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Establishment of 3D chromatin structure after fertilization and the metabolic switch at the morula-to-blastocyst transition require CTCF

The eukaryotic genome is tightly packed inside the nucleus, where it is organized in 3D at different scales. This structure is driven and maintained by different chromatin states and by architectural factors that bind DNA, such as the multi-zinc finger protein CTCF. Zygotic genome structure is established de novo after fertilization, but the impact of such structure on genome function during the first stages of mammalian development is still unclear. Here, we show that deletion of the Ctcf gene in mouse embryos impairs the correct establishment of chromatin structure, but initial lineage decisions take place and embryos are viable until the late blastocyst stage. Furthermore, we observe that maternal CTCF is not necessary for development. Transcriptomic analyses of mutant embryos show that the changes in metabolic and protein homeostasis programs that occur during the progression from the morula to the blastocyst depend on CTCF. Yet, these changes in gene expression do not correlate with disruption of chromatin structure, but mainly with proximal binding of CTCF to the promoter region of genes downregulated in mutants. Our results show that CTCF regulates both 3D genome organization and transcription during mouse preimplantation development, but mostly as independent processes.


In brief
The genome is organized in a precise 3D structure by chromatin-bound factors such as CTCF. However, its implication in gene expression is unclear. Andreu et al. describe mainly independent roles of CTCF in structuring the genome and in regulating the expression of key metabolic genes necessary for early mouse development.

INTRODUCTION
Chromatin is not randomly organized inside the eukaryotic cell nuclei. At the nuclear scale, individual chromosomes occupy distinct spatial territories during interphase (Cremer et al., 2006;Cremer and Cremer, 2010). Within chromosomes, the cell genome is compartmentalized into regions with similar chromatin characteristics, termed A-and B-compartments. A-compartments are transcriptionally active, gene rich, and nuclease sensitive, whereas B-compartments are relatively gene poor, transcriptionally silent, and insensitive to nucleases (Lieberman-Aiden et al., 2009). At intermediate length scales, high-resolution chromatin interaction maps have revealed that the genome folds into distinct modules called topologically associating domains (TADs). TADs are sub-megabase-sized chromatin domains in which genomic interactions are strong but sharply depleted on crossing the boundary to the adjacent one (Dixon et al., 2012;Hou et al., 2012;Nora et al., 2012;Sexton et al., 2012). Finally, at a smaller scale, distal genomic regions can physically interact, forming DNA loop structures that bring distant regulatory elements in close proximity to their target promoters (Dowen et al., 2014;Rao et al., 2014).
During early embryogenesis, there is a massive reorganization of the chromatin. In mature mouse sperm, TADs and compartments are already established (Jung et al., 2017;Du et al., 2017;Ke et al., 2017). However, mature oocytes arrested in metaphase II stage of meiosis are depleted of both structures (Du et al., 2017;Ke et al., 2017), as happens in mitotic cells when chromosomes are highly condensed (Naumova et al., 2013). After fertilization, chromatin adopts a more relaxed state and the strength of TADs and compartments is highly reduced, although some weak TADs and interaction domains are detected (Du et al., 2017;Ke et al., 2017;Flyamer et al., 2017;Collombet et al., 2020). During the first cleavage stages, a slow re-establishment of TADs and compartments starts but it is not completed until the eight-cell stage. Moreover, the progressive maturation of chromatin structure is conserved in early animal development, as has also been observed in Drosophila (Hug et al., 2017), zebrafish (Kaaij et al., 2018), and humans (Chen et al., 2019).
A characteristic feature of more than 75% of the borders of TADs and 86% of anchoring points of smaller DNA loops in mammals is the presence of the CTCF architectural protein (Dixon et al., 2012;Rao et al., 2014;Phillips-Cremins et al., 2013). CTCF is a ubiquitously expressed and highly conserved 11-zinc-finger DNA-binding protein (Filippova et al., 1996). It has been shown to be involved in transcriptional regulation, chromatin insulation, genomic imprinting, X-chromosome inactivation, and higher-order chromatin organization (Klenova et al., 1993;Bell et al., 1999;Cuddapah et al., 2009;   and different combinations of Ctcf mutant alleles at eight-cell, morula, and blastocyst stages immunostained with CTCF antibody (green). Nuclei were stained with DAPI (gray). Scale bar, 25 mm. Embryos were obtained from three and two independent litters from Ctcf fl/À 3 Ctcf fl/À or Zp3-Cre;Ctcf fl/À 3 Ctcf fl/À crosses, respectively, for all time points. The total number of embryos analyzed at each stage were, for eightcell, nine control, four Z KO, six M KO, and five MZ KO; for morula, nine control, three Z KO, six M KO, and seven MZ KO; and for blastocysts, 10 control, three Z KO, seven M KO, and eight MZ KO. (B-D) Quantification of the total cell number per embryo of early (B; 24-42 cells), mid (C; 43-63 cells), and late (D; 64-80 cells) blastocyst stages for the different Ctcf genotypes. Each dot represents an individual embryo. Embryos were obtained from six and four (B), four and four (C) and three and two (D) independent litters from Ctcf fl/À 3 Ctcf fl/À or Zp3-Cre;Ctcf fl/À 3 Ctcf fl/À crosses, respectively. The reduced sample size of MZ embryos at the late blastocyst stage (D, n = 3) is due to reduced embryo viability. Horizontal bars represent means. ns, non-significant; ***p < 0.0005, **p < 0.005, and *p < 0.05 by Student's t test.
