <italic>Hox</italic> genes and of four housekeeping genes in both ES and E10.5 forebrain cells, as determined by RNA-seq. The large majority of <italic>Hox</italic> genes show low level activity in ES cells, whereas only few, very low, transcribed <italic>Hox</italic> genes are identified in E10.5 forebrain. In contrast, the expression levels of selected housekeeping genes are within a similar range (maximum threefold difference). (<bold>B</bold>) Overall gene expression patterns in ES and E10.5 forebrain cells are not significantly different. Box plots showing the overall distribution of RNA-seq signals per gene (RPKM), with colored boxes indicating the 25 to 75% range and whiskers indicating the 10 to 90% range. Differences between distributions were scored using a two-sided Welch two samples <italic>t</italic> test. (<bold>C</bold>) Quantitation of selected spliced <italic>Hox</italic> gene transcripts in ES cell and E10.5 forebrain samples as determined by RT-qPCR, with amounts in each sample relative to the <italic>Tubb2c</italic> gene. Below each sample, the specific product of a representative qPCR reaction is displayed. Color-coded dots are used to classify the different outcomes (see legend).</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.010">http://dx.doi.org/10.7554/eLife.02557.010</ext-link></p></caption><graphic xlink:href="elife02557fs007"/></fig></fig-group></p><p>These global 3D domains including the <italic>Hox</italic> clusters and their immediate flanking DNAs resemble the chromatin architecture found in embryonic forebrain cells, where the silent <italic>Hox</italic> clusters are covered by high levels of H3K27me3 only (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). A more quantitative comparison in 3D architectures between ES cells and E10.5 forebrain cells nevertheless indicated that in ES cells, <italic>Hox</italic> genes interacted more with the outside chromatin, relative to their interactions within the cluster, as compared to forebrain cells (<xref ref-type="fig" rid="fig1">Figure 1B,C</xref>, <xref ref-type="fig" rid="fig1s1 fig1s2 fig1s3 fig1s4 fig1s5">Figure 1—figure supplements 1–5</xref>). Therefore, despite the fact that the clusters are presumably inactive in both situations, the presence of bivalent marks in ES cells coincided with a 3D domain that has elevated relative levels of interactions with the directly surrounding regions, when compared to its counterpart in brain cells (<xref ref-type="fig" rid="fig1">Figure 1D</xref>, left and <xref ref-type="fig" rid="fig1s6">Figure 1—figure supplement 6A</xref>). This difference is more pronounced at the <italic>HoxB</italic> cluster (<xref ref-type="fig" rid="fig1">Figure 1B</xref>, right). In embryonic forebrain cells, <italic>HoxB</italic> forms a single 3D compartment, excluding the 80 kb large repeat-rich intergenic region located between <italic>Hoxb13</italic> and <italic>Hoxb9</italic>, which loops out (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). In contrast, both the <italic>Hoxb13</italic> and <italic>Hoxb9</italic> viewpoints revealed local 3D compartments in ES cells, matching again the extent of bivalent histone marks, yet these two compartments remained separated and did not fuse. Rather, they displayed increased interactions with the nearby chromatin, as if decreased internal interactions would increase contacts outside the cluster (<xref ref-type="fig" rid="fig1">Figure 1B,C</xref>, <xref ref-type="fig" rid="fig1s2 fig1s3 fig1s5">Figure 1—figure supplements 2, 3 and 5</xref>). In ES cells, the <italic>HoxB</italic> cluster is thus organized in two 3D compartments, which have more interactions with their genomic surroundings than in forebrain cells (<xref ref-type="fig" rid="fig1">Figure 1D</xref>, right and <xref ref-type="fig" rid="fig1s6">Figure 1—figure supplement 6B</xref>).</p><p>To have a possibly more unbiased view on how 3D compartments and the presence of H3K27me3 and H3K4me3 modifications relate to each other, in both ES and brain cells, we devised an approach to correlate 4C-seq signals with either H3K27me3 or H3K4me3 ChIP-seq signal (<xref ref-type="table" rid="tbl1">Table 1</xref>; ‘Materials and methods’). In both cell types, H3K27me3 marks strongly correlated with the 3D organization, suggesting a direct link between these two readouts. A considerably lower correlation was scored for <italic>HoxB</italic>, perhaps related to the absence of clustering of the two H3K27me3 marked sub-domains. In contrast, no particular correlation was observed between the 3D organization and the presence of H3K4me3 marks, in the bivalent state (<xref ref-type="table" rid="tbl1">Table 1</xref>), suggesting that H3K4me3 marks and/or the associated factors do not noticeably contribute to the formation of 3D compartments in ES cells.<table-wrap id="tbl1" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.011</object-id><label>Table 1.</label><caption><p>Spearman's rank correlation coefficient between pairs of 4C-seq and ChIP-seq samples</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.011">http://dx.doi.org/10.7554/eLife.02557.011</ext-link></p></caption><table frame="hsides" rules="groups"><thead><tr><th/><th colspan="3">ChIP-seq</th></tr><tr><th>4C-seq</th><th>Input</th><th>H3K27me3</th><th>H3K4me3</th></tr></thead><tbody><tr><td><italic>Hoxd13</italic> ES cells 1</td><td align="char" char=".">−0.14</td><td align="char" char="."><bold>0.52</bold></td><td align="char" char=".">0.24</td></tr><tr><td><italic>Hoxd13</italic> ES cells 2</td><td align="char" char=".">−0.07</td><td align="char" char="."><bold>0.40</bold></td><td align="char" char=".">0.22</td></tr><tr><td><italic>Hoxd13</italic> E8.5 PSM</td><td align="char" char=".">−0.03</td><td align="char" char="."><bold>0.58</bold></td><td align="char" char=".">0.13</td></tr><tr><td><italic>Hoxd13</italic> E10.5 Forebrain 1</td><td align="char" char=".">−0.12</td><td align="char" char="."><bold>0.67</bold></td><td align="char" char=".">0.26</td></tr><tr><td><italic>Hoxd13 E10.5</italic> Forebrain 2</td><td align="char" char=".">−0.09</td><td align="char" char="."><bold>0.69</bold></td><td align="char" char=".">0.25</td></tr><tr><td><italic>Hoxd13</italic> E10.5 Anterior trunk</td><td align="char" char=".">−0.07</td><td align="char" char="."><bold>0.80</bold></td><td align="char" char=".">0.30</td></tr><tr><td><italic>Hoxd9</italic> ES cells 1</td><td align="char" char=".">−0.08</td><td align="char" char="."><bold>0.63</bold></td><td align="char" char=".">0.28</td></tr><tr><td><italic>Hoxd9</italic> ES cells 2</td><td align="char" char=".">−0.13</td><td align="char" char="."><bold>0.59</bold></td><td align="char" char=".">0.26</td></tr><tr><td><italic>Hoxd9</italic> E8.5 PSM</td><td align="char" char=".">−0.05</td><td align="char" char="."><bold>0.31</bold></td><td align="char" char=".">0.29</td></tr><tr><td><italic>Hoxd9</italic> E10.5 Forebrain 1</td><td align="char" char=".">−0.08</td><td align="char" char="."><bold>0.66</bold></td><td align="char" char=".">0.26</td></tr><tr><td><italic>Hoxd9</italic> E10.5 Forebrain 2</td><td align="char" char=".">−0.12</td><td align="char" char="."><bold>0.61</bold></td><td align="char" char=".">0.28</td></tr><tr><td><italic>Hoxd9</italic> E10.5 Anterior trunk</td><td align="char" char=".">−0.15</td><td align="char" char="."><bold>0.67</bold></td><td align="char" char=".">0.47</td></tr><tr><td><italic>Hoxd4</italic> ES cells 1</td><td align="char" char=".">0.01</td><td align="char" char="."><bold>0.48</bold></td><td align="char" char=".">0.11</td></tr><tr><td><italic>Hoxd4</italic> ES cells 2</td><td align="char" char=".">−0.07</td><td align="char" char="."><bold>0.50</bold></td><td align="char" char=".">0.29</td></tr><tr><td><italic>Hoxd4</italic> E8.5 PSM</td><td align="char" char=".">−0.04</td><td align="char" char=".">0.04</td><td align="char" char="."><bold>0.38</bold></td></tr><tr><td><italic>Hoxd4</italic> E10.5 Forebrain 1</td><td align="char" char=".">−0.05</td><td align="char" char="."><bold>0.59</bold></td><td align="char" char=".">0.24</td></tr><tr><td><italic>Hoxd4</italic> E10.5 Forebrain 2</td><td align="char" char=".">−0.04</td><td align="char" char="."><bold>0.58</bold></td><td align="char" char=".">0.27</td></tr><tr><td><italic>Hoxd4</italic> E10.5 Anterior trunk</td><td align="char" char=".">−0.07</td><td align="char" char=".">0.16</td><td align="char" char="."><bold>0.59</bold></td></tr><tr><td><italic>Hoxc13</italic> ES cells 1</td><td align="char" char=".">−0.03</td><td align="char" char="."><bold>0.39</bold></td><td align="char" char=".">0.20</td></tr><tr><td><italic>Hoxc13</italic> E8.5 PSM</td><td align="char" char=".">−0.03</td><td align="char" char="."><bold>0.55</bold></td><td align="char" char=".">−0.03</td></tr><tr><td><italic>Hoxc13</italic> E10.5 Forebrain 1</td><td align="char" char=".">−0.07</td><td align="char" char="."><bold>0.57</bold></td><td align="char" char=".">0.18</td></tr><tr><td><italic>Hoxc13</italic> E10.5 Anterior trunk</td><td align="char" char=".">−0.05</td><td align="char" char="."><bold>0.82</bold></td><td align="char" char=".">0.00</td></tr><tr><td><italic>Hoxb13</italic> ES cells 1</td><td align="char" char=".">−0.05</td><td align="char" char="."><bold>0.12</bold></td><td align="char" char=".">0.02</td></tr><tr><td><italic>Hoxb13</italic> ES cells 2</td><td align="char" char=".">−0.08</td><td align="char" char=".">−0.01</td><td align="char" char="."><bold>0.15</bold></td></tr><tr><td><italic>Hoxb13</italic> E8.5 PSM</td><td align="char" char=".">0.10</td><td align="char" char="."><bold>0.29</bold></td><td align="char" char=".">−0.17</td></tr><tr><td><italic>Hoxb13</italic> E10.5 Forebrain 1</td><td align="char" char=".">0.02</td><td align="char" char="."><bold>0.48</bold></td><td align="char" char=".">0.09</td></tr><tr><td><italic>Hoxb13</italic> E10.5 Forebrain 2</td><td align="char" char=".">0.08</td><td align="char" char="."><bold>0.44</bold></td><td align="char" char=".">0.10</td></tr><tr><td><italic>Hoxb13</italic> E10.5 Anterior trunk</td><td align="char" char=".">−0.03</td><td align="char" char="."><bold>0.49</bold></td><td align="char" char=".">0.26</td></tr><tr><td><italic>Hoxb9</italic> ES cells 1</td><td align="char" char=".">0.01</td><td align="char" char="."><bold>0.47</bold></td><td align="char" char=".">0.09</td></tr><tr><td><italic>Hoxb9</italic> ES cells 2</td><td align="char" char=".">0.03</td><td align="char" char="."><bold>0.34</bold></td><td align="char" char=".">0.04</td></tr><tr><td><italic>Hoxb9</italic> E8.5 PSM</td><td align="char" char=".">−0.04</td><td align="char" char=".">−0.30</td><td align="char" char="."><bold>0.57</bold></td></tr><tr><td><italic>Hoxb9</italic> E10.5 Forebrain 1</td><td align="char" char=".">0.02</td><td align="char" char="."><bold>0.63</bold></td><td align="char" char=".">0.19</td></tr><tr><td><italic>Hoxb9</italic> E10.5 Forebrain 2</td><td align="char" char=".">0.03</td><td align="char" char="."><bold>0.59</bold></td><td align="char" char=".">0.16</td></tr><tr><td><italic>Hoxb9</italic> E10.5 Anterior trunk</td><td align="char" char=".">0.06</td><td align="char" char=".">−0.01</td><td align="char" char="."><bold>0.69</bold></td></tr><tr><td><italic>Hoxa13</italic> ES cells 1</td><td align="char" char=".">0.10</td><td align="char" char="."><bold>0.52</bold></td><td align="char" char=".">0.14</td></tr><tr><td><italic>Hoxa13</italic> E8.5 PSM</td><td align="char" char=".">0.10</td><td align="char" char="."><bold>0.58</bold></td><td align="char" char=".">0.12</td></tr><tr><td><italic>Hoxa13</italic> E10.5 Forebrain 1</td><td align="char" char=".">0.07</td><td align="char" char="."><bold>0.60</bold></td><td align="char" char=".">0.22</td></tr><tr><td><italic>Hoxa13</italic> E10.5 Anterior trunk</td><td align="char" char=".">0.06</td><td align="char" char="."><bold>0.73</bold></td><td align="char" char=".">0.20</td></tr></tbody></table><table-wrap-foot><fn><p>Spearman's rank correlation coefficient between pairs of 4C-seq and ChIP-seq samples in different samples (see section ‘Material and methods’ for methodology). For each 4C-seq sample, the highest correlating ChIP-seq sample is highlighted in bold.</p></fn></table-wrap-foot></table-wrap></p></sec><sec id="s2-2"><title><italic>Hox</italic> genes are transcribed at low levels in ES cells</title><p>It was previously reported that genes covered by bivalent marks in ES cells can be transcribed at low levels, resulting in detectable spliced transcripts (<xref ref-type="bibr" rid="bib49">Stock et al., 2007</xref>). We assessed whether the observed difference in the strength and homogeneity of the interaction profiles between ES cells and embryonic brain cells was associated with distinct levels of background transcription. In ES cells, RNA-seq detected transcription for most <italic>Hox</italic> genes, though generally at very low level (<xref ref-type="fig" rid="fig1s7">Figure 1—figure supplement 7A,B</xref>). RT-qPCR of a subset of transcripts confirmed that some of these low-level transcripts (particularly the <italic>Hoxd13</italic> and <italic>Hoxb13</italic> transcripts) constitute genuine processed transcripts (<xref ref-type="fig" rid="fig1s7">Figure 1—figure supplement 7C</xref>). In contrast, transcription of <italic>Hox</italic> genes in E10.5 forebrain cells was rarely detected, and no reliable spliced transcripts were detected (<xref ref-type="fig" rid="fig1s7">Figure 1—figure supplement 7</xref>). Therefore, when <italic>Hox</italic> genes are decorated by bivalent chromatin marks, they appear more permissive for background transcription as compared to other cell types where they are covered by H3K27me3 marks only, likely illustrating the increased resistance to transcription of the latter condition. In this context, posterior <italic>Hox</italic> genes seems to be more prone to background transcriptional activation in ES cells than more anterior <italic>Hox</italic> genes, in contrast to their subsequent dynamics of activation in future embryonic tissues where anterior genes come first. This may reflect the presence of strong enhancers in their vicinity (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>).</p></sec><sec id="s2-3"><title>Dynamics of 3D compartments during sequential <italic>Hox</italic> gene activation</title><p>Next, we assessed whether this large 3D domain observed in ES cells is modified when <italic>Hox</italic> genes become activated in the pre-somitic mesoderm (PSM) or instead, whether the previously observed positive and negative compartments are only established at a later stage to fix and memorize particular combinations of <italic>Hox</italic> gene activities determined at earlier stages and at various body levels. For this purpose, we compared the 4C-seq profiles from ES cells with those obtained from early embryonic E8.5 PSM cells dissected out at Theiler stage 13, posterior from the approximate level of the 12<sup>th</sup> to 14<sup>th</sup> forming somite (<xref ref-type="fig" rid="fig2">Figure 2A</xref>, scheme; <xref ref-type="fig" rid="fig2s1 fig2s2">Figure 2—figure supplements 1 and 2</xref>). In the most caudal aspect of this latter cellular territory, transcriptional activation had progressed up to the <italic>Hoxd9</italic> gene, whereas the <italic>Hoxd10</italic> to <italic>Hoxd13</italic> loci remained silent (<xref ref-type="bibr" rid="bib47">Soshnikova and Duboule, 2009</xref>). This cellular population was thus composed of a mixture of cells positive and negative for <italic>Hoxd9</italic> expression, whereas all cells were negative for <italic>Hoxd13</italic>. Conversely, the majority of cells expressed <italic>Hoxd4</italic>. The inactive <italic>Hoxd13</italic> viewpoint interacted mostly with the domain labeled by H3K27me3, at the centromeric side of the cluster (<xref ref-type="fig" rid="fig2">Figure 2A</xref>, bottom left). In contrast, the active <italic>Hoxd4</italic> gene essentially interacted with the other transcribed genes on the telomeric side of the cluster, labeled by H3K4me3 marks (<xref ref-type="fig" rid="fig2">Figure 2A</xref>, bottom right). The same bi-modal 3D organization was observed for all <italic>Hox</italic> gene clusters (<xref ref-type="fig" rid="fig2s1 fig2s2">Figure 2—figure supplements 1 and 2</xref>).