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CTCF binding landscape in jawless fish with reference to Hox cluster evolution

Scientific Reportsvolume 7, Article number: 4957 (2017) | Download Citation

Abstract

The nuclear protein CCCTC-binding factor (CTCF) contributes as an insulator to chromatin organization in animal genomes. Currently, our knowledge of its binding property is confined mainly to mammals. In this study, we identified CTCF homologs in extant jawless fishes and performed ChIP-seq for the CTCF protein in the Arctic lamprey. Our phylogenetic analysis suggests that the lamprey lineage experienced gene duplication that gave rise to its unique paralog, designated CTCF2, which is independent from the previously recognized duplication between CTCF and CTCFL. The ChIP-seq analysis detected comparable numbers of CTCF binding sites between lamprey, chicken, and human, and revealed that the lamprey CTCF protein binds to the two-part motif, consisting of core and upstream motifs previously reported for mammals. These findings suggest that this mode of CTCF binding was established in the last common ancestor of extant vertebrates (more than 500 million years ago). We analyzed CTCF binding inside Hox clusters, which revealed a reinforcement of CTCF binding in the region spanning Hox1-4 genes that is unique to lamprey. Our study provides not only biological insights into the antiquity of CTCF-based epigenomic regulation known in mammals but also a technical basis for comparative epigenomic studies encompassing the whole taxon Vertebrata.

Introduction

The CCCTC-binding factor (CTCF), containing the C2H2 Zn finger-type DNA binding domains, plays a pivotal role in chromatin organization as an insulator1, 2. Recent studies have revealed that CTCF binds to two-part motifs consisting of core and upstream motifs3, 4, and that the deposition of CTCF binding sites topologically determines chromatin structure2, 5,6,7. In mammalians, it was shown that CTCF binding sites propagated through retrotransposition3, 8. So far, our knowledge on molecular functions of CTCF is confined mostly to mammalians, whereas CTCF orthologs have been identified in diverse bilaterian animals9. Remarkably, CTCF orthologs are not retained in some bilaterians, which were also revealed to lack clusters of Hox genes9, 10. Studies on mammals have elucidated the involvement of CTCF binding in the regulation of chromatin topology in Hox clusters2, 11, 12. These findings pinpoint the importance of understanding the mechanism of Hox gene regulation from an epigenomic viewpoint.

Lampreys, used in this study, belong to Cyclostomata, a taxon that diverged from the future gnathostome (jawed vertebrate) lineage more than 500 million years ago, and are characterized by the lack of the articulating jaw13. Whole genome sequencing and analysis was first carried out for the sea lamprey Petromyzon marinus and later for the Arctic lamprey Lethenteron camtschaticum (or Japanese lamprey L. japonicum)14, 15. These genomic studies revealed its unique features, such as programmed genomic rearrangement and peculiar characters of protein-coding sequences associated with highly biased base composition towards increased GC-content14, 16. As these features could be influenced by epigenomic regulation through modifications of chromatin or DNA, it is crucial to investigate epigenomic regulation in this animal group.

In this study we identified two lamprey CTCF homologs, including CTCF2, which was duplicated in the lamprey lineage independently from the other gene duplication that gave rise to CTCFL (also called Boris17). We performed, to our knowledge, the first ever analysis with chromatin immunoprecipitation and sequencing (ChIP-seq) for a jawless fish, enabled by antibody validation and ChIP protocol optimization. Our results indicate the establishment of the binding property of CTCF, mediated by the two-part motif, in the last common ancestor of cyclostomes and gnathostomes, and also detected a transposon-associated propagation of the CTCF binding site in the lamprey, as in mammals. Focusing on Hox clusters, we also compared CTCF binding patterns between lamprey and gnathostomes.

Results

Generating Gene Models for L. camtschaticum

In order to allow comprehensive gene search and related gene-based analyses, genome-wide prediction of protein-coding genes was performed for the L. camtschaticum genome assembly LetJap1.0, for which no established gene model was available. The program AUGUSTUS18 was trained to adapt to this species using conserved genes and was employed for inference of protein-coding exons by incorporating RNA-seq reads (Supplementary Table S1) of embryonic and tissue samples (as evidence of exons) and protein sequences of other species (as evidence of coding regions). The resultant set of 34,320 predicted genes was further refined to recover false-negative exons and genes and to bridge split exons to retrieve longer open reading frames. This resulted in the final set of 34,435 genes, designated as the gene model ‘GRAS-LJ’ (Supplementary Fig. S1 and Supplementary Table S2; also see Supplementary Materials and Methods for the detailed procedures).