(E) Confocal images of control (Ctcf fl/À ), zygotic, maternal, and maternal-zygotic Ctcf mutants immunostained with anti-CDX2 (green) and anti-NANOG (cyan) antibodies. Nuclei were stained with DAPI (gray). Arrowheads indicate cells double positive for CDX2 and NANOG. Scale bar, 25 mm. Embryos were obtained from Dixon et al., 2012). Homozygous CTCF deletion results in early embryonic lethality (Fedoriw et al., 2004;Heath et al., 2008;Wan et al., 2008;Moore et al., 2012;Chen et al., 2019). In somatic cells, conditional knockouts (KOs) showed additional important roles for CTCF in cell-cycle progression, apoptosis, and differentiation (Heath et al., 2008;Soshnikova et al., 2010;Li and Lu, 2007;Arzate-Mejía et al., 2018). However, global depletion of CTCF in cell cultures leads to subtle and context-specific changes in gene expression (Zuin et al., 2014;Nora et al., 2017;Hyle et al., 2019;Kubo et al., 2021), although chromatin structure is affected with a reduction of TAD insulation and intra-TAD loop formation. However, we are still missing a clear understanding of the role of CTCF in the initial establishment of chromatin structure, and its impact on regulated gene expression.
Here, we have studied the requirement of CTCF protein during mouse preimplantation development, when 3D chromatin structure is radically reorganized after fertilization, by genetically eliminating the maternal, zygotic, or both contributions of CTCF. Our results show that CTCF is not necessary for the first phases of mouse preimplantation development, but, after blastocyst cavitation, it is absolutely required for developmental progression.

RESULTS
CTCF loss leads to embryonic lethality at the late blastocyst stage To eliminate maternal contribution of CTCF, we used females carrying a CTCF floxed allele (Heath et al., 2008) and the Zp3-Cre driver. This Cre driver is expressed during the growing phase of oocytes prior to the completion of the first meiotic division, and, when combined with a floxed allele, it will result in no mRNA or protein generated by the mother to be deposited in the oocyte as it is formed (Lewandoski et al., 1997). Both maternal and maternal-zygotic mutant embryos (M KO and MZ KO) develop up to the blastocyst stage, and they proliferate and undergo cavitation just as zygotic mutant embryos (Z KO) and controls ( Figure 1A). We confirmed the absence of CTCF protein by immunostaining ( Figure 1A), and subsequent image quantification ( Figure S1A). Quantitative differences at the eight-cell stage are subtle, although there is clear reduction when comparing M and MZ KOs. This may be due to the lower levels of CTCF protein we detect at the eight-cell stage that increase sharply at the morula stage, in agreement with previous observations on CTCF expression during preimplantation development (Olbrich et al., 2021). On the other hand, M KO morula and blastocyst stage embryos exhibit nuclear CTCF protein just as controls ( Figure S1A), showing that, as they are heterozygous for the deletion, zygotic transcription quickly rescues the lack of maternal CTCF protein. Finally, we confirmed the absence of CTCF protein in MZ KO embryos at all stages examined ( Figures 1A and S1A).
Despite being able to form blastocysts, Z and MZ KO embryos suffer a block in development some hours later ( Figure S1B). M KO instead develop normally ( Figure S1B), are born, and grow to adulthood without any apparent phenotype. We did not detect differences in the number of cells among genotypes at the early (24-42 cells; Figure 1B) or mid-blastocyst stage Figure 1C). However, in the late blastocyst stage (64-80 cells) we observed a striking reduction in the number of cells in Z and MZ KO embryos compared with wild-type (WT) and heterozygous embryos, or M KO embryos, respectively ( Figure 1D). While Z mutants show a similar median number of cells in late blastocysts compared with mid-blastocysts (40-45 cells; Figures 1C  and 1D), MZ mutants have even fewer cells at this later stage (approximately 30 cells, compared with approximately 50 at the mid-blastocyst stage; Figures 1C and 1D). It is true that we have a reduced sample size of MZ KO at the late blastocyst stage (n = 3, Figure 1D), but this is due to embryo viability, as we consistently obtain fewer than expected embryos of this genotype at this late stage. Next, we examined whether this growth arrest is due to a decrease in proliferation or to increased apoptosis, as the number of cells in mutant embryos decreases. We labeled proliferating cells in mid-blastocyst stage M (with a phenotype equivalent to WT embryos) or MZ KO embryos with phosphorylated histone H3, and apoptotic cells with TUNEL staining. However, MZ KO show no significant differences in proliferation or cell death compared with M KO (Figures S1C and S1D). As CTCF has been described to have a role in maintaining genome stability (Lang et al., 2017), we examined the degree of DNA damage in control and mutant blastocyst. Indeed, we found that there was an increase in gH2AX staining in nuclear foci in maternal-zygotic mutant compared with maternal mutants (Figures S1E and S1F). These results suggest that cells from the preimplantation embryo with no CTCF divide and grow normally but at a certain stage are unable to maintain cell viability, and show a moderate increase in DNA damage. Nevertheless, the exact reasons for embryo lethality remain unclear.