<fig-group><fig id="fig2" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.012</object-id><label>Figure 2.</label><caption><title>Bi-modal 3D organization of <italic>Hox</italic> clusters upon sequential activation.</title><p>(<bold>A</bold>) Quantitative local 4C-seq signal for the <italic>Hoxd13</italic> (left, centromeric side of <italic>HoxD</italic> cluster) and <italic>Hoxd4</italic> (right, telomeric side of <italic>HoxD</italic> cluster) viewpoints, either in ES (orange), or E8.5 pre-somitic mesoderm (cyan) cells. Below, the H3K27me3 and H3K4me3 ChIP-seq profiles are aligned. The colinear expression status of <italic>Hoxd</italic> genes in each sample is schematized below the ChIP-seq profiles, with active genes in blue and inactive genes in red. Ratios between the 4C-seq signals in different samples are indicated between the profiles. The boundaries separating active from inactive <italic>Hox</italic> gene compartments are indicated by dashed lines. The locations of <italic>Hoxd</italic> genes (red) and other transcripts (black) are shown below. The samples are shown on the left and cartoons summarizing the genome organizations are indicated on the right. (<bold>B</bold>) Spearman's rank correlation coefficient between pairs of 4C-seq and ChIP-seq samples, in early and late embryonic material.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.012">http://dx.doi.org/10.7554/eLife.02557.012</ext-link></p></caption><graphic xlink:href="elife02557f002"/></fig><fig id="fig2s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.013</object-id><label>Figure 2—figure supplement 1.</label><caption><title>Upon sequential activation, the <italic>HoxD</italic> cluster adopts a bi-modal 3D organization.</title><p>Quantitative local 4C-seq signals for the indicated <italic>Hoxd</italic> gene viewpoints. Profiles are displayed for ES (orange) and E8.5 pre-somitic mesoderm (cyan) cells. The viewpoints are indicated with arrowheads and excluded regions around the viewpoints are indicated with light grey boxes. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The ratios between 4C-seq signals are indicated between the respective profiles, with the signal in one particular color indicating that the viewpoint interacts more with fragments in the sample of the same color. The locations of <italic>Hoxd</italic> genes (red) and other transcripts (black) are shown below. Only the region covered by the random 4C-seq library is shown.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.013">http://dx.doi.org/10.7554/eLife.02557.013</ext-link></p></caption><graphic xlink:href="elife02557fs008"/></fig><fig id="fig2s2" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.014</object-id><label>Figure 2—figure supplement 2.</label><caption><title>Upon sequential activation, other <italic>Hox</italic> clusters adopt a bi-modal 3D organization as well.</title><p>Quantitative local 4C-seq signals for the indicated <italic>Hox</italic> gene viewpoints in other <italic>Hox</italic> clusters. Profiles are displayed for ES (orange) and E8.5 pre-somitic mesoderm (cyan) cells. The viewpoints are indicated with arrowheads and excluded regions around the viewpoints are indicated with light grey boxes. Below, the H3K27me3 and H3K4me3 ChIP-seq signals are aligned. The ratios between 4C-seq signals are indicated between the respective profiles, with the signal in one particular color indicating that the viewpoint interacts more with fragments in the sample of the same color. The locations of both <italic>Hox</italic> genes (red) and other transcripts (black) are shown below. Only regions covered by the random 4C-seq libraries are shown.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.014">http://dx.doi.org/10.7554/eLife.02557.014</ext-link></p></caption><graphic xlink:href="elife02557fs009"/></fig></fig-group></p><p>We correlated 4C-seq signals with ChIP-seq data in both early and late embryonic samples (<xref ref-type="fig" rid="fig2">Figure 2B</xref>; <xref ref-type="table" rid="tbl1">Table 1</xref>) and <italic>Hoxd13</italic> always strongly correlated with H3K27me3 histone marks. In contrast, in both the E8.5 tail bud and the E10.5 anterior trunk, the interactions of the active <italic>Hoxd4</italic> gene correlated primarily with H3K4me3 marks. The contacts established by <italic>Hoxd9</italic> correlated both with H3K27me3 and H3K4me3, either in E8.5 tailbuds, or in E10.5 anterior trunk, likely due to the presence of both expressing and non-expressing cells. At both stages where <italic>Hox</italic> clusters are partially active, the patterns of 3D compartmentalization and histone marks thus strongly correlated. Therefore, step wise <italic>Hox</italic> gene transcriptional activation, at least for the <italic>Hoxd9</italic> to <italic>Hoxd13</italic> genes, is accompanied by a conformational separation between active and inactive domains, which pre-figures their 3D organization at later developmental stages along the AP-axis (<xref ref-type="fig" rid="fig2">Figure 2</xref>; <xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>).</p></sec><sec id="s2-4"><title>Posterior <italic>Hoxd</italic> genes switch autonomously between 3D compartments</title><p>Temporal colinearity was initially defined as the sequential activation of <italic>Hox</italic> genes according to their positions in the clusters (<xref ref-type="bibr" rid="bib26">Izpisua-Belmonte et al., 1991</xref>; <xref ref-type="bibr" rid="bib12">Duboule, 1994</xref>). However, studies on the global transcriptional organization of the <italic>HoxD</italic> cluster, at least in the developing spinal cord, revealed two large and regulatory-independent modules, which separate ‘posterior’ genes (the <italic>AbdB</italic>-related <italic>Hoxd9</italic> to <italic>Hoxd13</italic> genes) from the rest of the gene cluster (<xref ref-type="bibr" rid="bib50">Tschopp et al., 2012</xref>). Also, in different developmental contexts such as the limbs and the cecum, groups of neighboring <italic>Hoxd</italic> genes are activated as single regulatory blocks (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>; <xref ref-type="bibr" rid="bib7">Delpretti et al., 2013</xref>). We thus assessed whether the transition in chromatin domains also occurred stepwise or, alternatively, if large domains consisting of multiple genes were initially organized in space, followed by sequential gene transcription within these domains.</p><p>We first compared the 3D cluster architecture over the course of embryonic development, between the E8.5 PSM and dissected E9.5 tail buds (<xref ref-type="fig" rid="fig3">Figure 3</xref>). This latter sample was obtained after cutting off the most caudal part of E9.5 embryos (Theiler stage 15) right after the incipient hind limb bud, that is at ca. somite 26–27 level. Accordingly, this sample contained the tail bud proper as well as some tissue localized slightly more rostral. During this 24 hr time interval, the <italic>Hoxd10</italic> and <italic>Hoxd11</italic> genes become robustly activated in these cells, which are derived from a sub-population of the sample dissected at E8.5. In the E8.5 PSM, <italic>Hoxd10</italic> and <italic>Hoxd11</italic> are still silenced.<fig id="fig3" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.015</object-id><label>Figure 3.</label><caption><title>Activated <italic>Hoxd</italic> genes switch compartments.</title><p>Quantitative local 4C-seq signals for the <italic>Hoxd13</italic>, <italic>Hoxd11 Hoxd9</italic> and <italic>Hoxd4</italic> viewpoints in either E8.5 pre-somitic mesoderm (cyan), E9.5 tail bud (brown) or E10.5 tail bud (purple) cells. The colinear expression status of <italic>Hoxd</italic> genes is schematized below each profile and, on the left, below each cartoon. Ratios between 4C-seq signals in different samples are indicated between the corresponding profiles. The boundaries between active and inactive <italic>Hox</italic> gene compartments are indicated by dashed lines and regions displaying important changes in interactions, as discussed in the text, are highlighted. Black arrows point towards opposing interacting behaviors due to the heterogeneous activity state of the viewpoint in the sample. The locations of <italic>Hoxd</italic> genes (red) and other transcripts (black) are shown below.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.015">http://dx.doi.org/10.7554/eLife.02557.015</ext-link></p></caption><graphic xlink:href="elife02557f003"/></fig></p><p>The re-organization of compartmentalization occurring along with gene activation over this 24 hr period was clearly revealed by comparing the profile obtained when using <italic>Hoxd11</italic> as a viewpoint with those obtained with either <italic>Hoxd4</italic> or <italic>Hoxd13</italic> (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top ratio between E8.5 PSM and E9.5 Tail bud). When switching from an inactive to an active state, <italic>Hoxd11</italic> re-deployed its interactions from the inactive, centromeric compartment (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top ratio: blue shaded area) to the active telomeric compartment (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top ratio: brown shaded area). These negative and positive compartments can be identified by the interaction profile of either <italic>Hoxd13</italic> (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top left) or <italic>Hoxd4</italic> (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top right), respectively. Accordingly, <italic>Hoxd4</italic> shifted its interactions towards the centromeric (active) part of the cluster in both E9.5 and E10.5 samples (<xref ref-type="fig" rid="fig3">Figure 3</xref>; right), to contact <italic>Hoxd10</italic>, <italic>Hoxd11</italic> and, to some extent, <italic>Hoxd12</italic> (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top ratio: brown shaded area). Of note, the dissected E8.5 PSM contained a mixture of cells either positive or negative for <italic>Hoxd9</italic> transcription, which coincided with this gene showing conspicuous contacts with both extremities of the gene cluster, depending whether it was active (right) or inactive (left) (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top, black arrows). In contrast, the E9.5 dissection contains a more homogenous cell population, strongly expressing this gene. As a consequence, in this latter sample, <italic>Hoxd9</italic> contacts more strongly the now expressed <italic>Hoxd10</italic> and <italic>Hoxd11</italic> genes, whereas the interactions with <italic>Hoxd13</italic> or <italic>Evx2</italic> are strongly diminished (<xref ref-type="fig" rid="fig3">Figure 3</xref>; compare top with middle panels).</p><p>In the E10.5 tail bud, the terminal part of the cluster containing <italic>Hoxd12</italic> and <italic>Hoxd13</italic> has been activated, as shown by the increased contacts established by <italic>Hoxd13</italic> with the telomeric part of the cluster, indicating that the full <italic>HoxD</italic> array had been processed and that all genes were now (at least in part) included into a ‘positive’ compartment (<xref ref-type="fig" rid="fig3">Figure 3</xref>, bottom ratio: purple shading). While considerably increased, contacts of <italic>Hoxd13</italic> with the telomeric part of the cluster were however weak (<xref ref-type="fig" rid="fig3">Figure 3</xref>, bottom, black arrows), most likely reflecting the restricted expression of <italic>Hoxd13</italic> at this stage, in a small subset of the dissected cells. In the same context, contacts established both by <italic>Hoxd4</italic> and <italic>Hoxd9</italic> with the centromeric part of the cluster extended towards the end of the cluster along with the developmental stage, such that both reached <italic>Hoxd12</italic> in E10.5 samples, whereas <italic>Hoxd13</italic> was only weakly contacted, corresponding to the results obtained when using <italic>Hoxd13</italic> as a bait (<xref ref-type="fig" rid="fig3">Figure 3</xref>; bottom left).</p><p>Therefore, it appears that the colinear time sequence in <italic>Hox</italic> genes activation is paralleled by a progressive transition in the chromatin structure, with a positive domain gaining in size along with time, at the expense of the negative domain, as best seen by the extension of <italic>Hoxd4</italic> contacts. At E8.5, these interactions extended up to <italic>Hoxd8-Hoxd9</italic>. In E9.5 samples <italic>Hoxd10</italic> was clearly contacted, and in E10.5 <italic>Hoxd11</italic> and <italic>Hoxd12</italic> were also involved (<xref ref-type="fig" rid="fig3">Figure 3</xref>, right column). These dynamic topologies suggest a stepwise transition of the genes from the negative to the positive compartment, rather than the switch of large groups of multiple transcription units, following a discrete and global chromatin re-organization.</p></sec><sec id="s2-5"><title>Memorizing bimodal chromatin configurations</title><p>During axial extension, <italic>Hox</italic> genes are activated in the most posterior aspect of the elongating embryo (<xref ref-type="bibr" rid="bib9">Deschamps and van Nes, 2005</xref>). It is thus possible that cells implementing this stepwise transition in chromatin domains can fix and memorize their bimodal distribution once they exit the posterior zone of activation, leading to the colinear <italic>Hox</italic> conformations observed along the AP-axis (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). Accordingly, one would expect cellular territories along the developing body axis to maintain the same bimodal combinations as those established at the time of their origin, during early axial extension. We looked at the similarities in bimodal profiles between posterior samples dissected at different times on the one hand, and various samples micro-dissected at different body levels, from E10.5 embryos, on the other hand (<xref ref-type="fig" rid="fig4">Figure 4</xref>).<fig id="fig4" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.016</object-id><label>Figure 4.</label><caption><title>The bimodal 3D organization of <italic>Hox</italic> cluster may help memorize states of colinear expression.</title><p>Quantitative local 4C-seq signals for the <italic>Hoxd13</italic>, <italic>Hoxd11 Hoxd9</italic> and <italic>Hoxd4</italic> viewpoints, in samples taken at various anterior to posterior positions along the developing body axis from E10.5 embryos. Anterior trunk (red), lumbo-sacral trunk (blue) and tail bud (purple) tissues were used and the approximate expression status of <italic>Hoxd</italic> genes in every sample is schematized below each profile (as for <xref ref-type="fig" rid="fig3">Figure 3</xref>). Ratios between 4C-seq signals in the different samples are indicated between the corresponding profiles. The boundaries between active and inactive <italic>Hox</italic> gene compartments are indicated by dashed lines and regions displaying important changes in interactions, as discussed in the text, are highlighted. The locations of <italic>Hoxd</italic> genes (red) and other transcripts (black) are shown below. On the right, cartoons summarizing the 3D genome organization of the <italic>HoxD</italic> cluster are indicated.