Identification of Cyclostome CTCF Genes

A BLASTP search was performed to identify CTCF homologs in our gene model, using the human CTCF peptide sequence (NCBI NP_006556.1) as a query. This identified a component of GRAS-LJ, g14920.t1 (3621 bp) that is harbored by the scaffold KE994200.1 of the genome assembly LetJap1.0. The sequence was extended by using RNA-seq data to obtain the transcript sequence of 5781 bp that encompasses the putative full-length open reading frame (ORF), which is markedly longer [1207 amino acids (aa)] than that of already reported orthologs [human, 727 aa (NCBI NP_0065561); fruitfly, 818 aa (NCBI NP_648109)]. The gene from which this cDNA was derived was designated LjCTCF (NCBI KX830966).

Unexpectedly, our search for sequences similar to CTCF identified another one that was different from LjCTCF. This was based on the components of GRAS-LJ g14288.t1 and g23728.t1, predicted on the scaffolds KE994121.1 and APJL01108776.1 respectively, and the sequence was extended to 1475 bp with our RNA-seq data and 3′ RACE to cover its putative full-length ORF (396 aa). The gene from which this sequence was derived was designated LjCTCF2 (NCBI KX830967).

The identified LjCTCF and LjCTCF2 sequences were used to further identify their orthologs in sea lamprey (P. marinus) and inshore hagfish (Eptatretus burgeri). The deduced amino acid sequences of LjCTCF and LjCTCF2 contain eleven and eight Zn finger domains, respectively. Multiple sequence alignment revealed that the elongated stretch upstream to the N-terminus accounts for the large length of LjCTCF (Fig. 1a), and that similarity was not observed between the lamprey CTCF and CTCF2 sequences outside the Zn finger domains (Supplementary Fig. S2).

Figure 1
Figure 1

Properties of L. camtschaticum CTCF and CTCF2 genes. (a) Protein domain structures, in comparison with human CTCF and CTCFL. The Zn finger domains (ZF) were identified by MOTIF Search (http://www.genome.jp/tools/motif/). See SI Materials and Methods for detail procedures of LjCTCF and LjCTCF2 sequence determination by cDNA cloning. (b) Expression levels of LjCTCF (white) and LjCTCF2 (black) in embryos and adult tissues. FPKM, fragments per kilobase of exon per million mapped sequence reads. (c) Whole-mount in situ hybridization on stage 26.5 embryos. Riboprobes were designed in non-conserved regions downstream to the Zn finger domains to avoid cross-hybridization (for the magnified view and the result with upstream riboprobes, see Supplementary Fig. S10). Note that the difference in the intensities of the expression signals between the two genes do not correspond to that of actual expression levels, as indicated by our RNA-seq data in (b). Scale bars: 500 μm. (d) Molecular phylogenetic tree. This tree was inferred with the maximum-likelihood approach using 208 aligned amino acid sites (see Supplementary Tables S8 and S9 for the list of sequences used). Two stickleback sequences are included (upper, Ensembl ENSGACP00000003270; lower, ENSGACP00000020939). At each branch node in the tree, only bootstrap value of no less than 60, and the posterior probability inferred with the Bayesian approaches are shown.

Expression Patterns of Lamprey CTCF and CTCF2

Expression levels of LjCTCF and LjCTCF2 were quantified employing the aforementioned RNA-seq data and our gene model ‘GRAS-LJ’. Both LjCTCF and LjCTCF2 were expressed in all the tissues analyzed, while LjCTCF was expressed at a higher level (Fig. 1b). Their detailed embryonic expression patterns were analyzed with whole-mount in situ hybridization. This resulted in broad expression signals mainly in the craniofacial regions and axial structures, which is shared between the two genes (Fig. 1c). This pattern is similar to that documented for the CTCF ortholog of zebrafish19, clawed frog20, and mouse21 in that CTCF transcripts were widely distributed, particularly in the brain, branchial arches, and axial structure including the neural tube.