A possibility that could be occurring was that initial lineage specification in the mouse embryo was not taking place correctly, and that mis-specified cells were leading to embryo arrest. Therefore, we performed immunostainings to detect key factors involved in the commitment of the trophectoderm (CDX2) and the inner cell mass (SOX2 and NANOG) in midstage blastocysts. We observed no obvious change in the levels of CDX2 and SOX2 between genotypes, and these were correctly expressed in restricted patterns in the trophectoderm (TE) and the inner cell mass (ICM), respectively ( Figures 1E and 1F). Although the general levels of NANOG expression remained mostly unchanged and we do not observe differences in the number of cells expressing both CDX2 and NANOG across genotypes ( Figure S1G), there is a tendency for TE from mutants to express higher levels of NANOG (Figure S1H). Nevertheless, and despite these subtle changes in the expression of NANOG, overall we can conclude that there is no gross mis-specification of the TE and ICM in CTCF depleted blastocysts and that the first cell fate decision is correctly specified.
We next attempted to see if the second lineage-specification event, that taking place in the ICM to produce the epiblast and the primitive endoderm, characterized by the segregated expression of NANOG and GATA6 respectively (Chazaud et al., 2006), occurs properly. However, this takes place in the late blastocyst, when Ctcf mutants are abnormal and a much-reduced cell number ( Figures 1D and S1B), which could make interpretation of expression patterns of these two markers difficult. Therefore, we carried out staining for NANOG and GATA6 in mid to late blastocyst (64-80 cells), at a stage where there is still not a mutually exclusive pattern of expression of NANOG and GATA6, but rather a mix of single and double-positive cells for each of the markers (Plusa et al., 2008). We detect the presence of single-and doublepositive cells in MZ KO embryos ( Figure S1I), just as we do in M KO embryos. Our analysis suggests that the specification of primitive endoderm and epiblast initiates correctly in the absence of CTCF, but, due to embryo lethality, we do not know if full segregation of the two lineages is completed in the mutants.

Reorganization of early chromatin structure in Ctcf mutant embryos
We developed single-blastocyst Hi-C adapting the volumes and timings from the single-cell Hi-C protocol (Nagano et al., 2015). This was necessary, as at this stage (mid-blastocyst), mutant embryos are phenotypically normal and undistinguishable from littermates ( Figures 1A and 1C). We therefore needed to generate individual embryo libraries, genotype on the Hi-C sequence data, and then merge individual libraries for the analysis ( Figures S2A and S2B). To optimize the number of libraries generated, we used embryos from the cross of Ctcf fl/À ; Zp3-Cre tg/+ females with Ctcf fl/À males, which will produce 50% M KO and 50% of MZ KO (see STAR Methods). As we have shown before that M KO are normal and reach adulthood, we assumed these embryos would be equivalent to WT controls or to zygotic heterozygotes (Ctcf fl/À ). Nevertheless, we also included control CD-1 embryos in our study. Overall, we produced 40 high-quality single-blastocyst libraries (13 WT, 14 M KO, and 13 MZ KO; Figure S2C) with an average of 2.2 million valid contacts each (Table S1). Unsupervised clustering of our single blastocyst Hi-C datasets resulted in the classification of MZ KO libraries separately from WT and M KO (Figure 2A), validating our initial expectation that M KO would be equal to WT controls. Visual inspection of merged-bygenotype contact probability maps revealed a clear difference in chromatin organization in MZ KO blastocysts compared with their littermates (M KO) or control embryos (WT), presenting a less defined structure ( Figure 2B). Genome-wide comparison between WT and MZ KO showed a general reduction in insulation, which did not happen when comparing WT with M KO ( Figure 2C). By plotting the frequency of contacts in relation to the distance they span, we observed a noticeable increase in the MZ KO embryos of contacts in the 200 kb to 1 Mb range, which could be associated with TAD organization, as well as a decrease in con-tacts of 10-50 Mb, a genomic distance associated with compartmentalization ( Figure 2D). The observation of an increase in contacts in the TAD range does not imply a more structured genome, as it is accompanied by a decrease in insulation (Figure 2C) in the mutants, most probably due to more interactions taking place between adjacent TADs.
Analysis of compartment formation in MZ KO blastocysts revealed that compartments remain mainly unaffected compared with WT blastocysts. However, we detected a slight increased frequency of compartments changing their identity from B to A when comparing WT with MZ KO, as opposed to WT compared with M KO (4.01% versus 2.51%; Figures 2E and 2F). We also observed an overall reduction in the strength of compartments, as apparent from the less-defined checkerboard pattern in MZ KO embryos (Figures 2G and 2H), associated with an increased interaction between adjacent compartments in the case of CTCF loss ( Figure 2I).
TAD assembly in preimplantation development is partially dependent on CTCF CTCF-depleted blastocysts showed reduction in the number of TADs (2,898 TADs detected in WT, 2,915 in M KO, and 2,497 in MZ KO), leading to an increase in median size ( Figure 3A). We also observed a high degree of TAD reorganization, even in genomic loci that will not be active until later stages in development, such as the Wnt6-Epha4-Pax3 locus ( Figure  TADs also showed a decrease in their reproducibility score (a statistical measure of the robustness of a TAD; see STAR Methods) and in the density of intra-TAD contacts (number of interactions by TAD size) ( Figure 3A). Changes in insulation score (IS), a measure of the interactions passing across a given genomic position, at the TAD boundaries are less pronounced in MZ KO blastocysts ( Figure 3B), confirming a weaker insulation of these domains when CTCF is not present. Likewise, we detect a reduction in the difference of IS at CTCF peaks when using CTCF chromatin immunoprecipitation (ChIP) data from naive embryonic stem (ES) cells ( Figure 3C). Consistent with these results, clear differences in TAD strength are appreciated when data from TADs are stacked in metaplots, showing a greatly diminished signal in MZ KO compared with WT or M KO embryos ( Figure 3D).