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.016">http://dx.doi.org/10.7554/eLife.02557.016</ext-link></p></caption><graphic xlink:href="elife02557f004"/></fig></p><p>The profile obtained from E8.5 PSM (<xref ref-type="fig" rid="fig3">Figure 3</xref>, top), right at the onset of <italic>Hoxd9</italic> activation globaly aligned with that observed in the ‘anterior trunk’ sample at E10.5 (<xref ref-type="fig" rid="fig4">Figure 4</xref>, top), that is a cellular domain with a posterior boundary positioned approximately at the <italic>Hoxd9</italic> anterior limit of expression. In both cases, <italic>Hoxd9</italic> clearly contacted the negative domain, as defined by the <italic>Hoxd13</italic> contacts (<xref ref-type="fig" rid="fig4">Figure 4</xref>, top), whereas some weak contacts were also scored with the positive domain, as determined by <italic>Hoxd4</italic> contacts, indicating that the posterior limit of the dissection was slightly below the <italic>Hoxd9</italic> boundary. These contacts were somehow stronger, in proportion, in the E8.5 than in the E10.5 dissection.</p><p>At E9.5 (Theiler stage 14), the ‘tail bud’ (i.e., from the start of the non-segmented mesoderm) was dissected from ca. somite 22 to 25 and caudally. At this stage, <italic>Hoxd9</italic> is robustly transcribed, whereas <italic>Hoxd11</italic> has just started transcriptional activation. The interaction profiles obtained in this tissue were most similar to those obtained when a fragment of E10.5 trunk was dissected out that grossly corresponded to the future lumbo-sacral region, at levels 22 to 28, that is the AP levels supposedly produced in the E9.5 tail bud (<xref ref-type="fig" rid="fig4">Figure 4</xref>, middle). At this AP level, <italic>Hoxd9</italic> is fully activated and this was reflected by the quasi absence of contact with <italic>Hoxd13</italic> whereas, conversely, strong interactions appeared with the active part of the cluster (<xref ref-type="fig" rid="fig4">Figure 4</xref>, top ratio: red and blue shading). This was controlled by using <italic>Hoxd4</italic> as bait, since contacts were now clearly scored with <italic>Hoxd9</italic> and <italic>Hoxd10</italic> and, to a lesser extent, with <italic>Hoxd11</italic> (<xref ref-type="fig" rid="fig4">Figure 4</xref>, top ratio: blue shading).</p><p>In this lumbo-sacral sample, neither <italic>Hoxd12</italic> nor <italic>Hoxd13</italic> are as yet transcribed, which coincided with the absence of contact between <italic>Hoxd13</italic> and the active part of the gene cluster (<xref ref-type="fig" rid="fig4">Figure 4</xref>, middle, left). On the other hand, <italic>Hoxd11</italic> expectedly displayed a mixed interaction profile, contacting both the negative and positive domains, likely reflecting the presence of both expressing and non-expressing cells (<xref ref-type="fig" rid="fig4">Figure 4</xref>, middle). In the most caudal piece of the E10.5 mouse embryo, interactions between <italic>Hoxd12</italic>, <italic>Hoxd13</italic> and the positive domain were finally detected, suggesting that the entire cluster falls into a single spatial domain (<xref ref-type="fig" rid="fig4">Figure 4</xref>, bottom ratio: purple shading). Here again, however, though the interactions were significant, they were not particularly strong, suggesting the presence of a mixed cell population. Based on these data, we propose that the bimodal distributions are frozen in those cells leaving the zone of proliferation, at the caudal aspect of the embryo where temporal colinearity is potentially processed. These 3D structures, and hence the <italic>Hox</italic> transcription programs, will thus be maintained and memorize the various AP levels from which they originate.</p></sec><sec id="s2-6"><title>Temporal colinearity within a constitutive framework of long-range interactions</title><p>In different developmental contexts, the transcriptional activity of <italic>Hoxd</italic> genes coincides with an overall remodeling of long-range chromatin interactions with the flanking gene deserts, which harbor essential enhancer elements active in these developing tissues (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>; <xref ref-type="bibr" rid="bib4">Berlivet et al., 2013</xref>; <xref ref-type="bibr" rid="bib7">Delpretti et al., 2013</xref>). Colinear activation of <italic>Hoxd</italic> genes along the developing trunk is thought to primarily rely on regulatory influences intrinsic to the gene cluster itself (<xref ref-type="bibr" rid="bib48">Spitz et al., 2001</xref>). However, and even though their importance remains unclear, contributions of the flanking regulatory landscapes in this process have been proposed (<xref ref-type="bibr" rid="bib53">Tschopp et al., 2009</xref>; <xref ref-type="bibr" rid="bib52">Tschopp and Duboule, 2011b</xref>). Therefore, we assessed whether or not the reported changes in local interactions are associated with variations in long-range contacts during temporal colinearity, as was observed during limb and intestinal development (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>; <xref ref-type="bibr" rid="bib7">Delpretti et al., 2013</xref>).</p><p>By using a recently developed analytical methodology (<xref ref-type="bibr" rid="bib57">Woltering et al., 2014</xref>), we found that all interrogated <italic>Hoxd</italic> genes displayed substantial interactions with the flanking gene deserts (<xref ref-type="fig" rid="fig5">Figure 5A</xref>, <xref ref-type="fig" rid="fig5s1">Figure 5—figure supplement 1A</xref>). The quantification of interactions over both the centromeric and telomeric gene deserts revealed a gene-specific interaction preference towards either one or the other desert (<xref ref-type="fig" rid="fig5">Figure 5B</xref>), similar to what was previously described in limb bud cells (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>). However, in marked contrast, the dynamics of these long-range chromatin interactions were moderate, if any, and no clear modification in the contact profiles were detected between the inactive state in ES cells, and the subsequent transcriptional activation (<xref ref-type="fig" rid="fig5">Figure 5B,C</xref>). Hierarchical clustering of global patterns of long-range interactions revealed that the <italic>Hoxd4</italic>, <italic>Hoxd9</italic> and <italic>Hoxd11</italic> viewpoints systematically cluster together, whereas the <italic>Hoxd13</italic> viewpoint always behaves as outlier (<xref ref-type="fig" rid="fig5s1">Figure 5—figure supplement 1B</xref>).<fig-group><fig id="fig5" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.017</object-id><label>Figure 5.</label><caption><title>Sequential <italic>Hoxd</italic> gene activation occurs without drastic remodeling of long-range interactions.</title><p>(<bold>A</bold>) Distribution of long-range contacts in both the centromeric and telomeric gene deserts surrounding the <italic>HoxD</italic> cluster. Smoothed 4C-seq signals (11 fragment window size) are shown for the <italic>Hoxd13</italic> and <italic>Hoxd4</italic> gene viewpoints in ES and E9.5 tail bud cells. The analyzed genomic interval is the same as in <xref ref-type="bibr" rid="bib57">Woltering et al. (2014)</xref>. The location of topological domains (TADs) in ES cells are obtained from <xref ref-type="bibr" rid="bib11">Dixon et al. (2012)</xref> and indicated on the top with the <italic>HoxD</italic> cluster and both the centromeric and telomeric gene deserts indicated by arrows. The dashed lines demarcate the domain of high signal over the <italic>HoxD</italic> cluster, which is excluded from the analysis. (<bold>B</bold>) Summaries of the distributions in long-range signals within the centromeric and telomeric gene deserts surrounding the <italic>HoxD</italic> cluster, for all <italic>Hoxd</italic> genes assayed at various stages of their sequential activation. Each <italic>Hoxd</italic> gene specifically interacts with either the centromeric or the telomeric gene desert and these privileged contacts remain largely invariant during transcriptional activation. (<bold>C</bold>) Cumulative signals over the centromeric and telomeric gene deserts and the <italic>HoxD</italic> cluster for all <italic>Hoxd</italic> genes assayed at various stages of their sequential activation.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.017">http://dx.doi.org/10.7554/eLife.02557.017</ext-link></p></caption><graphic xlink:href="elife02557f005"/></fig><fig id="fig5s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.018</object-id><label>Figure 5—figure supplement 1.</label><caption><title>Temporal colinearity occurs without dynamic long-range interactions.</title><p>(<bold>A</bold>) Distribution of long-range contacts in the centromeric and telomeric gene deserts surrounding the <italic>HoxD</italic> cluster. Smoothed 4C-seq signals (11 fragment window size) for the indicated <italic>HoxD</italic> viewpoints either in ES (orange), E8.5 pre-somitic mesoderm (cyan), E9.5 tail bud (brown) or E10.5 tail bud (purple) cells over the same genomic interval as analyzed in <xref ref-type="bibr" rid="bib57">Woltering et al. (2014)</xref>. Genomic location of the <italic>HoxD</italic> cluster and surrounding genes is indicated below. TADs observed in ES cells (from <xref ref-type="bibr" rid="bib11">Dixon et al. 2012</xref>) are indicated on the top. The positions of both the <italic>HoxD</italic> cluster and the centromeric and telomeric gene deserts are indicated by arrows. The dashed lines demarcate the domain of high signals over the <italic>HoxD</italic> cluster, which is excluded from the analysis. (<bold>B</bold>) Hierarchical clustering of global patterns of long-range interactions in the surrounding gene deserts, for <italic>Hoxd</italic> viewpoints in ES cells and at different stages of sequential <italic>Hox</italic> gene activation. The <italic>Hoxd4</italic>, <italic>Hoxd11</italic> and <italic>Hoxd13</italic> viewpoints are consistently clustered together, with the <italic>Hoxd13</italic> behaving as an outlier. The correlations between samples (indicated by heatmaps) were calculated using Spearman's ranking of smoothed 4C-seq signals (11 fragment window size) over the combined genomic intervals as used in <xref ref-type="bibr" rid="bib57">Woltering et al. (2014)</xref>, with the <italic>HoxD</italic> cluster itself excluded. The samples were subsequently clustered (top) according to standard hierarchical clustering. (<bold>C</bold>) Hierarchical clustering of global patterns of long-range interactions in the surrounding gene deserts for <italic>Hoxd</italic> viewpoints in autopod (digits) and zeugopod (limbs) cells. Data are from <xref ref-type="bibr" rid="bib57">Woltering et al. (2014)</xref>.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.018">http://dx.doi.org/10.7554/eLife.02557.018</ext-link></p></caption><graphic xlink:href="elife02557fs010"/></fig><fig id="fig5s2" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.02557.019</object-id><label>Figure 5—figure supplement 2.</label><caption><title>Comparison between HiC and 4C-seq datasets obtained in ES cells.</title><p>(<bold>A</bold>) Virtual 4C carried out from HiC datasets, using bins covering the indicated <italic>Hoxd</italic> genes as viewpoints. Bins used as viewpoints are indicated in red. The interactions with bins covering the surrounding centromeric and telomeric TADs are given in light orange. TADs in ES cells (obtained from <xref ref-type="bibr" rid="bib11">Dixon et al. 2012</xref>) are indicated on the top and the location of the <italic>HoxD</italic> cluster is indicated below. The dashed lines demarcate the assigned TAD boundaries in ES cells. (<bold>B</bold>) Comparison of the distribution of long-range signals in both the centromeric and telomeric gene deserts, as obtained either by virtual 4C (light orange, data from <xref ref-type="bibr" rid="bib11">Dixon et al. 2012</xref>) or by 4C-seq (bright orange, this study). Despite large differences in both the size of the viewpoints and the resolution, the distribution is largely similar. The distribution of the HiC bin covering the promoters of the <italic>Hoxd13</italic> and <italic>Hoxd11</italic> genes behaves as a mix of the two individual 4C-seq viewpoints. (<bold>C</bold>) Coordinates of the centromeric and telomeric TADs surrounding the <italic>HoxD</italic> cluster (from <xref ref-type="bibr" rid="bib11">Dixon et al. 2012</xref>). (<bold>D</bold>) Detailed location of the HiC bins covering the <italic>HoxD</italic> cluster. (<bold>E</bold>) 4C-seq and virtual 4C patterns obtained when using a viewpoint covering the regulatory region CNS39 (<xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>), within the telomeric gene desert. In contrast to <italic>Hoxd</italic> gene viewpoints, the interactions observed with the centromeric gene desert are near background.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.019">http://dx.doi.org/10.7554/eLife.02557.019</ext-link></p></caption><graphic xlink:href="elife02557fs011"/></fig></fig-group></p><p>This clustering of interactions matches with the position of a previously mapped boundary between ‘topological associated domains’ (TADs; <xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>; <xref ref-type="bibr" rid="bib38">Nora et al., 2012</xref>). In ES cells indeed, two TADs cover approximately the gene deserts on either side and have their border at the level of the <italic>Hoxd12</italic>-<italic>Hoxd11</italic> genes (<xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>; <xref ref-type="fig" rid="fig5">Figure 5A</xref>, <xref ref-type="fig" rid="fig5s1">Figure 5—figure supplement 1A</xref>). Virtual 4C from HiC data with bins that cover the <italic>Hoxd</italic> genes show highly similar priming of interactions with the surrounding TADs when compared to our 4C-seq analysis, confirming that both approaches score similar chromatin behavior (<xref ref-type="fig" rid="fig5s2">Figure 5—figure supplement 2</xref>; <xref ref-type="bibr" rid="bib46">Sexton et al., 2012</xref> for analysis strategy). During limb development, genes located near this boundary (<italic>Hoxd9</italic> to <italic>Hoxd11</italic>) change their tropism and switch their contacts from one TAD to the other, such as to interact sequentially with the appropriate enhancers (<xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>). This structural re-organization is clearly illustrated in our hierarchical clustering with <italic>Hoxd11</italic> changing its association from <italic>Hoxd4</italic> to <italic>Hoxd13</italic> (<xref ref-type="fig" rid="fig5s1">Figure 5—figure supplement 1C</xref>). During temporal colinearity, however, such structural re-organization is not observed and hence the stepwise transcriptional activation of <italic>Hoxd</italic> genes appears to occur within a largely constitutive framework of long-range interactions; genes up to <italic>Hoxd11</italic> interact mostly with the telomeric domain, either before or after their activation, and <italic>Hoxd13</italic> always interacts with the centromeric domain. This opposed tropism for <italic>Hoxd</italic> genes in ES cells, as revealed by HiC and the consequent TAD structures, is somewhat at odds with the local clustering of <italic>Hoxd</italic> genes when in a negative state, which we report here by using 4C. This paradox is discussed below.</p></sec></sec><sec id="s3" sec-type="discussion"><title>Discussion</title><p>In this study we present the dynamics of local and long-range 3D chromatin organization during temporal colinear activation of <italic>Hox</italic> genes in vivo. Prior to their sequential activation, <italic>Hox</italic> genes are organized into local 3D chromatin compartments that encompass all bivalently marked chromatin (<xref ref-type="bibr" rid="bib5">Bernstein et al., 2006</xref>). In ES cells, however, these 3D compartments appear less defined than the fully inactive and H3K27me3-only marked compartments observed in differentiated cells. This difference in discreteness was paralleled by the amount of observable background transcription, which was substantially higher in ES cells that in brain cells, though the biological impact of these very low level transcripts, if any, remains to be determined. This supports the view whereby a generally relaxed chromatin organization in ES cell may accompany the plasticity required for cell-faith commitment (<xref ref-type="bibr" rid="bib33">Mattout and Meshorer, 2010</xref>). Interestingly, in ES cells grown in the presence of two kinase-inhibitors and thought to be in a somehow more naïve developmental state, H3K27me3 marks are almost absent from <italic>Hox</italic> clusters (<xref ref-type="bibr" rid="bib32">Marks et al., 2012</xref>). Considering the high correlation between the presence of H3K27me3 marks and the existence of 3D chromatin compartments, we would anticipate <italic>Hox</italic> clusters within these ES cells not to group into 3D compartments as distinct as those visible in ‘canonical’ ES cells. We therefore hypothesize that structuring into fully inactive 3D compartments is a gradual process occurring over the course of several days during embryonic development.