Timings of CTCF-CTCF2 and CTCF-CTCFL Duplications

The identification of the second lamprey homolog of CTCF, CTCF2, prompted us to analyze its evolutionary origin. Our phylogenetic analyses indicated that the duplication between CTCF and CTCF2 occurred in the lamprey lineage, after the split of the hagfish lineage (Fig. 1d). Also, the long terminal branches of CTCF2 indicated its elevated evolutionary rate. Prior to concluding that CTCF2 is a lamprey-specific paralog, we sought to carefully examine the possibility that CTCFL and CTCF2 were orthologs, by means of an in-depth analysis in which the likelihoods of all possible tree topologies are computed and statistically evaluated. This analysis did not support the exclusive phylogenetic clustering of CTCFL with CTCF2 (Table 1).

Table 1 Tree topology test for the phylogenetic relationship between CTCFL and CTCF2.

Previously, CTCFL was suggested to have duplicated in the lineage leading to amniotes, by an analysis employing nucleotide sequences without multiple substitutions taken into account22. To scrutinize the origin of CTCFL more closely, we used amino acid sequences from all major vertebrate lineages, including cyclostomes and chondrichthyans. Our molecular phylogenetic analysis suggested that the gene duplication giving rise to CTCFL occurred earlier in vertebrate evolution than previously suggested, that is, before the split of the chondrichthyan lineage (Fig. 1d and Table 2). We further examined if this early duplication timing was supported with variable tree inference methods and sequence datasets. Almost all of the phylogenetic trees inferred exhibited the duplication between CTCF and CTCFL before the split of the chondrichthyan lineage, while no trees supported the timing of this duplication in the amniote ancestor (Supplementary Fig. S3).

Table 2 Tree topology test for the duplication timing between CTCF and CTCFL

Immunodetection of Lamprey CTCF Protein

For ChIP-seq, we chose a commercially available anti-CTCF antibody that targets the C-terminus region (CST #3418S). Using this antibody, we performed western blotting against protein lysates of human GM12878 cells, a chicken embryo, and the adult liver and embryos of Arctic lamprey. This experiment identified positive bands of approximately 140 kDa for the human and chicken samples, while the band for lamprey was detected at over 250 kDa (Fig. 2a). The increased molecular weight of lamprey LjCTCF protein was consistent with its elongated amino acid sequence mentioned above (Fig. 1a).

Figure 2
Figure 2

Identification of CTCF proteins. (a) Western blotting of CTCF proteins, using an anti-CTCF antibody. Protein extracts from human GM12878 cells, stage 32 chicken embryo, adult lamprey liver, and stage 27 lamprey embryos were used for the analysis. β-actin (ACTB) or histone H3 was used as a loading control protein. (b) Immunoprecipitation. Silver-stained SDS PAGE gel of IP proteins showing chicken CTCF protein of approximately 140 kDa, and lamprey CTCF protein of >250 kDa. Detailed procedures of western blotting and immunoprecipitation are described in Supplementary Materials and Methods. Note that the band positions may not be accurate possibly because of posttranslational modification.

Immunoprecipitation was performed to examine whether the antibody specifically recognizes CTCF proteins. Silver staining after SDS-PAGE yielded clear single bands at around 140 kDa for chicken and over 250 kDa for lamprey (Fig. 2b). The gel bands were excised and analyzed with mass spectrometry, which resulted in identification of chicken CTCF and Arctic lamprey CTCF (Supplementary Table S3).

Genome-wide Detection of CTCF Binding Sites

Using the validated anti-CTCF antibody, ChIP assays were performed on Arctic lamprey, chicken, and human samples. Enrichment in the ChIP assays was confirmed by quantitative polymerase chain reaction (qPCR) (Supplementary Fig. S4). ChIP-seq yielded 16–23 million 80nt-long single reads per library, which were mapped to the genomes of the individual species (see Fig. 3a for lamprey and chicken). Proportions of mapped reads were 72–73% for lamprey, 90% for chicken, and 95% for human (Supplementary Table S4). The number of peaks called by MACS223 did not markedly differ between the lamprey liver and embryos (Fig. 3b), and more than 70% of the peaks were shared between the data derived from the liver and embryos (Fig. 3c). This pattern that CTCF binding landscape is intrinsic to individual genomes irrespective of cell types was previously suggested for human24, 25. Numbers of significant peaks were comparable among the analyzed species (Fig. 3b and Supplementary Table S5). Nucleotide sequences of core motifs and upstream motifs identified de novo by the MEME program26, were highly similar among the species (Fig. 3d) and identical to the motifs known for mammals3, 4, 24. The validity of the identified binding sites was confirmed with the observation that ChIP-seq peaks with higher fold enrichment more frequently harbor the CTCF binding motif sequence (Fig. 3e).