Intriguingly, this decrease of insulation is independent of the presence of CTCF at the TAD borders ( Figure S3C). This would suggest that other effects, different from direct binding of CTCF to chromatin but triggered by its absence, contribute to the global loss of chromatin structure at these stages. These could be changes in the expression of other factors that would also be involved in genome organization. Otherwise, the global depletion of CTCF before the structuring of the genome at these early developmental stages might lead to an overall weakening of TADs and chromatin structure. We also observe a much higher number/proportion of TADs where the position of their borders is not stable in the MZ KO compared with the control ( Figure 3E), once more showing how 3D structure is disrupted in embryos that lack CTCF.
Interestingly, if we compare our results with those obtained in ES cells depleted of CTCF (Nora et al., 2017;Kubo et al., 2021), we observe similar loss of insulation and ectopic contacts across TAD boundaries upon CTCF depletion ( Figure 3F). However, as discussed above, in the embryo, this occurs independently of CTCF occupancy of TAD boundaries, which does not happen in ES cells. On the other hand, the minor but reproducible reduction in the strength of compartments we observe was also detected in ES and immune cells depleted of CTCF, where up to a 10% reduction in strength has been described (Nora et al., 2017;Stik et al., 2020).

Transcriptional changes in blastocyst-stage mutant embryos
We analyzed the transcriptional changes in single blastocysts when mutants for Ctcf still do not show any overt phenotype ( Figures 1A and 1C), although there is a measurable effect on chromatin structure. We studied the expression of control Article ll OPEN ACCESS mutants revealed an enrichment of genes involved in metabolic pathways (glycolysis, such as Gpi1 or Slca1; fatty acid metabolism, Acadsb and Acsl6; lysosomal enzymes for amino and nucleotide sugar metabolism, Neu1 and Hexa; oxidative phosphorylation, Ndufb7, Ndufb10, Ndufab1, Uqcrc1, or Ppa2) and DNA replication for downregulated genes, and protein homeostasis (the ribosome, various Rpl and Rps genes; the proteasome, Psmd12 or Sem1; and the spliceosome, Hspa or Snrpb) and cell cycle for upregulated genes ( Figure 4B; Table S3). Interestingly, we did not observe changes in lineage determination genes, with the exception of Nanog, which matches our previous observations with whole-mount immunohistochemistry ( Figures 1E, S1G, and S1H).

Binding to promoters of downregulated genes suggests that CTCF activates gene expression
To characterize the relationship of direct CTCF binding with changes in gene expression, we checked binding of CTCF, as identified by ChIP sequencing (ChIP-seq) in naive ES cells, along the loci of all genes expressed at the blastocyst stage. We used CTCF binding data from ES cells because of the lack of goodquality data in early mouse embryos. The observed conservation of CTCF binding in different cell types from the same organism and also across different species (Cuddapah et al., 2009;Martin et al., 2011) made us consider that the majority of CTCF peaks found in naive ES cells would also be present in the embryo. We first analyzed the distribution of CTCF in a 2-kb region surrounding the transcriptional start site (TSS), finding that downregulated genes have a much higher tendency to have CTCF bound at their promoter region than non-DEGs. We also observe this trend for upregulated genes, although to a lesser extent. Interestingly, there is hardly binding of CTCF to the TSS region of genes that are not expressed in the blastocyst ( Figure 4C). We confirmed these observations by quantifying the proportion of upregulated, downregulated, and non-DEGs with CTCF binding at their promoters that is enriched for both categories of DEGs ( Figure 4D). Interestingly, if we extend our search to other regions, we find that downregulated genes are enriched for the binding of CTCF to at least one intron ( Figure 4D). Overall, these results show that proximal binding of CTCF to promoters is associated with positive regulation of gene expression. The observation of enrichment for intron binding only in downregulated genes is intriguing and could indicate cooperative mechanisms involving short-range CTCF-mediated interactions.
If we examine intergenic regions, we find an opposite behavior: genes upregulated in mutant blastocyst are more likely to present binding of CTCF in upstream or downstream noncoding regions than downregulated or non-DEGs ( Figure 4D). This is suggestive of CTCF acting upon cis-regulatory elements located in these regions. This could be explained by upregulated genes being localized at regions with lower gene density, and therefore with larger intergenic regions associated with each gene, with more complex and numerous intergenic regulatory elements. However, upregulated genes on average have a similar distribution of intergenic distances than non-DEGs; therefore, such bias cannot account for the observed enrichment of CTCF at these regions. On the other hand, we observed that downregulated genes tend to be nearer to neighboring genes, and therefore in more gene-dense regions than non-DEG or upregulated genes ( Figure 4E).