</p><sec id="s3-1"><title>The spatial dynamics of temporal colinearity</title><p>We also observe that the sequential transcriptional activation of <italic>Hox</italic> genes in the PSM coincides with a gene-by-gene transfer or positioning from the inactive H3K27me3-decorated compartment to a newly formed 3D compartment containing active genes only. This indicates that the presence of <italic>Hox</italic> genes in 3D compartments of various extents, along the developed body axis (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>) is not an a posteriori mechanism used to fix and secure the long-term maintenance of various states of activity, fixed earlier by ‘classical’ transcriptional regulations acting <italic>in trans</italic>. Instead, it suggests that such spatial structures are instrumental in the precise regulation of their transcriptional timing. This is observed at least for the <italic>Hoxd9</italic> to <italic>Hoxd13</italic> genes and we infer that the same process occurs in the part of the cluster containing from <italic>Hoxd1</italic> to <italic>Hoxd8</italic>. It is however not possible to assess this experimentally due to technical limitations associated with the size of the embryonic material at the corresponding developmental stages.</p><p>Temporal colinearity was originally proposed as a mechanism to translate time into spatial coordinates, in different ontogenic contexts (<xref ref-type="bibr" rid="bib12">Duboule, 1994</xref>; <xref ref-type="bibr" rid="bib24">Gerard et al., 1997</xref>; <xref ref-type="bibr" rid="bib17">Durston et al., 2012</xref>). While our results support this idea, the transcriptional timing associated with gene clustering may not be an absolute prerequisite to achieve the proper spatial distributions of <italic>Hox</italic> genes products, as suggested by the multiple cases where single mammalian <italic>Hox</italic> transgenes could largely recapitulate the major expression specificities along the AP-axis (<xref ref-type="bibr" rid="bib28">Krumlauf, 1994</xref>; <xref ref-type="bibr" rid="bib13">Duboule, 2007</xref>; <xref ref-type="bibr" rid="bib53">Tschopp et al., 2009</xref>; <xref ref-type="bibr" rid="bib51">Tschopp and Duboule, 2011a</xref>). In this context, it is possible that the progressive transition of <italic>Hox</italic> genes from an inactive to an active 3D compartment reflects the existence of a mechanism whose major aim would not be to precisely regulate a time sequence but instead, to protect the most ‘posterior’ <italic>Hox</italic> genes from a premature exposure to activating factors, a situation shown to block posterior elongation and hence to be detrimental to the embryo (<xref ref-type="bibr" rid="bib58">Young et al., 2009</xref>; <xref ref-type="bibr" rid="bib31">Mallo et al., 2010</xref>). The necessity to actively prevent the most posterior genes from premature activation is supported by their basal transcriptional activity in ES cells, where <italic>Hox</italic> clusters are less discrete than in subsequent negative tissues such as fetal brain cells. Such a basal activity, which was not scored in these latter cells, may reflect the rather generic nature of the activating signals, the general mechanism underlying temporal colinearity thus relying on de-repression.</p><p>Studies using internal <italic>Hox</italic> cluster deletions and duplications indeed showed that the relative position of <italic>Hox</italic> genes, rather than their promoters, determines their responses to activating signals (<xref ref-type="bibr" rid="bib53">Tschopp et al., 2009</xref>). In this view, graded signals emanating from the posterior aspect of the developing embryo would lead to a progressive de-repression of <italic>Hox</italic> clusters, implying that these clusters would display some directional sensitivity. While the nature of the activating factors is elusive, a link with the segmentation clock was proposed (<xref ref-type="bibr" rid="bib16">Dubrulle et al., 2001</xref>; <xref ref-type="bibr" rid="bib59">Zakany et al., 2001</xref>). Concerning the directional sensitivity, <italic>Polycomb</italic> group (Pc-G) gene products may play an important role in this process, as the distribution of H3K27me3 marks correlate with the size of the inactive 3D compartments. Recently, a somewhat graded distribution of both EZH2 and RING1B, two proteins members of the PRC2 and PRC1 complexes, respectively, was described over the <italic>HoxD</italic> cluster in ES cells, with the highest signals covering the most ‘posterior’ genes (<xref ref-type="bibr" rid="bib30">Li et al., 2011</xref>). Directionality may therefore derive from a weaker ‘anterior’ repression exerted by the Pc system. In this context, progressive alterations of the repressive system should sensitize the transcriptional threshold, while keeping on with directionality. This effect was observed in <italic>Cbx2</italic>−/− mutant embryos (a component of the PRC1 complex formerly known as M33), where the efficiency of the PRC1 complex was moderately decreased: RA treatment resulted in premature yet colinear activation of <italic>Hoxd</italic> genes (<xref ref-type="bibr" rid="bib3">Bel-Vialar et al., 2000</xref>). Alternatively, collinear activation may rely upon a different kind of model involving for example biophysical forces (<xref ref-type="bibr" rid="bib1">Almirantis et al., 2013</xref>). Future experiments where the process will be witnessed at the cellular level in real time may be informative in this context.</p></sec><sec id="s3-2"><title>Transcriptional maintenance</title><p><italic>Hox</italic> genes are originally activated in the most posterior aspect of the gastrulating embryo. This initial wave of activation seems to involve first a poised transcriptional status (<xref ref-type="bibr" rid="bib19">Forlani et al., 2003</xref>), followed by an apparent anterior forward spreading (<xref ref-type="bibr" rid="bib10">Deschamps and Wijgerde, 1993</xref>; <xref ref-type="bibr" rid="bib23">Gaunt and Strachan, 1994</xref>; <xref ref-type="bibr" rid="bib21">Gaunt, 2001</xref>), which will ultimately lead to the positioning and initiation of the expression domains in the pre-somitic mesoderm (PSM). The colinear processing of this early phase may involve preparatory modifications in the chromatin status, making the system poised for activation by factors emanating from posterior cells (<xref ref-type="bibr" rid="bib19">Forlani et al., 2003</xref>). In this view, the observed anterior forward spreading in expressing cells (<xref ref-type="bibr" rid="bib10">Deschamps and Wijgerde, 1993</xref>; <xref ref-type="bibr" rid="bib23">Gaunt and Strachan, 1994</xref>; <xref ref-type="bibr" rid="bib21">Gaunt, 2001</xref>) may reflect a prolonged exposure to low levels of signals diffusing from the posterior end of the primitive streak (<xref ref-type="bibr" rid="bib19">Forlani et al., 2003</xref>). A second (non-exclusive) possibility is that it illustrates the initial difficulty to maintain a robust boundary in Pc repression in a gene cluster where some anterior genes are fully active, with a tendency for the nearby-located genes to be de-repressed and activated.</p><p>However, our results suggest that once the expression is finally established within the PSM, the boundary between the active and inactive compartments remain rather stable for the next couple of days, until the axial skeleton is fully determined. In this view, these chromatin domains may represent part of the machinery used to fix a given state of activation and thus translate a temporal parameter into spatial coordinates. As such, early heterochronies in <italic>Hox</italic> gene activation within the PSM will lead to subsequent re-positioning of the expression boundary, as previously observed (<xref ref-type="bibr" rid="bib24">Gerard et al., 1997</xref>).</p></sec><sec id="s3-3"><title>Long-range contacts</title><p>By using genetic approaches, it was previously argued that the time-sequenced activation of <italic>Hoxd</italic> genes primarily uses regulatory influences located within the gene cluster itself (<xref ref-type="bibr" rid="bib48">Spitz et al., 2001</xref>), with some contributions coming from more distant flanking regions (<xref ref-type="bibr" rid="bib53">Tschopp et al., 2009</xref>; <xref ref-type="bibr" rid="bib52">Tschopp and Duboule, 2011b</xref>). We now report that such a transcriptional activation is implemented with little-if any-differences in the interaction profiles between the target genes and their neighboring gene deserts, unlike the situation observed during limb development where new contacts appear upon gene activation (<xref ref-type="bibr" rid="bib34">Montavon et al., 2011</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>). However, temporal colinearity does occur within a framework of constitutive long-range interactions, which may provide a scaffold helping the bimodal separation of active and inactive genes to take place. Further experiments with mice carrying large re-arrangements of these two gene deserts will be necessary to clearly weight the importance of flanking regions in the implementation of the <italic>Hox</italic> clock.</p><p>Finally, while the comparison between published HiC data (<xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>) and our 4C datasets are generally highly consistent (e.g., <xref ref-type="fig" rid="fig5">Figure 5</xref>, <xref ref-type="fig" rid="fig5s1">Figure 5—figure supplement 1</xref>; <xref ref-type="bibr" rid="bib2">Andrey et al., 2013</xref>), the data reported here using ES cells raise an apparent paradox. HiC analysis in ES cells identified a boundary between topological domains positioned around the <italic>Hoxd12</italic> to <italic>Hoxd11</italic> gene (<xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>; <xref ref-type="fig" rid="fig5s2">Figure 5—figure supplement 2D</xref>) and such boundaries are thought to impose or reflect a physical separation between the two interaction landscapes (e.g., <xref ref-type="bibr" rid="bib37">Nora et al., 2013</xref>). As a consequence, <italic>Hoxd13</italic> should display more interactions with its flanking gene desert than with the other part of the <italic>HoxD</italic> gene cluster. Yet, by using several viewpoints in a 4C set-up, the <italic>HoxD</italic> cluster in ES cells appears to form a single negative compartment, despite the interspersed presence of this TAD boundary (<xref ref-type="fig" rid="fig1">Figure 1</xref>). In fact, a detailed analysis of the HiC dataset reveals that the <italic>HoxD</italic> cluster itself forms a ‘micro-TAD’, displaying strong internal interactions, in agreement with the 4C results reported here. As such, we consider it likely that the TAD boundary identified by HiC in ES cells (<xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>) represents an average description of two distinct configurations (<xref ref-type="fig" rid="fig6">Figure 6</xref>). For each allele, either the most posterior <italic>Hoxd13</italic> gene forms stable interactions within the TAD on the centromeric side or, alternatively, the <italic>Hoxd11</italic> to <italic>Hoxd1</italic> genes interact with the TAD on the telomeric side (<xref ref-type="fig" rid="fig6">Figure 6</xref>). As a consequence, for each allele the entire <italic>HoxD</italic> 3D compartment becomes located towards a single TAD, on one side of the cluster with a physical separation from the other side. Molecule(s) causing these interactions are elusive and may include proteins that mediate constitutive loops between <italic>Hoxd</italic> gene promoters and their regulatory elements. The CTCF protein, which may play a role in scaffolding TADs, binds multiple sites around the <italic>Hoxd13</italic> to <italic>Hoxd8</italic> region (<xref ref-type="bibr" rid="bib18">Ferraiuolo et al., 2010</xref>; <xref ref-type="bibr" rid="bib41">Phillips-Cremins et al., 2013</xref>). Because formaldehyde crosslinking has a very short range of action (<xref ref-type="bibr" rid="bib39">Orlando et al., 1997</xref>), the system generates a graded pattern of 4C and HiC interactions from the location of the actual binding sites. Therefore, while in ES cells and for each allele, TAD borders are likely located at either side of the <italic>HoxD</italic> 3D compartment (<xref ref-type="fig" rid="fig6">Figure 6A</xref>, black lines), our analysis of a large population of cells reflects the equilibrium that exists between these two situations (<xref ref-type="fig" rid="fig6">Figure 6E</xref>). At later stages, when the <italic>HoxD</italic> cluster adopts a bimodal 3D organization, the tethering of interactions, as illustrated by the existence of TADs on either side, may help implement the separation between activated and repressed <italic>Hox</italic> genes, thereby potentially reducing deleterious regulatory interferences and premature activation of the most posterior <italic>Hox</italic> genes.<fig id="fig6" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.020</object-id><label>Figure 6.</label><caption><title>Model of dynamic bi-modal 3D compartmentalization during temporal colinearity.</title><p>(<bold>A</bold>) Schematic organization of topological domains in ES cells (from <xref ref-type="bibr" rid="bib11">Dixon et al. 2012</xref>) matching the centromeric and telomeric gene deserts, with an apparent boundary assigned near the <italic>Hoxd11</italic> gene (grey diagonal lines). All <italic>Hoxd</italic> genes in ES cells have considerable interactions on either side of the cluster, suggesting that this border is more diffuse and hence the entire <italic>HoxD</italic> cluster can be integrated in either TAD (diagonal black lines). (<bold>B</bold>) Various states of activity for <italic>Hoxd</italic> genes in different samples, analyzed during sequential activation. The assigned TAD boundary in ES cells is indicated by the dashed line. (<bold>C</bold>) Conceptual 2D representation of chromatin organization within the <italic>HoxD</italic> cluster chromatin compartment and surrounding centromeric and telomeric TADs in ES cells. (<bold>D</bold>) Schemes illustrating the dynamics of local 3D compartmentalization for the <italic>HoxD</italic> cluster (red and blue compartments) vs the constitutive nature of interactions in the context of the surrounding TADs during sequential activation. (<bold>E</bold>) A dynamic equilibrium to explain the paradox in the observed local vs long-range interactions. Genes located at the centromeric or telomeric extremities of the <italic>HoxD</italic> cluster form stable interactions with DNA sequences located with the flanking gene deserts, thereby dragging the <italic>HoxD</italic> 3D chromatin compartments into either one of the TADs. Within a cellular population, this process is in equilibrium, resulting in a read-out where <italic>Hoxd</italic> genes have a graded preference to interact with either the centromeric or the telomeric deserts, despite being organized into a single 3D chromatin compartment.