Figure 3
Figure 3

ChIP-seq peaks and binding motifs in lamprey, chicken and human cells. (a) ChIP-seq results for lamprey and chicken CTCF. Shown are the lamprey Hox α cluster and chicken Hox B cluster. (b) Numbers of peaks identified by MACS2. The “consensus peaks” (black bar) were identified by taking an intersect of peaks called in each replicate and in the merged replicates (see Materials and Methods for details about peak selection). (c) Overlap of consensus peaks in the lamprey embryo and adult liver tissue. (d) CTCF core and upstream motifs identified by MEME. The motifs were identified in two parts and are almost identical between the different species analyzed (also see Supplementary Fig. S11). (e) Peaks in various fold enrichment ranges harboring the core motif and core + upstream motifs. Numbers of ChIP-seq peaks are shown with white bars. Proportions of peaks containing a core motif (▪) and those containing a core + upstream motif () are indicated as lines.

Association of CTCF Peaks with Repeats

For mammals, it was shown that CTCF binding sites propagated through retrotransposition, and that the types of retrotransposons that contributed to this process differ even between mammalian taxa3, 8. We examined how CTCF binding sites propagated in the Arctic lamprey genome. To this end, a de novo repeat sequence library was constructed and used for identification of genomic locations of repetitive sequences (see Materials and Methods for details). We assessed possible overlap between ChIP-seq peaks and the repetitive sequences identified above with Fisher’s exact test. This revealed statistically significant positional association of LTR retrotransposons and DNA hAT transposons with CTCF ChIP-seq peaks (q < e-10, odds ratio > 10; Table 3). Particularly remarkable among these was the repetitive sequence #274 in the ‘hAT-Tip100’ repeat subclass, because as many as 80% of the repetitive sequences categorized into this group overlapped ChIP-seq peaks (Table 3). These peaks harbored canonical CTCF binding motifs at a high proportion, without CpG sites for potential DNA methylation, and tended to exhibit high fold enrichment in ChIP-seq, which was validated with qPCR (Supplementary Fig. S5). We queried the hAT-Tip100 sequence in search of counterparts in the genome of a closely related species, Petromyzon marinus, which did not yield any high-similarity hit—the most similar sequence showed an E-value of 0.25 for a 28 bp-long nucleotide sequence stretch. This suggests that the propagation of this repetitive element occurred after the split of these two lamprey species, estimated at 30–10 million years ago27.

Table 3 Association of CTCF binding with repetitive elements.

CTCF Binding in Hox Clusters

We analyzed distributions of repetitive sequences and CTCF binding sites in Hox gene clusters. First, as performed above for lamprey, repetitive sequences were identified in the whole genomes of chicken, mouse, and human (see Materials and Methods for details). This comparison, based on a uniform condition, revealed a higher extent of repeat intrusion into lamprey Hox gene clusters (37.6% versus 0.5–10.9% in the other species; Supplementary Table S6). The composition of repetitive sequences in the lamprey Hox clusters highly resembled that of its whole genome.

To perform a cross-species comparison of CTCF binding patterns, we focused on peaks with high fold enrichment (“significant peaks”; see Materials and Methods) and scanned them for binding motifs with their orientations (Fig. 4). The comparison, involving mouse, dog, and opossum for which CTCF ChIP-seq data was publicly available (Supplementary Figs S6 and S7), revealed a high similarity of CTCF binding pattern in the Hox clusters among the gnathostomes analyzed (Fig. 4). The gnathostome commonalities include 1) outward binding motifs located near Evx or Hox13 as well as near Hox5 and 2) shared positioning of binding motifs between the clusters (e.g. between Hox8 and -9 genes, and between Hox5 and -6 genes). In lamprey, we detected more CTCF binding sites (up to 18 sites in lamprey, compared to up to 10 in other species, per cluster), and the outward binding motifs, especially those in the 3′ regions, were located closer to the end of the clusters than in gnathostomes. More conspicuously, we identified multiple CTCF binding sites between Hox1 and -4, in four out of the five clusters covering this segment, whereas no or few peaks were detected in this segment in any jawed vertebrate species analyzed.