Reorganization of chromatin structure has a limited impact on transcription
To assess the possible link between structural and transcriptional changes due to the lack of CTCF, we first analyzed changes in the IS surrounding the TSS of DEG. Genes that are downregulated in Ctcf mutant blastocysts show a trend for higher insulation at their TSS in WT conditions compared with those upregulated or with non-DEG ( Figure 5A). In order to quantify the extent of the change in insulation, we have calculated the difference in Delta IS (D-Delta IS, Figure 5A; see STAR Methods) around the TSS of expressed genes for WT, M KO, and MZ KO embryos. While there are no differences between different group of genes when comparing M KO with WT embryos, when we compare MZ KO with WT blastocysts, the changes in D-Delta IS are significantly higher between both up-and downregulated genes (right and left panels, Figure 5A) and non-differentially expressed (middle panel, Figure 5A; Figure S3D): However, these differences can be explained by increased insulation surrounding CTCF bound regions ( Figure 3C) as promoter regions of DEGs are more frequent bound by CTCF, most notably downregulated genes ( Figure 4C). In fact, differences in Delta IS are higher in DEGs that show binding of CTCF to their promoters ( Figures S4A and S4B).
We next examined the distribution of DEG in compartments, taking into account whether they are stable in MZ KO embryos compared with controls or change their identity from A to B and vice versa. We observe that genes that change their expression have a higher tendency to be localized in A-compartments and less in B-compartments compared with non-DEGs (Figure 5B). We hardly find DEGs located to compartments that change their identity from B to A. However, we see a slight enrichment of downregulated genes in compartments changing from A to B ( Figure 5B). We next analyzed whether DEGs are related to TAD stability. For this, we assigned each expressed gene to a TAD and examined whether it was preserved, or one or both boundaries were lost, when comparing controls with mutants. In this case, we could not detect differences between DEGs and non-DEGs regarding the behavior of the TAD that contains them upon CTCF loss ( Figure 5C).
These results are consistent with what we observe at specific loci. Slc2a1 gene, encoding the GLUT1 glucose transporter, is downregulated in MZ KO embryos ( Figure 4A), and is localized in a TAD that shows a displacement of one of its borders without affecting the expression of any the immediately neighboring genes. However, the rewiring of contacts at the sub-TAD level results in loss of interactions near Slc2a1, as can be appreciated in the Hi-C matrix and the virtual 4C ( Figure S4C). Also, this region contains DNAse I hypersensitive sites (DHSs) mapped in morulastage embryos (Lu et al., 2016), suggestive of the presence of open chromatin and cis-regulatory elements. The adjacent TAD is split in two in MZ KO blastocysts, but only two genes inside them have their expression affected. For example, Ppcs, which encodes an enzyme involved in coenzyme A biosynthesis, is inside one of the split TADs and is the only gene dysregulated (in this case upregulated) inside the TAD, although an increase in contacts is detected in the region ( Figure S4C). Similarly, if we focus on the Nanog locus, we find some pluripotency genes that, together with Nanog, are upregulated in MZ mutants (Dppa3, Gdf3; Figures 4A and S4D). They are in the same TADs that have their borders expanded and suffer a reshuffling of the local interactions in CTCF-depleted embryos ( Figure S4D). Curiously, in the same TAD and neighboring Nanog sits Slc2a3, encoding GLUT3, another glucose transporter, which does not change its transcriptional activity. Again, this region contains DHSs suggestive of the presence of putative cis-regulatory elements. Moreover, by searching for the presence of DHSs within 10 kb upstream the TSS or 10 kb downstream the TTS of expressed genes, we observed that both up-and downregulated genes were enriched in open regions compared with non-DEGs irrespective of their location in A-or B-compartments (Figure S4E). Together, these results suggest that, although lack of CTCF affects 3D genome structure at different scales, only local reshuffling of the contacts at the sub-TAD level can partially explain the changes in expression we observe.
The transcriptional switch of metabolic programs from the morula to the blastocyst is impaired in Ctcf mutants As CTCF has been suggested to have a role in the coordination of dynamic transitions in expression (Sams et al., 2016;Gomez-Velazquez et al., 2017), we investigated whether genes changing expression in the morula-to-blastocyst transition were more susceptible to being affected by the lack of CTCF. To do so, we complemented our transcriptomic analysis on blastocysts described above with single-embryo RNA sequencing (RNA-seq) at the early morula stage (16-20 cells; Figure 1A) of control, Z, and MZ mutants. Samples from different genotypes were not resolved by principal-component analysis ( Figure S5A), and the comparison of MZ KO with control morulae showed only 55 DEGs, most of them being pseudogenes ( Figure S5C; Table S2). Therefore, we observe that the loss of maternal and zygotic CTCF has little to no effect on transcription before the blastocyst stage, once more reinforcing the notion of its dispensability for the very first stages of mouse development.
We next used our single-embryo expression data from controls to examine the transcriptional changes occurring at the morulato-blastocyst transition. We identified 114 downregulated genes and 702 upregulated genes in this developmental transition (Figure 6A; Table S2). As expected, ICM and embryonic pluripotency genes (such as Carm1, Dppa1, Tdgf1, or Zic3) or TE genes (Cdx2, Eomes, Gata3, or Tfap2a) (Stirparo et al., 2021), are upregulated from the morula to the blastocyst ( Figure 6A; Table S2). However, as observed before ( Figures 1E and 1F), they do not show much change in Ctcf mutant blastocysts ( Figure S5F). KEGG analysis showed that several of the pathways and genes changing in the transition to blastocyst are dysregulated in the opposite way to that of MZ KO mutants (Figures 4B and 6B). Oxidative Article ll OPEN ACCESS phosphorylation, amino and nucleotide sugar metabolism, or lysosome pathways are upregulated from morula to blastocyst ( Figure 6B), while they are downregulated in MZ KO blastocyst compared with controls ( Figure 4B). Equally, genes related to the ribosome are downregulated in the blastocyst, but upregulated in Ctcf mutants (Figures 4A and 4B; 6A and 6B; Table S3).