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.020">http://dx.doi.org/10.7554/eLife.02557.020</ext-link></p></caption><graphic xlink:href="elife02557f006"/></fig></p></sec></sec><sec id="s4" sec-type="materials|methods"><title>Materials and methods</title><sec id="s4-1"><title>Animal care, tissue sampling, ES cell culture and sample preparation</title><p>All experiments were performed in agreement with the Swiss law on animal protection (LPA). Tissue samples were isolated at the indicated time points (with maximum 6 hr delay), with day E0.5 being noon on the day of the vaginal plug. Tissue pieces for 4C-sequencing, ChIP-sequencing, RNA-sequencing and Reverse Transcriptase-qPCR were isolated in PBS and subsequently transferred to PBS supplemented with 10% Fetal Calf Serum. 4C-seq and ChIP-seq material was incubated for 45 min with 1 mg/ml collagenase (Sigma-Aldrich, St. Louis, MO), and 4C-seq material was further made single cell using a cell strainer (BD Falcon).</p><p>Mouse ES cells were grown under feeder-free conditions on gelatinized plates in Dulbecco's modified Eagle's medium (DMEM, Life Technologies, Carlsbad, CA) supplemented with 17% fetal calf serum, 1 × non-essential amino acids (Life Technologies), 1 × Pen–Strep (Life Technologies), Sodium Pyruvate (Life Technologies), 0.1 mM β-mercaptoethanol, and 1000 U/ml LIF.</p><p>Embryonic 4C-seq samples consisted of pooled material from multiple embryos: 129 embryos for E8.5 pre-somatic mesoderm samples, 196 embryos for E9.5 tail bud, 143 embryos for E10.5 tail bud or E10.5 lumbo-sacral trunks and around 20 embryos for each E10.5 forebrain sample. For embryonic ChIP samples, 50 μg of chromatin was cross-linked at a time, of which 10 μg was used per ChIP. To obtain 50 μg of chromatin, 750 E8.5 pre-somatic mesoderm samples or 10 E10.5 forebrains were pooled. Total RNA from E10.5 forebrain was isolated from single embryos. ES cell 4C-seq and ChIP-seq samples were prepared from samples consisting of 20 million cells. 10 μg of ES cell chromatin was used per ChIP. Total RNA was isolated from 1 million cells.</p></sec><sec id="s4-2"><title>4C-sequencing</title><p>4C–seq libraries were constructed as previously described (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). NlaIII (New England Biolabs, Ipswich, MA) was used as the primary restriction enzyme and DpnII (New England Biolabs) was used as the secondary restriction enzyme. For each viewpoint, a total of 1 μg (E9.5 tail bud, E10.5 tail bud, E10.5 lumbo-sacral trunks, E10.5 forebrain and ES cells) or 50 ng (E8.5 pre-somatic mesoderm) of each 4C-seq library was amplified using 16 individual PCR reactions with inverse primers including Illumina Solexa adapter sequences (primer sequences in <xref ref-type="table" rid="tbl2">Table 2</xref>). Illumina sequencing was done on multiplexed samples, containing PCR amplified material of up to 7 viewpoints, using 100 bp Single end reads on the Illumina HiSeq system according to the manufacturer's specifications.<table-wrap id="tbl2" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.021</object-id><label>Table 2.</label><caption><p>4C-seq Inverse primer sequences</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.021">http://dx.doi.org/10.7554/eLife.02557.021</ext-link></p></caption><table frame="hsides" rules="groups"><thead><tr><th>Viewpoint</th><th>Inverse primer</th><th>Sequence</th></tr></thead><tbody><tr><td rowspan="4"><italic>Hoxd13</italic></td><td rowspan="2">iHoxd13 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAAAAATCCTAGACCTGGTCATG</td></tr><tr><td>chr2:74504328-74504348</td></tr><tr><td rowspan="2">iHoxd13 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGAGGCCGATGGTGCTGTATAGG</td></tr><tr><td>chr2:74505579-74505598</td></tr><tr><td rowspan="4"><italic>Hoxd11</italic></td><td rowspan="2">iHoxd11 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAAGCATACTTCCTCAGAAGAGGCA</td></tr><tr><td>chr2:74523621-74523643</td></tr><tr><td rowspan="2">iHoxd11 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGACTAGGAAAATTCCTAATTTCAGG</td></tr><tr><td>chr2:74523881-74523903</td></tr><tr><td rowspan="4"><italic>Hoxd9</italic></td><td rowspan="2">iHoxd9 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTACGAACACCTCGTCGCCCT</td></tr><tr><td>chr2:74536168-74536185</td></tr><tr><td rowspan="2">iHoxd9 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGACCCTCAGCTTGCAGCGAT</td></tr><tr><td>chr2:74536797-74536814</td></tr><tr><td rowspan="4"><italic>Hoxd4</italic></td><td rowspan="2">iHoxd4 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAAGGACAATAAAGCATCCATAGGCG</td></tr><tr><td>chr2:74561330-74561353</td></tr><tr><td rowspan="2">iHoxd4 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGATCCAGTGGAATTGGGTGGGAT</td></tr><tr><td>chr2:74562171-74562191</td></tr><tr><td rowspan="4"><italic>Hoxc13</italic></td><td rowspan="2">iHoxc13 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAGATAATTTTCCTGAGACATTGTAAC</td></tr><tr><td>chr15:102756108-102756132</td></tr><tr><td rowspan="2">iHoxc13 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGAGCTCAATGTTCCCTTCCCTAACG</td></tr><tr><td>chr15:102755251-102755273</td></tr><tr><td rowspan="4"><italic>Hoxb13</italic></td><td rowspan="2">iHoxb13 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAGGACTGTTCCTCGGGGCTAT</td></tr><tr><td>chr11:96057673-96057692</td></tr><tr><td rowspan="2">iHoxb13 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGAATCTGGCGTTCAGAGAGGCT</td></tr><tr><td>chr11:96057448-96057467</td></tr><tr><td rowspan="4"><italic>Hoxb9</italic></td><td rowspan="2">iHoxb9 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAAGATTGAGGAGTCTGGCCACTT</td></tr><tr><td>chr11:96136070-96136091</td></tr><tr><td rowspan="2">iHoxb9 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGATCATCAAACCAAGCAGGGCA</td></tr><tr><td>chr11:96136671-96136690</td></tr><tr><td rowspan="4"><italic>Hoxa13</italic></td><td rowspan="2">iHoxa13 forward<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTAACACTTGCACAACCAGAAATGC</td></tr><tr><td>chr6:52212211-52212232</td></tr><tr><td rowspan="2">iHoxa13 reverse<xref ref-type="table-fn" rid="tblfn1">*</xref></td><td>CAAGCAGAAGACGGCATACGAGGCGAGGCTCAGGCTTTTAT</td></tr><tr><td>chr6:52212476-52212495</td></tr><tr><td rowspan="4"><italic>CNS(39)</italic></td><td rowspan="2">iCNS(39) forward<xref ref-type="table-fn" rid="tblfn2">†</xref></td><td>AATGATACGGCGACCACCGAACACTCTTTCCCTACACGACGCTCTTCCGATCTATCCAAGGAGAAAGGTGTTGGTC</td></tr><tr><td>chr2:74975258-74975279</td></tr><tr><td rowspan="2">iCNS(39) reverse<xref ref-type="table-fn" rid="tblfn2">†</xref></td><td>CAAGCAGAAGACGGCATACGACAGGGCGTTGGGTCACTCT</td></tr><tr><td>chr2:74975670-74975687</td></tr></tbody></table><table-wrap-foot><fn><p>Location of primers according to NCBI37 (mm9).</p></fn><fn id="tblfn1"><label>*</label><p>Primers from Noordermeer D, Leleu M, Splinter E, Rougemont J, De Laat W, Duboule D. 2011. The dynamic architecture of <italic>Hox</italic> gene clusters. Science 334:222–225.</p></fn><fn id="tblfn2"><label>†</label><p>Primers from Andrey G, Montavon T, Mascrez B, Gonzalez F, Noordermeer D, Leleu M, Trono D, Spitz F, Duboule D. 2013. A switch between topological domains underlies <italic>HoxD</italic> genes collinearity in mouse limbs. Science 340:1234167.</p></fn></table-wrap-foot></table-wrap></p><p>4C-seq reads were sorted, aligned, and translated to restriction fragments using the 4C-seq pipeline of the BBCF HTSstation (available at <ext-link ext-link-type="uri" xlink:href="http://htsstation.epfl.ch">http://htsstation.epfl.ch</ext-link>; <xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>; <xref ref-type="bibr" rid="bib6">David et al., 2014</xref>) according to ENSEMBL Mouse assembly NCBIM37 (mm9). 4C-seq patterns were corrected vs previously generated random 4C–seq libraries (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>), consisting of BACs covering the mouse Hox clusters (<italic>HoxD</italic>: RP23-331E7; <italic>HoxC</italic>: RP23-430C12; <italic>HoxB</italic>: RP23-381I12 and RP23-196F5; <italic>HoxA</italic>: RP24-298M24). After random correction, three restriction fragments were removed that returned aberrant values (<italic>HoxD</italic>: chr2:74’597’000-74’597’732; chr2:74’608’796-74’609’312, <italic>HoxB</italic>: chr11:95’999’958-96’000’916) due to sequence abnormalities in the BAC template (confirmed by Sanger sequencing; not shown). Normalization and further data processing was done as previously described (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). Quantitative log2 ratios were calculated by dividing the quantitative fragment count between tissue samples. Unprocessed 4C-seq data is available from the Gene Expression Omnibus (GEO) repository under accession number GSE55344. Random corrected tracks are available from <ext-link ext-link-type="uri" xlink:href="http://duboule-lab.epfl.ch/data">http://duboule-lab.epfl.ch/data</ext-link>.</p><p>The directionality of long-range interactions was calculated as previously described (<xref ref-type="bibr" rid="bib57">Woltering et al., 2014</xref>). In <xref ref-type="fig" rid="fig5">Figure 5A</xref>, the smoothed 4C-seq patterns (running mean, window size 11) were obtained using the 4C-seq pipeline of the BBCF HTSstation (available at <ext-link ext-link-type="uri" xlink:href="http://htsstation.epfl.ch">http://htsstation.epfl.ch</ext-link>; <xref ref-type="bibr" rid="bib6">David et al., 2014</xref>). HiC data on topological associated domains (TADs) from ES cells were obtained from (<ext-link ext-link-type="uri" xlink:href="http://chromosome.sdsc.edu/mouse/hi-c/database.php">http://chromosome.sdsc.edu/mouse/hi-c/database.php</ext-link>; <xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>). Two TADs located centromeric and telomeric of the clusters were selected, covering genomic coordinates chr2:73400000-75960000 (discussed in <xref ref-type="bibr" rid="bib57">Woltering et al., 2014</xref>). Spearman correlation of long-range patterns was done over the region covering these TADs, with signal on the <italic>HoxD</italic> cluster itself removed (excluded region: chr2:74484971-74607492). Conventional hierarchical clustering was done to score for relationships between viewpoints.</p></sec><sec id="s4-3"><title>ChIP-sequencing</title><p>ChIP was performed as previously described (<xref ref-type="bibr" rid="bib36">Noordermeer et al., 2011</xref>). Cells were fixed for 5 min in a 2% formaldehyde solution at room temperature. ChIP-seq samples were fragmented to a range of 200–500 bp using tip sonication (Misonix S4000, Misonix, Farmingdale, NY), For all ChIP assays, 10 μg of cross-linked chromatin was used. Antibodies used: anti Histone H3K27me3 (#17-622; Millipore, Billerica, MA) and anti H3K4me3 (#17-614; Millipore). ChIP-seq libraries were constructed from 6 to 10 nanograms of immune-precipitated DNA according to the manufacturers instructions (Illumina, San Diego, CA). Sequencing was done using 50 or 100 bp Single end reads on the Illumina HiSeq system according to the manufacturer's specifications. ChIP-seq reads were mapped to ENSEMBL Mouse assembly NCBIM37 (mm9), and extended to 100 bp if read lengths smaller than 100 bp were used, using the ChIP-seq pipeline of the BBCF HTSstation (available at <ext-link ext-link-type="uri" xlink:href="http://htsstation.epfl.ch">http://htsstation.epfl.ch</ext-link>; <xref ref-type="bibr" rid="bib6">David et al., 2014</xref>). ChIP-seq data is available from the Gene Expression Omnibus (GEO) repository under accession numbers GSE55344 and GSE31570.</p></sec><sec id="s4-4"><title>Correlation of 4C-seq and ChIP-seq samples</title><p>Random corrected 4C-seq and ChIP-seq samples were correlated by ranking experimental values within restriction fragments (<xref ref-type="table" rid="tbl1">Table 1</xref>). First, to each NlaIII restriction fragment covered by the random 4C tracks within the regions visualized in <xref ref-type="fig" rid="fig1 fig2">Figures 1 and 2</xref> (<italic>HoxD</italic> cluster: chr2:74454783-74622413; <italic>HoxC</italic> cluster: chr15:102715179-102909417; <italic>HoxB</italic> cluster: chr11:95992344-96244915; <italic>HoxA</italic> cluster: chr6:52058584-52234371), the average ChIP-seq signal was assigned for each condition. Restriction fragment within individual samples were ranked based on their 4C-seq or ChIP-seq value and subsequently the Spearman's rank correlation coefficient was calculated between pairs of samples.</p></sec><sec id="s4-5"><title>RNA-sequencing and Reverse Transcriptase-qPCR</title><p>Total RNA from tissue samples was isolated using Trizol LS reagent (Life Technologies). Total RNA from ES cell samples was isolated using Trizol reagent (Life Technologies). For RNA-seq, the RNA was depleted from rRNAs and, subsequently, strand-specific total RNA-seq libraries were constructed according to the manufacturers instructions (Illumina). Sequencing was done using 50 bp Single end reads on the Illumina HiSeq system according to the manufacturer's specifications. RNA-seq reads were mapped to ENSEMBL Mouse assembly NCBIM37 (mm9) and translated into reads per gene (RPKM) using the RNA-seq pipeline of the BBCF HTSstation (available at <ext-link ext-link-type="uri" xlink:href="http://htsstation.epfl.ch">http://htsstation.epfl.ch</ext-link>; <xref ref-type="bibr" rid="bib6">David et al., 2014</xref>). RNA-seq data is available from the Gene Expression Omnibus (GEO) repository under accession numbers GSE55344. For RT-qPCR, cDNA was synthesized after DNAseI treatment (Life Technologies) using SuperScript III (Life Technologies) and oligo-dT primers (Life Technologies), using the manufacturer's instructions. For ES cells and E10.5 forebrain, 2 μg of RNA was used as input for the cDNA synthesis, for E10.5 posterior trunk 1 μg of RNA was used. Products were quantified by qPCR using EXPRESS SYBR GreenER mixes (Life Technologies) on a CFX96 PCR Detection System (BioRad, Hercules, CA). Sequences of intron-spanning primers are provided in <xref ref-type="table" rid="tbl3">Table 3</xref>.<table-wrap id="tbl3" position="float"><object-id pub-id-type="doi">10.7554/eLife.02557.022</object-id><label>Table 3.</label><caption><p>RT-qPCR primer sequences</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.02557.022">http://dx.doi.org/10.7554/eLife.02557.022</ext-link></p></caption><table frame="hsides" rules="groups"><thead><tr><th>Fragment</th><th>Primer</th><th>Sequence</th></tr></thead><tbody><tr><td>mRNA</td><td>mRNA Tubb2c forward<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>GCAGTGCGGCAACCAGAT chr2:25080064-25080081</td></tr><tr><td><italic>Tubb2c</italic></td><td>mRNA Tubb2c reverse<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>AGTGGGATCAATGCCATGCT chr2:25079711-25079730</td></tr><tr><td>mRNA</td><td>mRNA Tbp forward<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>TTGACCTAAAGACCATTGCACTTC chr17:15644342-15644365</td></tr><tr><td><italic>Tbp</italic></td><td>mRNA Tbp reverse<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>TTCTCATGATGACTGCAGCAAA chr17:15650497-15650518</td></tr><tr><td>mRNA</td><td>mRNA Hoxd13 forward<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>GGTGTACTGTGCCAAGGATCAG chr2:74507077-74507098</td></tr><tr><td><italic>Hoxd13</italic></td><td>mRNA Hoxd13 reverse<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>TTAAAGCCACATCCTGGAAAGG over intron boundry</td></tr><tr><td>mRNA</td><td>mRNA Hoxd9 forward<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>GCAGCAACTTGACCCAAACA over intron boundry</td></tr><tr><td><italic>Hoxd9</italic></td><td>mRNA Hoxd9 reverse<xref ref-type="table-fn" rid="tblfn3">*</xref></td><td>GGTGTAGGGACAGCGCTTTTT chr2:74537278-74537298</td></tr><tr><td>mRNA</td><td>mRNA Hoxd4 forward</td><td>TCAAGCAGCCCGCTGTGGTC chr2:74565709-74565728</td></tr><tr><td><italic>Hoxd4</italic></td><td>mRNA Hoxd4 reverse</td><td>TCTGGTGTAGGCCGTCCGGG chr2:74566355-74566374</td></tr><tr><td>mRNA</td><td>mRNA Hoxb13 forward</td><td>GTCCATTCTGGAAAGCAG chr11:96056334-96056351</td></tr><tr><td><italic>Hoxb13</italic></td><td>mRNA Hoxb13 reverse</td><td>AAACTTGTTGGCTGCATACT chr11:96057389-96057408</td></tr><tr><td>mRNA</td><td>mRNA Hoxb9 forward</td><td>GGCAGGGAGGCTGTCCTGTCT chr11:96133282-96133302</td></tr><tr><td><italic>Hoxb9</italic></td><td>mRNA Hoxb9 reverse</td><td>GCCAGTTGGCAGAGGGGTTGG chr11:96135938-96135958</td></tr></tbody></table><table-wrap-foot><fn><p>Location of primers according to NCBI37 (mm9).