Figure 4
Figure 4

Distribution of CTCF binding sites in Hox clusters. Coding region of genes are indicated with gray boxes. CTCF binding sites are indicated with green arrowheads and bars. Arrowheads indicate the orientations of core motifs inferred by the FIMO program. Green bars indicate CTCF binding sites without a core motif. CTCF binding sites of the Arctic lamprey that overlap repeats are indicated with asterisks (see Results for details). Arrowheads in dashed boxes represent shared relative positions of CTCF binding sites between multiple Hox clusters. This figure includes only “significant peaks” defined in Materials and Methods. See Supplementary Fig. S6 for an equivalent scheme in mouse, dog, and opossum, and Supplementary Fig. S7 for detailed locations of all the peaks in Hox clusters. For the Arctic lamprey, we analyzed only Hox α-ε clusters that were identified in continuous sequences harboring multiple Hox genes.

In order to infer whether the vertebrate ancestor already possessed CTCF binding sites between Hox1 and -4 (see Results), we attempted to infer in silico the CTCF binding landscape in the Hox cluster of amphioxus, as well as the elephant shark, using the FIMO program, because ChIP-seq data on CTCF was not available for these species. The prediction referring to the motifs of the species analyzed with ChIP-seq data in this study, however, detected only a few CTCF binding sites inside the amphioxus and elephant shark Hox clusters. In fact, this negative result was not validated, as this method alone did not accurately predict CTCF binding sites in the human Hox clusters, either. The only species with CTCF ChIP-seq data that potentially serves as an outgroup is the fruitfly. In its Antennapedia complex (ANT-C), we identified outstanding ChIP-seq peaks between pb and Dfd genes, orthologous to vertebrate Hox2 and Hox4 genes, respectively (Supplementary Fig. S8).

Discussion

The present study encompassed the whole phylogeny of vertebrate CTCF and its relatives, now including the jawless vertebrate homologs. Out of the two lamprey CTCF genes identified in our gene model GRAS-LJ, LjCTCF is thought to be the canonical CTCF ortholog because it is expressed generally at higher levels and possesses the same number of Zn finger domains in its protein product as its gnathostome counterparts (Fig. 1). LjCTCF2 was revealed to be relatively lowly expressed and more rapidly-evolving, retaining fewer Zn finger domains (Fig. 1). Importantly, our in-depth phylogenetic analysis ruled out the possibility that lamprey CTCF2 is orthologous to CTCFL, and rather suggested the origin of CTCF2 in the lamprey lineage (Fig. 5). We also propose that CTCFL arose through gene duplication before the split of the chondrichthyan lineage (Fig. 1d and Supplementary Fig. S3). The hypothesis by Hore et al., postulating the origin of CTCFL early in amniote evolution22, was not statistically rejected (p ≥ 0.069; Table 2) and is preferred by maximum parsimony principle as it assumes fewer gene losses. However, Hore et al. inferred the phylogenetic tree employing a simple evolutionary model (p-distance based on nucleotide substitutions) and tree inference method (neighbor-joining method). We inferred phylogenetic trees using a similar sequence set as Hore et al. but employing more modern phylogenetic approaches, which did not suggest a recent duplication timing (Supplementary Fig. S3e,f,k and l). Tree inference using coding nucleotide sequences may cause phylogenetic mispositioning because the third codon positions lose phylogenetic signatures if substitutions are saturated. Indeed, our analysis using peptide sequences or nucleotides in the first and second codon positions favored a more ancient origin of CTCFL (Supplementary Fig. S3a–d and g–j).

Figure 5
Figure 5

Possible scenarios of the establishment of CTCF binding patterns in vertebrate Hox clusters. Alternative evolutionary scenarios of CTCF gene duplication and CTCF binding patterns in Hox clusters are depicted. CTCF-based Hox regulation spanning Hox1-4 genes was established either at the ancestor of bilaterians (Scenario A) or in the cyclostome lineage (Scenario B).

Our ChIP-seq experiments for lamprey CTCF were enabled by validation of antibody specificity across species and our optimized ChIP protocol. Our results on the lamprey revealed high similarities in the number and distribution of ChIP-seq peaks between different tissues or between different life stages (e.g., between liver and embryos; Fig. 3b and c), as previously shown for human24, 25. This observation validates the cross-species comparison of CTCF ChIP-seq peak distribution, even between different tissues and between embryonic stages. Our genome-wide comparison revealed a high similarity in numbers of CTCF binding sites, in spite of their variable genome sizes (1.2 Gbp for chicken, 1.6 Gbp for the Arctic lamprey, and 3.5 Gbp for human)15, as well as in the core and upstream binding motif sequences between Arctic lamprey, chicken, and mammals (Fig. 3b,c and d). We conclude that the establishment of CTCF binding property, known for mammals, occurred in the last common ancestor of extant vertebrates at the latest.