If we perform a genome-wide comparison of the changes in gene expression of MZ KO blastocysts with the changes that occur during the morula-to-blastocyst transition, we observe an inverse correlation for both sets of DEGs ( Figure 6C). In fact, while we identify 127 genes that are downregulated in MZ KO compared with controls and also upregulated in the Article ll OPEN ACCESS morula-to-blastocyst transition, only 19 genes show the opposite trend (upregulated in MZ KO and downregulated in the morula-to-blastocyst transition; Figure S5E; Table S3). Moreover, clustering of metabolism and ribosomal genes previously identified as DEGs grouped together E2.5 controls and E3.5 MZKO, with both these showing a similar signature of expression ( Figures 6D and 6E). Overall, these results suggest that loss of Ctcf leads to a failure to correctly implement the changes in gene expression, mainly related to metabolism and protein homeostasis, which would normally occur when the morula progresses to the blastocyst stage.

DISCUSSION
The role of CTCF in structuring the genome in 3D is undisputed (Ong and Corces, 2014). Equally, the relationship between for chromatin structure and the regulation of gene expression, although subject to debate, is also clear (Ibrahim and Mundlos, 2020). The limited transcriptional effect of CTCF deletion, as observed for example in ES cells (Nora et al., 2017;Kubo et al., 2021), is therefore surprising. This could be the result of CTCF being necessary for the initial establishment of regulatory interactions along chromatin but not for the posterior maintenance of these contacts. Therefore, depletion of CTCF would not affect established transcriptional programs but could be needed for changes in gene expression associated with differentiation events (Gomez-Velazquez et al., 2017;Arzate-Mejía et al., 2018). In this view, 3D chromatin structure mediated by CTCF is not required for transcriptional homeostasis but is essential for cell state transitions that involve deployment of different expression programs.
We have explored this issue by taking advantage of the first stages of mouse development, starting from fertilization, when the maternal and paternal haploid genomes come together to establish the embryonic diploid genome. We find that maternal CTCF is not required for development or adult life, nor for the establishment of 3D chromatin structure or transcriptional programs in the preimplantation mouse embryo. Our results are in contradiction with previous studies (Wan et al., 2008;Moore et al., 2012), surely due to the different approaches used to deplete CTCF (microinjection of RNAi versus genetic deletion). Furthermore, we observe that CTCF is dispensable for the first stages of development, as the block we observe in zygotic and maternal-zygotic mutants only occurs after blastocyst formation. This lack of phenotype suggests that CTCF is not required in the first hours of development before the zygotic gene is expressed and protein produced. Another possibility is that maternal CTCF is rescued by the paternal contribution, as it has been shown that CTCF protein is present on the mature sperm genome, bound to a small number of specific sites (Jung et al., 2017). Nevertheless, this low amount of protein is not sufficient to establish correct chromatin structure in embryos lacking maternal and zygotic CTCF.
Despite limitations in resolution of the contact maps, we observe that complete depletion of CTCF in blastocyst nuclei generates a less organized chromatin structure with an overall reduction in the isolation of interaction domains. Compartment identity remains mainly unaffected, but we have found higher fre-quency of inter-compartment interactions that translates into a reduction of compartment strength. TADs do not completely disappear in mutant embryos, but again their strength is much reduced and there is a reorganization of domains due to more promiscuous genomic contacts. Indeed, the increase in the number of long-range contacts (200 kb to 1 Mb) suggests the formation of larger DNA loops, probably as a consequence of cohesin not encountering obstacles at CTCF binding sites during loop extrusion (Wutz et al., 2020;Hansen 2020).
The fact that interaction domains are established de novo during preimplantation development in the absence of CTCF, albeit with a strong reduction in domain isolation, argues that other complementary factors or processes must be taking place to initially structure the genome. Similar conclusions have been reached in other systems, as for example the human embryo (Chen et al., 2019). In this scenario, CTCF could be acting as a scaffolding factor that stabilizes and maintains 3D organization generated by other means. Another possibility is that 3D structure is inherited from pre-patterned chromatin present in the separate gamete haploid genomes. However, recent work has shown that the majority of chromatin domains present in the maternal or paternal genomes prior to fertilization are not preserved in the embryo (Collombet et al., 2020). At present, there are no clear candidates for this triggering factor of genome structure, and at this point we cannot rule out that it appears as a side effect of other processes involving genome dynamics, such as DNA replication, transcription (Pope et al., 2014;Du et al., 2017;Ke et al., 2017), or lamina-chromatin interactions (Borsos et al., 2019).
The absence of CTCF triggers changes in expression related to metabolism and protein homeostasis in the blastocyst, while, surprisingly, there are no changes yet at the morula stage. Interestingly, in the morula-to-blastocyst transition, there is a metabolic switch in the use of energy (Kaneko, 2016;Zhang et al., 2018) and a downregulation of ribosomal components . Therefore, changes in the expression of metabolic pathways and structural components of the ribosomes in this transition do not take place in the CTCF mutants, which we believe is the cause of the developmental block of late blastocysts. It is interesting to note that metabolic transitions are important in other developmental transitions in the early mouse embryos, as is the case of the 2C state (Rodriguez-Terrones et al., 2020).