</p></fn><fn id="tblfn3"><label>*</label><p>Primers from Montavon T, Le Garrec JF, Kerszberg M, Duboule D. 2008. Modeling <italic>Hox</italic> gene regulation in digits: reverse collinearity and the molecular origin of thumbness. Genes Dev 22:346–359.</p></fn></table-wrap-foot></table-wrap></p></sec></sec></body><back><ack id="ack"><title>Acknowledgements</title><p>We thank the members of the Duboule labs in Lausanne and Geneva for useful discussion and are grateful to Joost Woltering for sharing autopod and zeugopod 4C-data. We thank Mylène Docquier, Christelle Barraclough, Céline Delucinge and Natacha Civic from the Geneva Genomics Platform and Jacques Rougemont from the Bioinformatics and Biostatistics Core Facility (BBCF) of the Ecole Polytechnique Fédérale (EPFL) in Lausanne for their assistance in generating and analyzing high throughput data. Computations were performed at the Vital-IT (<ext-link ext-link-type="uri" xlink:href="http://www.vital-it.ch">http://www.vital-it.ch</ext-link>) Center for high-performance computing of the SIB Swiss Institute of Bioinformatics using tools developed by the BBCF (<ext-link ext-link-type="uri" xlink:href="http://bbcf.epfl.ch">http://bbcf.epfl.ch</ext-link>).</p></ack><sec sec-type="additional-information"><title>Additional information</title><fn-group content-type="competing-interest"><title>Competing interests</title><fn fn-type="conflict" id="conf1"><p>The authors declare that no competing interests exist.</p></fn></fn-group><fn-group content-type="author-contribution"><title>Author contributions</title><fn fn-type="con" id="con1"><p>DN, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article</p></fn><fn fn-type="con" id="con2"><p>ML, Conception and design, Acquisition of data, Analysis and interpretation of data</p></fn><fn fn-type="con" id="con3"><p>PS, Acquisition of data, Analysis and interpretation of data</p></fn><fn fn-type="con" id="con4"><p>EJ, Conception and design, Acquisition of data</p></fn><fn fn-type="con" id="con5"><p>FC, Organized and planified the production of all the mice used, Conception and design</p></fn><fn fn-type="con" id="con6"><p>DD, Conception and design, Analysis and interpretation of data, Drafting or revising the article</p></fn></fn-group><fn-group content-type="ethics-information"><title>Ethics</title><fn fn-type="other"><p>Animal experimentation: All experiments were performed in agreement with the Swiss law on animal protection (LPA) under license 1008/3482/0 to DD.</p></fn></fn-group></sec><sec sec-type="supplementary-material"><title>Additional files</title><sec sec-type="datasets"><title>Major datasets</title><p>The following dataset was generated:</p><p><related-object content-type="generated-dataset" document-id="Dataset ID and/or url" document-id-type="dataset" document-type="data" id="dataro1"><name><surname>Noordermeer</surname><given-names>D</given-names></name>, <name><surname>Leleu</surname><given-names>M</given-names></name>, <name><surname>Duboule</surname><given-names>D</given-names></name>, <year>2014</year><x>, </x><source>Temporal dynamics and developmental memory of 3D chromatin architecture at Hox gene loci</source><x>, </x><object-id pub-id-type="art-access-id">GSE55344</object-id><x>; </x><comment>NCBI Gene Expression Omnibus with open, unrestricted access.</comment></related-object></p><p>The following previously published dataset was used:</p><p><related-object content-type="generated-dataset" document-id="Dataset ID and/or url" document-id-type="dataset" document-type="data" id="dataro2"><name><surname>Noordermeer</surname><given-names>D</given-names></name>, <name><surname>Leleu</surname><given-names>M</given-names></name>, <name><surname>Duboule</surname><given-names>D</given-names></name>, <year>2011</year><x>, </x><source>The Dynamic Architecture of Hox Gene Clusters</source><x>, </x><object-id pub-id-type="art-access-id">GSE31570</object-id><x>; </x><comment>NCBI Gene Expression Omnibus with open, unrestricted access.</comment></related-object></p></sec></sec><ref-list><title>References</title><ref id="bib1"><element-citation publication-type="journal"><person-group person-group-type="author"><name><surname>Almirantis</surname><given-names>Y</given-names></name><name><surname>Provata</surname><given-names>A</given-names></name><name><surname>Papageorgiou</surname><given-names>S</given-names></name></person-group><year>2013</year><article-title>Evolutionary constraints favor a biophysical model explaining hox gene collinearity</article-title><source>Current Genomics</source><volume>14</volume><fpage>279</fpage><lpage>288</lpage><pub-id pub-id-type="doi">10.2174/13892029113149990003</pub-id></element-citation></ref><ref id="bib2"><element-citation publication-type="journal"><person-group 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An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see <ext-link ext-link-type="uri" xlink:href="http://elifesciences.org/review-process">review process</ext-link>). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.</p></boxed-text><p>Thank you for sending your work entitled “Temporal dynamics and developmental memory of 3D chromatin architecture at <italic>Hox</italic> gene loci” for consideration at eLife. Your article has been favorably evaluated by a Senior editor and 3 reviewers, one of whom is a member of our Board of Reviewing Editors.</p><p>The Reviewing editor (Robb Krumlauf) and the other reviewers discussed their comments before we reached this decision. The combined consensus view is that in principle this is suitable for <italic>eLife</italic> but there are a number of major and minor issues that need to be addressed or considered before it is acceptable for publication in <italic>eLife</italic>. The Reviewing editor has assembled the following comments to help you prepare a revised submission.</p><p>In several recent papers this group, as well as others, have shown that the Hox loci are split in active and inactive domains, with these domains associating with either centromeric or telomeric gene desert flanking the locus. The current manuscript largely builds on these earlier findings, confirming that these domains also form during temporal colinearity, and adds more details to this emerging story.</p><p>In the current work the authors perform similar 4C studies to their published work, but analyze different tissues. Here the authors focus on exploring how the temporal dynamics of Hox gene activation along the antero/posterior axis correlate with the 3D architecture of the clusters. They show that inactive Hox genes are organized in a single topological domain, as shown in brain and ES cells. However, as the collinear activation of these genes takes place, they move one-by-one, instead of in groups from an inactive to an active bimodal 3D domain. The main result is that active Hox genes occupy the same domain, and have more constant interactions with the gene desert regions. Whether the move is the consequence of gene activation, or is part of the activation process itself remains an open question. These domains coincide, with regions decorated by or depleted of H3K27me3 repression marks, respectively. Interestingly, once this bimodal 3D organization is defined for a particular A/P position, it remains fixed as development progress. Finally, they show that these internal switches within the Hox clusters are independent of the external contact that Hox genes make with the flanking gene deserts, which remain largely invariant through development and correlate with the Topological domains (TADs) defined by HiC studies. This is interesting, as the authors previously showed that these interactions do change and contribute to Hox gene expression during limb development.</p><p>Overall the results confirm the previous work, but extend this to a temporal dimension during development. This is a very interesting descriptive work that introduces a temporal dimension to the process of chromatin 3D organization of the Hox loci during collinear activation. There are a number of major and minor issues that need to be addressed or considered before it is acceptable for publication in <italic>eLife</italic>.</p><p><italic>Major</italic> <italic>comments:</italic></p><p>1) The authors propose an equilibrium model to explain the apparently contradiction between the 4C-seq and the HiC data observed in ES cells. Looking at the 4C-seq data from all other developmental stages, it seems is that an equilibrium between TAD constrains and the Hox interactions within the cluster could be occurring in all tissues and stages. Thus, it seems that, within the Hox clusters (that it, not considering long-range contacts), the architecture restrictions imposed by the TADs are much stronger than the 3D rearrangements caused by the progressive transition from inactive to active domains during A/P specification. Indeed, for the external genes, the increment of the contacts seen during these transitions, although detectable, is rather small compared with the number of interactions identified at their corresponding side of the TAD. It seems that genes closer to the TADs (specially Hox11 genes) are the ones showing the larger changes in their contacts as they transit from an active to an inactive state. This resembles very much with what the authors have shown for long-range contacts of these same genes during the distal limb differentiation, suggesting a more permeability of the TAD border for genes in very close proximity. It would be interesting to plot the number of reads within the cluster that lie at each side of the TADs for each gene and at each stage and A/P position. This could help to determine the putative constrains imposed by TADs onto the 3D dynamics associated to Hox gene collinear activation. This should also be discussed with more detail.</p><p>2) Could it be that these strong restrictions imposed by TADs contribute to preventing the premature activation of the most posterior genes along the body axis, which is detrimental for proper development?</p><p>3) The authors need to more quantitatively analyze 4C data tracks in order to more rigorously define domains. The work depends on analysis of 4C data. One limitation of this data is that it is less suited for identification of domains than Hi-C, as only single view points are analyzed. Typically topological domains are more readily detectable, both quantitatively and statistically, by Hi-C or 5C, because those methods provide matrices of interaction in which domains are more readily, and rigorously, identified. The authors need to describe their 4C data more quantitatively when they infer domain boundaries: in the current manuscript it seems that domain boundaries are either taken from published Hi-C data (which is from a different tissue/cell type and thus may be incorrect in certain aspects, even though many domain features may be conserved), or are based solely on visual inspection of the data, which is not quantitative. Overall the visual inspection of the data is quite satisfying and domain boundaries are certainly visible yet I think the manuscript would be strengthened by including some statistical metrics to better define a 4C domain. This will also allow more quantitative detection of the precise boundary, which is a key factor in the current work, and would allow a more quantitative comparison with domains of histone modifications. The approach to correlate 4C data with histone modifications described in Table I is a start, but it does not show whether domain boundaries coincide. For instance, 4C signal often appear to extend beyond the domain of histone modification (e.g., <xref ref-type="fig" rid="fig3">Figure 3</xref>, Hoxd13 in E9.5 tail bud).</p><p>4) Topological domains are much more easily detected by Hi-C. It would be very informative to have high resolution (i.e., deeply sequenced) Hi-C maps for E8.5 PSMs, E10.5 Tail buds, and limb tissues. This would be a more useful Hi-C data set than the ES cells data from Dixon et al. If the authors have this type of analysis they should consider including it in the paper.</p><p>5) The section about memorizing chromatin configurations is hard to follow, as it requires visual comparison of 4C data tracks in <xref ref-type="fig" rid="fig4">Figure 4</xref> with those in <xref ref-type="fig" rid="fig3">Figure 3</xref>. Can the authors include a supplemental figure that displays these different profiles side-by-side (e.g., the E9.5 Tail bud tracks vs. the E10.5 Lumbo-sacral trum track and include a ratio plot)? The authors also need a more quantitative approach to compare such profiles, as the current comparison is based solely on visual inspections, and important differences between 4C profiles are rather subtle.</p><p>6) There are similarities of the model proposed by the authors in <xref ref-type="fig" rid="fig6">Figure 6</xref> and those with the work from Wendy Bickmore's laboratory where they have shown that the relocation of Hox clusters away from the chromosome territory (CT) occurs during their activation, or similarities to Spyros Papageorgiou's “cluster translocation model” hypothesis. In the former, the Hox complex is gradually excluded from CTs, which could correspond to the flanking TADs. In the latter, sequential activation of the complex would involve its translocation from a region of repression (chromatin) to activation where there would be greater access to upstream regulatory factors. Although, Spyros envokes morphogenic and electromotive (i.e., Coulomb force) components, nowhere do the authors mention these works in the content of their discussion or model.</p><p><italic>Minor</italic> <italic>points:</italic></p><p>1) The statement that ES cells have a less compact Hox locus conformation are difficult to support by just 4C data. FISH might address this if it is available. The authors may wish to soften their conclusion otherwise.</p><p>2) The explanation of the apparent paradox between Hi-C and 4C data discussed at the end of the paper is reasonable, but a more likely explanation is that Hi-C simply has a lower resolution (20Kb at best). The authors suggest an apparent paradox when comparing Hi-C data and 4C data: the Hi-C data suggest a TAD boundary around d12 in ES cells, whereas d13 interacts with the rest of the Hox locus as observed by 4C. Their interpretation could be true, but in my view a more simple explanation is that the resolution of Hi-C (20 Kb at best) may be too low, and in the Dixon et al. data the true boundary can easily be 20Kb to the right or the left. I think the argument that Hi-C detects a boundary, while 4C does not, because Hi-C represents the average of two distinct conformations is not valid: 4C should be equally detecting this same average.</p><p>3) In ES cells low basal expression of Hox genes is observed, and the locus appears to be in a single domain. The results certainly indicate that any local domain feature may be weaker in these cells as compared to forebrain. However, for the authors to conclude that this reflects a less compact organization they need FISH data. Further, can the authors rule out that the increased outside-the-domain interactions in ES cells are not due to increased random ligation in ES cells?</p><p>4) The authors find that genes switch autonomously between compartments. But it seems that d1-d8 act as a single domain, then D9-10-11 get included one by one, and d13 is always different (consistent with it being in a different domain based on Hi-C data). Perhaps the authors can either clarify this, or discuss this in the paper: do they think each gene is acting independently, or are there three groups (D1-D8; D9-11, D13)?