Distribution of ChIP-seq peaks in lamprey was shown to be associated with that of repetitive sequences, particularly retrotransposons and DNA transposons (Table 3; Supplementary Fig. S9). It was previously reported that CTCF binding sites are associated with retrotransposons in mammalian genomes3, 8. These suggest a similar process of CTCF binding site propagation within a genome, mediated by transposons, occurring independently in these different lineages.

A recent analysis suggested that some epigenomic features of mammalian Hox clusters, such as their global regulation involving distal enhancers, had not been established at the chordate ancestor28. In mice, the outward CTCF binding motif located between HoxA5 and HoxA6 functions as an anchor point for long range interaction12. Studies on cyclostomes should thus help pinpoint the timing of its establishment. Whole-genome sequencing on lampreys previously showed the elongation of Hox clusters14, 15, while their precise structures have not yet been determined. The present study provided a comparative landscape of repetitive sequences in Hox clusters (Supplementary Table S6), which revealed the contrast that lamprey Hox clusters are massively intruded by repetitive elements, whereas the Hox clusters of the gnathostomes analyzed are almost devoid of them. The increase in the number of CTCF binding sites inside the lamprey Hox clusters might be explained by transposon-based CTCF binding site propagation and maintenance (Fig. 4). Indeed, 72.7% (40 out of 55) of CTCF ChIP-seq peaks inside the Hox α-ε clusters were associated with repeat elements, which was similar to the proportion of their genome-wide association (72.3%; 19,981 out of 27,633 CTCF peaks) (Supplementary Fig. S9).

While the exact structures of lamprey Hox clusters remain to be determined, our CTCF ChIP-seq data has enabled the first vertebrate-wide comparison of CTCF binding patterns in Hox clusters. The shared positioning of the binding sites between different clusters might suggest that they originated before the split of the clusters. It showed that there is a common outward orientation of CTCF binding sites closest to the ends of Hox clusters, in both lamprey and gnathostomes (Fig. 4). Most striking was the contrast in the region between Hox1 and Hox4 intruded by CTCF binding sites in the lamprey, whereas no or few binding sites were identified in this segment in all the gnathostomes analyzed (Fig. 4 and Supplementary Fig. S6). The identification of CTCF motifs in the corresponding locations in the fruitfly Hox cluster (Supplementary Fig. S8) suggests an early origin of CTCF binding in the Hox1-4 region at the bilaterian ancestor, and that they subsequently decayed in the jawed vertebrate lineage (Scenario A in Fig. 5). On the other hand, Scenario B in Fig. 5 is supported by a notion that repeat-associated propagation of CTCF binding motifs might have occurred relatively recently, in light of the similar event in the mammalian lineage that occurred within these 100 million years at most.

Hox genes regulate the morphogenesis of various tissues in vertebrate embryos, and the origins of its molecular mechanisms have been intensively analyzed in lampreys, focusing on the hindbrain29, 30, pharyngeal arches31, and axial elements in the trunk32. It is of great interest to characterize possible influence of CTCF binding in the genomic region spanning Hox1 to Hox4 on morphological evolution marking the distinction between cyclostomes and the rest of the vertebrates, such as the acquisition of the articulating jaw. To address this point, it is crucial to characterize CTCF binding in chondrichthyan Hox clusters, which is currently ongoing.

Materials and Methods

Animals and Cells

Tissues of brain, eye, heart, intestine, liver, skeletal muscle, and pooled oocytes of an adult female, and the testis of an adult male Arctic lamprey, L. camtschaticum, from the Shiribetsu river, Hokkaido, Japan, were dissected, snap frozen in liquid nitrogen, and kept at −80 °C until use. L. camtschaticum embryos obtained by artificial fertilization were sampled as pools at stages 25 and 27, according to the embryonic staging of L. reissneri 33 and stored as above. Fertilized chicken eggs were obtained from a local farm. Chicken embryos used were at stage 25 according to the embryonic staging by Hamburger and Hamilton34, snap frozen, and kept at −80 °C until use. Human lymphoblastoid cell line GM12878, was purchased from the Coriell Cell Repositories and cultured in RPMI-1640 media (Gibco) supplemented with 15% FBS, 2 mM L-glutamine, and 1x antibiotic-antimycotic solution (Gibco), at 37 °C, 5% CO2. Animal care and experimental procedures were conducted in accordance with guidelines approved by the Institutional Animal Care and Use Committee (IACUC), RIKEN Kobe Branch.