On the one hand, we have not found a clear relationship between the transcriptional changes we observe in mutant embryos and the structural reorganization caused by CTCF absence. Changes in insulation or TAD organization are not translated into global changes of gene expression in those regions, suggesting that transcription is resilient to alterations in TADs. Although mutant blastocysts exhibit a rewiring of interactions, these local changes do not alter all the genes in a given region in the same way, indicating that there are other properties that influence the response of genes to modifications of the 3D structure of the genome. The different susceptibility of genes could depend on the proximity between promoters and their enhancers, enhancer-promoter intrinsic preferences (Zabidi et al., 2015;Yokoshi et al., 2020), or whether they are genes with dynamic or continuous and stable expression (Sams et al., 2016;Gomez-Velazquez et al., 2017;Stik et al., 2020 On the other hand, we see a link between CTCF binding at promoters and downregulation of the expression, suggesting another role of CTCF protein different from insulation. In fact, it has been shown that promoter-proximal CTCF can promote long-range enhancer-promoter contacts and facilitate gene activation (Kubo et al., 2021). It is noteworthy that, in the case of genes downregulated in mutants, we see enrichment of CTCF bound at introns together with promoters, suggestive of a more complex mechanism than transcriptional activation by binding to promoter regions.
In summary, we find that CTCF is necessary for a proper 3D organization of the chromatin after fertilization and for transcriptional regulation of the metabolic switch in the morula-to-blastocyst transition required for developmental progression beyond the blastocyst stage. A rather unexpected observation was that we did not see major disruption of lineage-specification events or the early expression of key factors, such as Sox2, Oct4, Cdx2, or Gata6, when comparing Ctcf mutant blastocysts with controls. Therefore, our mutant embryos, which show an impaired 3D genome structure, were able to properly progress through the first cell fate decision. This argues that the gene regulatory networks responsible for these early events are largely independent of chromatin organization and must rely on shortrange regulatory mechanism.
Limitations of the study A limitation of our study is the small amount of starting material we can obtain from the Ctcf mutants. The necessity to determine the genotype of individual embryos on sequence data from Hi-C or RNA-seq libraries generated from single embryos prevents us from being able to pool embryos of the same genotype to obtain a larger sample. A further limitation of our genetic models is the need to maintain and generate embryos from two separate breeding schemes, to obtain on one hand homozygous zygotic mutants and heterozygotes, and, on the other, maternal and maternal-zygotic mutants. Finally, our analyses come from a heterogeneous population, as at the blastocyst stage there are two distinct cell types, trophectoderm and ICM. Therefore, we have not been able to distinguish changes specific to one or another of these cell types caused by the loss of CTCF.

STAR+METHODS
Detailed methods are provided in the online version of this paper and include the following:

DECLARATION OF INTERESTS
embryos, Ctcf fl/individuals were inter-crossed. To obtain maternal and maternal-zygotic mutant embryos Zp3-Cre tg/+ ; Ctcf fl/females were crossed with Ctcf fl/males. In this way, all embryos will lack maternal contribution of Ctcf. As for the zygotic Ctcf deletion, 50% will be heterozygous (Ctcf fl/-) and 50% homozygous (Ctcf À/À ). Adults were genotyped by PCR of tail-tip DNA using primers detailed in Table S4. For preimplantation embryos, genotyping was performed directly on individually isolated embryos after antibody staining. The stage of embryos used is indicated in the text. This ranges from morula used for RNA-seq ( Figure S5) to blastocyst in all other experiments. No differences related to the sex of the embryos was expected, and therefore this was not determined.
Mice were housed and maintained in the animal facility at the Centro Nacional de Investigaciones Cardiovasculares (Madrid, Spain) in accordance with national and European legislation. Procedures were approved by the CNIC Animal Welfare Ethics Committee and by the Area of Animal Protection of the Regional Government of Madrid (ref. PROEX 196/14).
Cell lines R1 129 male mESCs were cultured in DMEM containing 10% fetal calf serum in the presence of LIF with MEK and GSK3 inhibitors (1 mM PD0325901, 3 mM CHIR99021), as previously described (Cuadrado et al., 2019). Plates were coated with 0.1% gelatin and no feeder cells were present. This mESC line is authenticated by the Mouse Genome Editing Core Unit at CNIO, where it is routinely used to generate mice.
Embryos where staged by counting the number of cells using DAPI staining, and subdivided at blastocyst stage in early (24-42 cells), mid (43-63 cells) and late (64-80 cells). Embryos where imaged on glass-bottomed dishes (Ibidi or MatTek) with a Leica SP5 or Leica SP8 confocal microscope.

Immunofluorescence quantification
For quantification of CTCF intensity, DAPI-stained nuclei were segmented using the StarDist deep-learning image segmentation method (Weigert et al., 2020) through ImageJ software (Schindelin et al., 2012). The mean CTCF was quantified within the segmented nuclei and averaged per each embryo.
For NANOG intensity quantification, IMARIS imaging software (v. 7.6.3) was used. Images were smoothed with Gaussian filter 3 3 3 3 1 and then nuclei were segmented in 3D reconstructions with the tool Spots detection based on DAPI, CDX2 and NANOG staining with a 7 mm isosurface. Spots were visually evaluated and corrected if necessary. Using spot-to-spot colocalization analysis, NANOG-CDX2 double-positive cells were selected, and mean fluorescence intensity was measured within the spots. NANOG intensity values for each blastomere were normalized to the brightest blastomere of each litter of embryos to minimize staining and confocal imaging variability. Statistical analyses were performed with GraphPad Prism v.8.4.3 (GraphPad Software, USA).