</p><p>5) The data in <xref ref-type="fig" rid="fig5">Figure 5</xref> is puzzling: Hox d11 is switching domains within the Hox cluster from E8.5 to E10.5, yet it does have the same bias to interact with the telomeric desert as the d4 gene. Does this mean that d4 and d11 are really in the same larger topological domain at all time points, but that there is a change in sub-domain organizations? Cell type specific re-organization of the internal organization of topological domains has been proposed before (e.g., Phillips-Cremins et al. Cell 2013).</p><p>6) All results combined, one can imagine a constant global domain organization with temporal changes in sub-domains. It would be very informative to have high resolution (i.e., deeply sequenced) Hi-C maps for E8.5 PSMs, E10.5 Tail buds, and limb tissues. This would be a more useful Hi-C data set than the ES cells data from Dixon et al. I do realize this may be difficult given the amount of material that this may require, but it would be a very interesting dataset as it may reveal the stage-specific sub-domains as well as invariant larger domains in a single dataset, with potential changes in the larger domains in limb.</p><p>7) In the legend of <xref ref-type="fig" rid="fig6">Figure 6A</xref>, the authors indicate that the 'apparent boundary' near the Hoxd11 gene is indicated by grey lines but do they not mean the dashed blue line passing from <xref ref-type="fig" rid="fig6">figure 6A</xref> into 6B? Or are they referring to the grey shaded triangle projecting from the middle of Hoxd11 onto the cartoon of the HoxD complex below it?</p><p>8) In regards to the replicates presented in supplemental data to <xref ref-type="fig" rid="fig1">Figure 1</xref>, where the contacts within and outside the HoxD complex are represented as a graphical ratio between ES cells and forebrain cells: visually this conveys the difference in the 3D organization of the inactive Hox complex between the ES cells and forebrain cells, however between replicates there is surprising variation in the ratio. Would we not expect less variation in the produced ratios between replicates which are produced from pooled samples of ES cells and forebrain cells? In the absence of the associated chromatin marks, this makes it hard to argue that in ES cells the HoxD cluster is less defined than in forebrain cells. Furthermore, one would worry that it would be easy to pick data from one replicate over another to fit the hypothesis regarding the remodeling of the HoxD architecture during sequential gene activation.</p></body></sub-article><sub-article article-type="reply" id="SA2"><front-stub><article-id pub-id-type="doi">10.7554/eLife.02557.024</article-id><title-group><article-title>Author response</article-title></title-group></front-stub><body><p>Major comments:</p><p><italic>1) The authors propose an equilibrium model to explain the apparently contradiction between the 4C-seq and the HiC data observed in ES cells. Looking at the 4C-seq data from all other developmental stages, it seems is that an equilibrium between TAD constrains and the Hox interactions within the cluster could be occurring in all tissues and stages. Thus, it seems that, within the Hox clusters (that it, not considering long-range contacts), the architecture restrictions imposed by the TADs are much stronger than the 3D rearrangements caused by the progressive transition from inactive to active domains during A/P specification. Indeed, for the external genes, the increment of the contacts seen during these transitions, although detectable, is rather small compared with the number of interactions identified at their corresponding side of the TAD. It seems that genes closer to the TADs (specially Hox11 genes) are the ones showing the larger changes in their contacts as they transit from an active to an inactive state. This resembles very much with what the authors have shown for long-range contacts of these same genes during the distal limb differentiation, suggesting a more permeability of the TAD border for genes in very close proximity</italic>.</p><p>We do not fully understand this comment. In the limb, <italic>Hoxd11</italic> clearly shifts its contacts from the telomeric domain (long range) to the centromeric domain (long range too).</p><p>During trunk development, we show here that this shift does <italic>not</italic> occur and that long-range contacts do not change significantly. Instead, what change are the short-range contacts, with <italic>Hoxd11</italic> shifting from a negative (H3K27me3) ‘micro’-domain to a positive domain including 3’ located genes. This is the main conclusion of the paper in fact, i.e., that the bimodal regulation observed in limbs is NOT at work in the trunk, where a much more progressive mechanism operates, with genes moving one after the other from one domain to the other (and <italic>not</italic> from one TAD to the other...). We have tried to make this point clearer in the text.</p><p><italic>It would be interesting to plot the number of reads within the cluster that lie at each side of the TADs for each gene and at each stage and A/P position. This could help to determine the putative constrains imposed by TADs onto the 3D dynamics associated to Hox gene collinear activation. This should also be discussed with more detail</italic>.</p><p>We have done this exercise and we now show the results below (<xref ref-type="fig" rid="fig7">Author response image 1</xref>). We have added this figure to the rebuttal rather than to the ‘paper’ since the correct interpretation of this calculation is difficult, due to three effects occurring simultaneously (see figure legend), of which the influence of the region flanking the baits, which we exclude in the counting of the reads, can’t be reliably determined. Importantly though, the numbers generally suggest that dynamics within the cluster (even without taking into account changes in the regions flanking the baits) are considerably more prominent that changes in the TADs.<fig id="fig7" position="float"><label>Author response image 1.</label><graphic xlink:href="elife02557f007"/></fig></p><p><italic>2) Could it be that these strong restrictions imposed by TADs contribute to preventing the premature activation of the</italic> <italic>most posterior genes along the body axis, which is detrimental for proper development</italic>?</p><p>Yes, this is an interesting possibility and the last sentence of the Discussion already touched upon this subject. We have tried to make it more explicit and changed it to: “…the tethering of interactions, as illustrated by the existence of TADs on either side, may help implement the separation between activated and repressed <italic>Hox</italic> genes, thereby potentially reducing deleterious regulatory interferences and premature activation of the most posterior <italic>Hox</italic> genes.”</p><p><italic>3) The authors need to more quantitatively analyze 4C data tracks in order to more rigorously define domains. The work depends on analysis of 4C data. One limitation of this data is that it is less suited for identification of domains than Hi-C, as only single view points are analyzed. Typically topological domains are more readily detectable, both quantitatively and statistically, by Hi-C or 5C, because those methods provide matrices of interaction in which domains are more readily, and rigorously, identified. The authors need to describe their 4C data more quantitatively when they infer domain boundaries: in the current manuscript it seems that domain boundaries are either taken from published Hi-C data (which is from a different tissue/cell type and thus may be incorrect in certain aspects, even though many domain features may be conserved), or are based solely on visual inspection of the data, which is not quantitative. Overall the visual inspection of the data is quite satisfying and domain boundaries are certainly visible yet I think the manuscript would be strengthened by including some statistical metrics to better define a 4C domain. This will also allow more quantitative detection of the precise boundary, which is a key factor in the current work, and would allow a more quantitative comparison with domains of histone modifications. The approach to correlate 4C data with histone modifications described in Table I is a start, but it does not show whether domain boundaries coincide. For instance, 4C signal often appear to extend beyond the domain of histone modification (e.g.,</italic> <xref ref-type="fig" rid="fig3"><italic>Figure 3</italic></xref><italic>, Hoxd13 in E9.5 tail bud)</italic>.</p><p>We do not fully understand these issues and we feel that the reviewers may have been confused. First, we do not try and define any TAD by 4C in this paper (even though the precision of 4C in defining TAD boundaries by using multiplexed baits is <italic>much</italic> higher than by using 5C or Hi-C – see Andrey et al., <italic>Science</italic>, 2013). We introduce the TAD boundary as defined by Dixon et al. only for a matter of discussion. What we show is precisely that we do not fit a TAD boundary during sequential gene activation. Secondly, in <xref ref-type="fig" rid="fig3">Figure 3</xref> no data on histone modifications is shown and hence we hardly understand what the referee means by: <italic>For instance, 4C signal often appear to extend beyond the domain of histone modification (e.g.,</italic> <xref ref-type="fig" rid="fig3"><italic>Figure 3</italic></xref><italic>, Hoxd13 in E9.5 tail bud).</italic> Instead, we show ratios for the difference in 3D organization between the data sets. In fact, we use these ratio plots to determine the borders between ‘interaction domains’ (not TADs). We consider this controlled approach based on initial normalization and subsequent calculations of ratios (described in detail in the supplemental information of <xref ref-type="bibr" rid="bib36">Noordermeer et al, 2011</xref> <italic>Science</italic>) considerably more precise than <italic>‘...based solely on visual inspection of the data’</italic> to determine positions of borders in these highly heterogeneous early embryonic tissue samples. We have tried to improve the quality of the text related to this part to prevent readers being confused.</p><p><italic>4) Topological domains are much more easily detected by Hi-C. It would be very informative to have high resolution (i.e., deeply sequenced) Hi-C maps for E8.5 PSMs, E10.5 Tail buds, and limb tissues. This would be a more useful Hi-C data set than the ES cells data from Dixon et al. If the authors have this type of analysis they should consider including it in the paper</italic>.</p><p>We of course agree that HiC data in these tissues would be a very helpful addition to the existing body of data. Unfortunately, these data are not available mostly for two reasons. Our experience with HiC (and similarly 4C-seq) is that generating data from tissues is much more complicated than using cell lines, especially when <italic>very</italic> large pools of tissue samples need to be collected, frozen, and later on combined (as would be the case here). Thus far, we have therefore been unable to generate HiC data from embryonic tissue samples. Further illustrating this, the fact that all published HiC data sets known to us (Liebermann-Aiden 2009, Dixon 2012, Jin 2013, Nagano 2013, Naumova, 2013, Sofueva 2013, Zuin 2013) are from cultured or circulating cells. The only exception is a mouse cortex HiC dataset (Dixon 2012) that is of much poorer quality due to low unique read numbers.</p><p><italic>5) The section about memorizing chromatin configurations is hard to follow, as it requires visual comparison of 4C data tracks in</italic> <xref ref-type="fig" rid="fig4"><italic>Figure 4</italic></xref> <italic>with those in</italic> <xref ref-type="fig" rid="fig3"><italic>Figure 3</italic></xref><italic>. Can the authors include a supplemental figure that displays these different profiles side-by-side (e.g., the E9.5 Tail bud tracks vs. the E10.5 Lumbo-sacral trum track and include a ratio plot)? The authors also need a more quantitative approach to compare such profiles, as the current comparison is based solely on visual inspections, and important differences between 4C profiles are rather subtle</italic>.</p><p>We agree that the comparison is not very easy. This paper presents comparisons between several parameters (first between genes at X time points in Y tissues), then between the ‘same’ tissue at various time points. This is of course intrinsically difficult as we cannot guarantee that the cells indicated at T1 will be indeed those present in the tissue at T2. The major problem there is this that of the embryological lineage, which we approximate (there is not other way to do this more precisely). In this context, to have a precise comparison of the 4C profiles would not really help the case as this remains an approximation. Also, we did not want to display several times the same profile, even if it was to address different questions. We have tried to improve the text of this section too, to make it clearer for the reader.</p><p><italic>6) There are similarities of the model proposed by the authors in</italic> <xref ref-type="fig" rid="fig6"><italic>Figure 6</italic></xref> <italic>and those with the work from Wendy Bickmore's laboratory where they have shown that the relocation of Hox clusters away from the</italic> <italic>chromosome territory (CT) occurs during their activation […]</italic></p><p>We now refer to this model. However, the Bickmore laboratory has shown that the Hox clusters loop out the chromosome territory when activated, yet <italic>not</italic> following a sequential activation. The referees say: <italic>‘’In the latter, sequential activation of the complex would involve its translocation from a region of repression (chromatin) to activation where there would be greater access to upstream regulatory factors’’.</italic> We do not remember that this nice model involved a ‘sequential activation’. The cluster was either outside or inside, but this was not linked to progressive activation.</p><p>Also, the Bickmore lab showed that upon activation, the <italic>HoxD</italic> cluster loops out of its CT only along the developing trunk, but not in the early limb bud (Morey et al., Development 2007). We therefore consider it unlikely that looping from the CT is equivalent to TAD switching, which once again seems to occur only in limbs thus far.</p><p><italic>[…] or similarities to Spyros Papageorgiou's “cluster translocation model” hypothesis. In the former, the Hox complex is gradually excluded from CTs, which could correspond to the flanking TADs. In the latter, sequential activation of the complex would involve its translocation from a region of repression (chromatin) to activation where there would be greater access to upstream regulatory factors. Although, Spyros envokes morphogenic and electromotive (i.e., Coulomb force) components, nowhere do the authors mention these works in the content of their discussion or model</italic>.</p><p>We have added references to these pieces of work now, yet we do not see anything in our paper, which can be ‘explained’ or ‘enlightened’ by the latter model. We nevertheless mention the electromotive possibility now.</p><p>Minor points:</p><p><italic>1) The statement that ES cells have a less compact Hox locus conformation are difficult to support by just 4C data. FISH might address this if it is available. The authors may wish to soften their conclusion otherwise</italic>.</p><p>We did soften our wording and wherever the word ‘compact’ or ‘compaction’ was used, we now use the words ‘defined’, ‘discrete’ or ‘structuring’ instead:. We have also changed the text here and there to make it less explicit. The use of FISH to illustrate this point is certainly not appropriate, at least considering data available at this locus (Bickmore laboratory and our laboratory, unpublished). The level of resolution (the cluster is ca. 100 kb large) and the fact that in both cases some ‘compaction’ is observed makes it unlikely to give a decisive answer.