RNA Extraction and RNA-seq Library Preparation

Tissues and embryos of L. camtschaticum were powderized while frozen, using SK-200 mill (Tokken) with stainless tubes and bullets. Tissue powders were quickly lysed in Trizol reagent (Thermo Fisher Scientific) to extract total RNAs. Libraries for RNA-seq were prepared using 1 μg total RNA following the standard protocol of TruSeq Stranded mRNA Sample Prep Kit (Illumina).


Whole-Mount In Situ Hybridization

Whole-mount in situ hybridization was performed following previous literature35. Riboprobes were designed for 5′ and 3′ regions of the LjCTCF and LjCTCF2 transcript sequences (Supplementary Fig. S10) and prepared using cDNA of embryos at stage 27 with oligonucleotide primers in Supplementary Table S7.


Molecular Phylogenetic Analysis

Deduced amino acid sequences of CTCF, CTCFL, and CTCF2 genes were aligned with the program MAFFT v7.22136 employing the L-INS-i method. From the entire multiple alignment, the stretch of Zn finger domains was extracted, and ambiguously aligned sites were removed with trimAl v1.437 with the ‘-automated1’ option followed by removal of gapped sites. Molecular phylogenetic trees with the maximum-likelihood (ML) framework were inferred using the program RAxML v8.2.438 assuming the PROTCATWAG model for amino acid sequences and the CATGTR model with assignment of distinct models to the codon positions for nucleotides. Credibility of the nodes in ML trees was evaluated with 1,000 bootstrap resamplings employing the rapid bootstrapping implemented in RAxML. Phylogenetic trees with the Bayesian framework were inferred with PhyloBayes v4.1b39 assuming the CAT-WAG-Γ model for amino acid sequences and the CAT-GTR-Γ model for nucleotide sequences. Tree-topology test was performed with a combination of CONSEL40 and RAxML assuming the PROTGAMMAWAG model. Accession numbers and sequences used for the analysis are listed in Supplementary Tables S8 and S9.


Chromatin Immunoprecipitation

Frozen liver tissue from an adult female lamprey, lamprey embryos at stage 27, and chicken embryos at stage 25, were powderized using a SK-200 mill and fixed in 1% formaldehyde/PBS (-) solution for 15 minutes at room temperature. Human GM12878 cells were fixed in 1% formaldehyde/PBS (-) solution for 5 minutes at room temperature. Chromatin lysates were prepared in lysis buffer (50 mM Tris-HCl pH8.0, 1% SDS, 10 mM EDTA, 1% proteinase inhibitor), by sonication with Covaris S220 or E220 (COVARIS).

ChIP was performed in ChIP dilution buffer (16.7 mM Tris-HCl (pH 8.0), 0.01% SDS, 1.1% (w/v) TritonX-100, 1.2 mM EDTA, 167 mM NaCl, 1% proteinase inhibitor) containing 4 mg/ml BSA (for lamprey and chicken embryonic samples) or 0.5 mg/ml BSA (for lamprey liver and human GM12878 cells) using lysates equivalent to 1 × 107 cells and protein A beads (Novex) coupled with 5 μl of anti-CTCF antibody (CST #3418 S). After 4 hours of IP reaction at 4 °C, beads were washed 4 times in a low salt buffer (20 mM Tris-HCl (pH 8.0), 0.1% SDS, 1% (w/v) TritonX-100, 2 mM EDTA, 150 mM NaCl), and two more times with a high salt buffer (20 mM Tris-HCl (pH 8.0), 0.1% SDS, 1% (w/v) TritonX-100, 2 mM EDTA, 500 mM NaCl). Chromatin complexes were eluted from the beads by agitation in elution buffer (10 mM Tris-HCl (pH 8.0), 300 mM NaCl, 5 mM EDTA, 1% SDS) and incubated overnight at 65 °C for reverse-crosslinking. Eluates were treated with RNase A and Proteinase K, and ChIP DNA was ethanol precipitated. Libraries were prepared using 20 ng of input DNA and 1 ng of ChIP DNA with KAPA LTP Library Preparation Kit (KAPA Biosystems) and custom synthesized TruSeq adaptors. The procedure of ChIP and library preparation is detailed in SI Materials and Methods.