ChIP-seq 4x10 7 mESCs growing at 70% of confluence were washed with PBS, trypsinized, resuspended in 20 mL of growing media and crosslinked with 1% formaldehyde for 15 min at RT. After quenching with 0.125 M Glycine, fixed cells were washed twice with PBS containing 1 mM PMSF and protease inhibitors, pelleted and lysed in lysis buffer (1%SDS, 10mM EDTA, 50mM Tris-HCl pH 8.1) at 2x10 7 cells/ml. 10 7 cells equivalent to 40-50 mg of chromatin were used per immunoprecipitation reaction with 10 mg of antibody. Sonication was performed with a Covaris system (shearing time 20 min, 20% duty cycle, intensity 6, 200 cycles per burst and 30 s per cycle) in a minimum volume of 2 mL. From 6 to 15 ng of immunoprecipitated chromatin (as quantitated by fluorometry) were electrophoresed on an agarose gel and independent sample-specific fractions of 100-200 bp were taken. Adaptor-ligated library was completed by limited-cycle PCR with Illumina PE primers (10-12 cycles). DNA libraries were applied to an Illumina flow cell for cluster generation and sequenced on the Illumina HiSeq2500.
Alignment of sequences to the reference mouse genome (mm10) was performed using 'Bowtie2 0 (version 2.3.3.1) under default settings (Langmead and Salzberg, 2012). Duplicates were removed using Picardtools (version 2.13.2) and peak calling was carried out using MACS2 (version 2.1.1.20160309) after setting the qvalue (FDR) to 0.05 and using the '-extsize' argument with the values obtained in the 'macs2 predictd' step (Zhang et al., 2008). dangling-ends, errors, random breaks or duplicates, those experiments with more than 800.000 valid-pairs were kept. Then, the experiments with the same genotype were merged and normalized. Due to WT and MZ KO merged experiments had similar number of reads, the M KO merged experiment was subsampled for a better comparison among samples. For genotyping on sequence, we considered as MZ KO those embryos whose libraries contained no reads in the 22 kb of the deleted region chr8:105662421-105684451 at Ctcf locus (Heath et al., 2008), using as a control other two random regions of the same size chr8:122710142-122732172 and chr8:122710142-122732172. Contact probability as a function of genomic distance was calculated as an intra-chromosomal contact frequency distribution, using logarithmically increasing genomic distance bins.
TADs were identified by using 40 and 100 kb resolution vanilla-normalized and decay-corrected matrices as input to the TAD detection algorithm implemented in TADbit, with identical results. A 25 kb resolution was used for the TAD pile-up contact enrichment maps generated with the tool coolpup (Flyamer et al., 2020). A-B compartments were detected by calculating the first component of a principal-component analysis (PCA) of chromosome-wide matrices (100 kb) and assigning A compartments to the genomic bin with positive PCA1 values and high GC content. Conversely, B compartments were assigned to the genomic bin with negative PCA1 values and low GC content. The same results were obtained using 200 and 500 kb windows. The insulation score and Delta signal were computed using a custom python script following the same methodology as Crane et al., 2015, extracting 10 kb resolution raw matrices and using a sliding window of 100 kb x 100 kb. The Observed/Expected metaplots were created using the tool coolpup (Flyamer et al., 2020) and unbalanced matrices. The TAD reproducibility score is the number of times a TAD is called by the algorithm. As the algorithm of TAD calling is carried out 10 times, TAD reproducibility values range from 1 to 10. TADs with a score lower than 5 will be considered not robust and are likely to be undetectable in replicates or at different resolutions (Serra et al., 2017). In the case of TAD analysis, 25 kb resolution matrices were used with the rescale option to scale all the TADs to the same size. For TAD classification a border is considered stable if the bin border in one condition is one bin more or one bin less of the other. In TSS analysis, the resolution used was 10 kb and they are centered in the TSS of the genes. Juicebox software was used for Hi-C data visualization and virtual 4C (Robinson et al., 2018). In order to quantify the differences in Delta IS between upregulated, downregulated and no-differentially expressed genes, we have extracted for each gene and condition the change in Delta IS (D-Delta IS). This value is defined as the difference between the Delta IS 80 kb upstream from the TSS and 80 kb downstream (which are the minimum and maximum values in all Delta IS metaplots). Then we have measured the differences of this variable between MZ KO and WT and between M KO and WT datasets for the three group of genes to analyze if there are significant differences between them.

Compartment dynamics and strength
Using 100 kb resolution matrices normalized by coverage, we have considered a bin as A or B if the eigenvector is greater than 0.008 or less than À0.008 respectively. Then, for each chromosome, we have added the interactions between A-A, B-B or A-B bins only if the distance of the two bins is greater than 2 Mb. Finally, the compartmentalization strength is defined by the following formula (AA + BB)/AB. Lower values of this metric are related with more AB contacts or less AA and BB contacts, meaning a lack of proper compartmentalization.

QUANTIFICATION AND STATISTICAL ANALYSIS
Statistical analyses were performed with GraphPad Prism v.8.4.3 (GraphPad Software, USA). All details regarding statistical analysis can be found in figure legends for both main and supplementary figures. All data show means. Differences were considered statistically significant when p < 0.05.