</p><p><italic>2) The explanation of the apparent paradox between Hi-C and 4C data discussed at the end of the paper is reasonable, but a more likely explanation is that Hi-C simply has a lower resolution (20Kb at best). The authors suggest an apparent paradox when comparing Hi-C data and 4C data: the Hi-C data suggest a TAD boundary around d12 in ES cells, whereas d13 interacts with the rest of the Hox locus as observed by 4C. Their interpretation could be true, but in my view a more simple explanation is that the resolution of Hi-C (20 Kb at best) may be too low, and in the Dixon et al. data the true boundary can easily be 20Kb to the right or the left. I think the argument that Hi-C detects a boundary, while 4C does not, because Hi-C represents the average of two distinct conformations is not valid: 4C should be equally detecting this same average</italic>.</p><p>We agree with the reviewers that the current resolution of HiC is not sufficient to precisely define the TAD border, but the paradox that we refer to is of a different nature: the four sampled <italic>Hoxd</italic> genes in ES cells are located in the same local 3D compartment, yet their ratio between centromeric <italic>versus</italic> telomeric contacts is highly graded (<xref ref-type="fig" rid="fig5">Figure 5B</xref>, <italic>Hoxd13</italic> ∼70% of total TAD contacts in the centromeric TAD and <italic>Hoxd4</italic> ∼ 20% of total TAD contacts in the centromeric TAD – once more TADs as defined by <xref ref-type="bibr" rid="bib11">Dixon et al. 2012</xref>). Because the genes are in the same compartment (our 4C data), one would expect that the genes would act as a single interaction unit. Our preferred solution to this paradox remains therefore the solution we discuss in <xref ref-type="fig" rid="fig6">Figure 6E</xref>. We have tried to clarify this paradox further in the text and figure legend.</p><p>This piece of discussion does not change the meaning of our results and was not strictly necessary for the readers. However, the fact is that a ‘boundary’ is seen in the ES cells HiC datasets, which we do not really observe when using 4C, unlike many other instances where our 4C data perfectly matched the TADs boundaries (Montavon et al., <italic>Cell</italic> 2011; Andrey et al., <italic>Science</italic> 2013). We think this is worth discussing, even though we do not have a clear explanation for it.</p><p><italic>3) In ES cells low basal expression of Hox genes is observed, and the locus appears to be in a single domain. The results certainly indicate that any local domain feature may be weaker in these cells as compared to forebrain. However, for the authors to conclude that this reflects a less compact organization they need FISH data</italic>.</p><p>As mentioned under minor point 1, we have re-phrased all sentences including the word ‘compacted’. In our opinion, obtaining meaningful FISH data will be problematic, as current FISH data measured compaction between fully inactive <italic>versus</italic> partially activated clusters (Chambeyron et al, <italic>Genes Dev</italic> 2004 and Morey et al, <italic>Development</italic> 2007), which is equivalent to our single <italic>versus</italic> bimodal organizations. To compare two distinct states of ‘compaction’ is likely out of the current resolution of this methodology.</p><p><italic>Further, can the authors rule out that the increased outside-the-domain interactions in ES cells are not due to increased</italic> <italic>random ligation in ES cells?</italic></p><p>This issue of increased random ligation as mentioned by the reviewers is indeed a valid point. We have added a new figure (<xref ref-type="fig" rid="fig1s6">Figure 1–figure supplement 6</xref>) where we compare the amount of signal in the surrounding TADs <italic>versus</italic> the rest of chromosome 2 or 11 (with the <italic>HoxD/HoxB</italic> clusters themselves excluded). As may be appreciated, the signal outside the surrounding TADs is in fact higher in terminally repressed forebrain cells. The increased interactions in ES cells are therefore limited only to the surrounding TADs. This excludes the possibility that the effect is due to random ligation. The consequences of these data are now discussed in the text.</p><p><italic>4) The authors find that genes switch autonomously between compartments. But it seems that d1-d8 act as a single domain, then D9-10-11 get included one by one, and d13 is always different (consistent with it being in a different domain based on Hi-C data). Perhaps the authors can either clarify this, or discuss this in the paper: do they think each gene is</italic> <italic>acting independently, or are there three groups (D1-D8; D9-11, D13)?</italic></p><p>This is a very interesting issue indeed, which was briefly alluded to in the paper and which we tried to make clearer now. <xref ref-type="bibr" rid="bib50">Tschopp et al. (2012)</xref> reported that in the CNS, activation seems to work as a ‘two blocks’ strategy (gene 1 to 8 and genes 9 to 13 – the latter being part of the <italic>AbdB</italic> sub-group). This was one of the reasons why we carried out these experiments, to see if the chromatin ‘micro-domains’ were only two or alternatively, if they were progressively installed during trunk extension. We reached the conclusion that the latter situation is observed. Yet of course this is seen <italic>only</italic> in the d9 to d13 part of the cluster. To verify this on the anterior part would require thousands of dissections, <italic>a fortiori</italic> including cell sheets, which are not as well defined (homogenous) as in later embryos. For the moment, we think this experiment is not feasible. The referees are right in mentioning that our model may <italic>not</italic> apply to the anterior part (<italic>d1</italic> to <italic>d8</italic>) where sequential activation may <italic>not</italic> be paralleled by progressive chromatin transition. The text has been clarified accordingly.</p><p><italic>5) The data in</italic> <xref ref-type="fig" rid="fig5"><italic>Figure 5</italic></xref> <italic>is puzzling: Hox d11 is switching domains within the Hox cluster from E8.5 to E10.5, yet it does have the same bias to interact with the telomeric desert as the d4 gene. Does this mean that d4 and d11 are really in the same larger topological domain at all time points, but that there is a change in sub-domain organizations? Cell type specific re-organization of the internal organization of topological domains has been proposed before (e.g., Phillips-Cremins et al. Cell 2013)</italic>.</p><p>See under minor point 2. This is in fact the paradox we discuss in <xref ref-type="fig" rid="fig5">Figure 5</xref>. This is exactly the point; when negative, <italic>Hoxd11</italic> contacts the telomeric domain in long range, even though it does contact <italic>Hoxd13</italic>, which itself contacts the centromeric domain in long range. Considering that we have HiC data only for ES cells, we prefer to restrict the comparison to HiC <italic>versus</italic> 4C data in such ES cells. The only HiC data set from tissue (cortex HiC data set from <xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>) does not really call a border inside the <italic>HoxD</italic> cluster, but rather tends to fuse the centromeric and telomeric ES-cell TADs into a large single TAD centered around the <italic>HoxD</italic> cluster.</p><p><italic>6) All results combined, one can imagine a constant global domain organization with temporal changes in sub-domains. It would be very informative to have high resolution (i.e., deeply sequenced) Hi-C maps for E8.5 PSMs, E10.5 Tail buds, and limb tissues. This would be a more useful Hi-C data set than the ES cells data from Dixon et al. I do realize this may be difficult given the amount of material that this may require, but it would be a very interesting dataset as it may reveal the stage-specific sub-domains as well as invariant larger domains in a single dataset, with potential changes in the larger domains in limb</italic>.</p><p>As discussed under major point 4, we agree with the reviewers that this data would be very insightful. Yet we (and most likely others) have not been able to generate such data from embryonic tissues. Also, HiC data from E8.5 mouse embryos is not yet a reasonable target. Our estimate indeed would be that between 2’000 and 4’000 E8.5 PSM dissections would be required to generate a single HiC library.</p><p><italic>7) In the legend of</italic> <xref ref-type="fig" rid="fig6"><italic>Figure 6A</italic></xref><italic>, the authors indicate that the 'apparent boundary' near the Hoxd11 gene is indicated by grey lines but do they not mean the dashed blue line passing from</italic> <xref ref-type="fig" rid="fig6"><italic>figure 6A</italic></xref> <italic>into 6B? Or are they referring to the grey shaded triangle projecting from the middle of Hoxd11 onto the cartoon of the HoxD</italic> <italic>complex below it?</italic></p><p>The mentioned grey lines are the two diagonal lines in the triangle representing the HiC data. These lines originate from the same point as where the dashed line (which is grey as well) originates. To clarify this issue, we have rewritten the legend, which now reads: “(A) Schematic organization of topological domains in ES cells [from (<xref ref-type="bibr" rid="bib11">Dixon et al., 2012</xref>)] matching the centromeric and telomeric gene deserts, with an apparent boundary assigned near the <italic>Hoxd11</italic> gene (grey diagonal lines). All <italic>Hoxd</italic> genes in ES cells have considerable interactions on either side of the cluster, suggesting that this border is more diffuse and hence the entire <italic>HoxD</italic> cluster can be integrated in either TAD (diagonal black lines).”</p><p><italic>8) In regards to the replicates presented in supplemental data to</italic> <xref ref-type="fig" rid="fig1"><italic>Figure 1</italic></xref><italic>, where the contacts within and outside the HoxD complex are represented as a graphical ratio between ES cells and forebrain cells: visually this conveys the difference in the 3D organization of the inactive Hox complex between the ES cells and forebrain cells, however between replicates there is surprising variation in the ratio. Would we not expect less variation in the produced ratios between replicates which are produced from pooled samples of ES cells and forebrain cells? In the absence of the associated chromatin marks, this makes it hard to argue that in ES cells the HoxD cluster is less defined than in forebrain cells. Furthermore, one would worry that it would be easy to pick data from one replicate over another to fit the hypothesis regarding the remodeling of the HoxD architecture during sequential gene activation</italic>.</p><p>It is correct that quite some variation can be observed between the replicates. Partially, this is due to the commonly observed variation at the individual fragment level in 4C experiments, combined with the log2 scale that is used to depict ratios. Relatively small changes therefore already can appear quite prominent in the graph. However, we initially shared the concerns of the reviewers and thus included two more safeguards in the analysis:</p><p>1) To overcome the effect of signal variation at the single fragment level, we rather decided to assess the change within the <italic>Hox</italic> complexes <italic>versus</italic> outside the <italic>Hox</italic> complexes. Our rationale here was that if the changes were purely experimental, no regional difference would be expected. To further exclude variation at the single fragment level, we decided not to look at the value of the ratio, but rather just at the sign (positive or negative). We realize that in our previous manuscript, the figures with these data were too small. In the current <xref ref-type="fig" rid="fig1s5">Figure 1–figure supplement 5</xref>, the data can be now better appreciated. For all those viewpoints where we compare in-<italic>versus</italic>-out for ES cells versus brain cells, we find a significant regional difference. For all those viewpoints, we can therefore conclude that ES cells have a significantly different distribution <italic>versus</italic> forebrain cells, when assessing their signal within their <italic>Hox</italic> cluster <italic>versus</italic> outside their <italic>Hox</italic> cluster.</p><p>2) To further rule out experimental variation, the replicate samples (replicate 2) were generated as true technical replicates and processed simultaneously with each step done using the same batch or reagents. The resulting data set was unfortunately not of the same high quality as the data from replicates 1 and this is primarily reflected by a lower number of genomic fragments outside of the <italic>Hox</italic> clusters with signal. Overall, these data sets still displayed the same pattern of interactions (see <xref ref-type="table" rid="tbl4">Author response table 1</xref>), but with less significant p-values due to the fewer useful fragments in the analysis. This is in contrast to the comparison between replicates, which is generally not significantly different (or, in the case of the <italic>Hoxb13</italic> gene ES rep 1 <italic>versus</italic> ES rep 2: very marginally). Within the data set, we do realize there is one large outlier: the comparison between the <italic>Hoxd13</italic> replicate 1 <italic>versus</italic> replicate 2. Within the overall replicate 2 experiment, this particular viewpoint was most problematic. To keep consistency within the experiment, and not hand pick which data to include, we decided to maintain it. Importantly, despite it being significantly different, the differences between ES cells and forebrain cells are always much more significant.<table-wrap id="tbl4" position="anchor"><label>Author response table 1.</label><table frame="hsides" rules="groups"><thead><tr><th>View point</th><th>ES rep 1 vs FB rep 1</th><th>ES rep 2 vs FB rep 1</th><th>ES rep 1 vs FB rep 2</th><th>ES all vs FB all</th><th>ES rep 1 vs ES rep 2</th><th>FB rep 1 vs FB rep 2</th></tr></thead><tbody><tr><td><italic>Hoxd13</italic></td><td>9.92E - 21</td><td>1.70E - 11</td><td>7.63E - 08</td><td>2.21E - 34</td><td>0.59</td><td>1.09E - 04</td></tr><tr><td><italic>Hoxd9</italic></td><td>3.02E - 02</td><td>0.36</td><td>0.91</td><td>2.61E - 03</td><td>0.90</td><td>0.27</td></tr><tr><td><italic>Hoxd4</italic></td><td>1.90E - 09</td><td>0.015</td><td>0.014</td><td>1.36E - 11</td><td>0.78</td><td>0.11</td></tr><tr><td><italic>Hoxb13</italic></td><td>1.37E - 04</td><td>7.65E - 03</td><td>0.010</td><td>4.42E - 14</td><td>0.037</td><td>0.51</td></tr><tr><td><italic>Hoxb9</italic></td><td>2.75E - 07</td><td>0.019</td><td>0.012</td><td>2.21E - 09</td><td>0.20</td><td>0.25</td></tr></tbody></table><table-wrap-foot><fn><p>p-values of difference in distribution between replicate samples. ES: ES cells, FB: forebrain.</p></fn></table-wrap-foot></table-wrap></p><p>To further rule out that the difference in discretion is an experimental effect, we have analyzed the replicate data in the same way as done for <xref ref-type="fig" rid="fig1s6">Figure 1–figure supplement 6</xref> to <xref ref-type="fig" rid="fig2">figure 2</xref> (discussed in Minor point 3 of this rebuttal). This figure (<xref ref-type="fig" rid="fig8">Author response image 2</xref>) hints that an increase of non-specific interactions may be the cause of the poorer quality of the replicate 2 data set (for all rep 2 samples the ‘distal chromosome’ category is increased). Importantly though, for all viewpoints the ES cell samples have increased interactions in the TADs in comparison to the forebrain data sets. These data therefore further support our finding that the compartments on the <italic>Hox</italic> clusters are less discrete in ES cells than in terminally repressed forebrain cells.<fig id="fig8" position="float"><label>Author response image 2.</label><graphic xlink:href="elife02557f008"/></fig></p></body></sub-article></article> |