Sequencing

All RNA-seq libraries and all but two ChIP-seq libraries were subjected to on-board cluster generation using HiSeq SR Rapid Cluster Kit v2 (Illumina) and sequenced on Rapid Run Mode of Illumina HiSeq 1500 (Illumina) to obtain single end 80 nt reads. Two ChIP-seq libraries (GM12878_input_rep1 and GM12878_ChIP_rep1) were subjected to cluster generation in cBot using TruSeq PE Cluster Kit v3 cBot-HS (Illumina) and sequenced in the high-output mode of HiSeq 1500 to obtain paired-end 101 nt reads. Only the forward reads from the paired-end data were used for analysis after trimming them to 80 nt, to adjust to those of the other samples. All the RNA-seq and ChIP-seq reads were processed by Trim Galore! v0.3.7 (http://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) at default parameters to trim reads at low quality sequences and Illumina TruSeq adaptor sequences. RNA-seq and ChIP-seq data are available at DDBJ Sequence Read Archive (DRA) under the accession DRA005605.


ChIP-seq Data Analysis

Arctic lamprey, chicken, and human ChIP-seq reads were mapped to the LetJap1.0, galGal5, and hg19 genome assemblies using Bowtie v0.12.841, respectively. For lamprey and chicken, uniquely mapped reads were obtained using ‘-m 1 -v 2 -a --best --strata’ options. Subsequently, unmapped reads were mapped using ‘-m 5 -n 2 -a --best --strata’ options, allowing each read to align with up to 5 locations, and combined with the uniquely mapped reads. For human, only uniquely mapped reads were obtained using ‘-m 1 -n 2 -a --best --strata’ options. Peak calling was performed using MACS2 v2.0.10 with default parameters, including a q-value cutoff of 0.01. Each of ChIP replicate 1, ChIP replicate 2, and a merger of these two replicates (“merged replicates”) was used as a treatment sample, while input replicate 1 was used as a control sample in all the three peak calling runs. Intersect peaks in replicate 1, replicate 2, and “merged replicates” identified by Bedtools v2.17.042, were annotated as “consensus peaks”. A subset of “consensus peaks” with their fold enrichment for “merged replicates” of ≥10 (for lamprey and chicken) and ≥20 (for human) was assigned as “significant peaks”. These enrichment cutoffs were determined referring to the averages of fold enrichment among “consensus peaks” of 16.2, 15.2, 13.3, and 29.1 for lamprey embryos, lamprey liver, chicken embryos, and human GM12878 cells, respectively (Supplementary Table S5).

Motif identification was performed using MEME v4.10.0. The top 2,000 peaks, ranked by fold enrichment, were used to identify enriched sequence motifs within ± 100 bp from the peak summit. Background 1st-order Markov model was made from the Top 2,000 peaks (summit ± 100 bp) with ‘-pseudo 0.01’ option. After identifying the 16 bp-long core CTCF motif by MEME, FIMO v4.10.143 was used with ‘--thresh 1e-4 and --motif-pseudo 0.01’ options to identify motif locations in all the peak regions (summit ± 100 bp). The core CTCF motif sequences identified by FIMO were extended for 20 bp to the 5′ direction to search for the previously described ‘upstream motif’ (also called ‘M2 motif’)3, 4. The analysis for ‘upstream motif’ was performed similarly to the core motif analysis, motif identified by MEME and motif located by FIMO with ‘--thresh 1e-3 and --motif-pseudo 0.01’ options.


Association of CTCF Binding to Repetitive Elements

RepeatModeler44 was ran on the genome assembly LetJap1.0 with default parameters, which resulted in 847 repeat elements including SINE (49 entries), LINE (202 entries), LTR (82 entries), and DNA transposon (145 entries). Positions of repeat elements were obtained by running RepeatMasker v4.0.545 against the LetJap1.0 genome using the custom repeat library obtained above.

Overlaps of CTCF binding sites with the repeat elements were analyzed with Bedtools targeting 41,421 consensus peaks of 100 bp (summit ± 50 bp) from the stage 27 embryo CTCF ChIP-seq data with fold enrichment of no less than 5. As a control to assess the significance of association with repeats, we arbitrarily selected from the LetJap1.0 genome assembly 50,000 non-overlapping genomic regions of 100 bp with no undetermined base (‘N’) inside. CTCF binding sites overlapping each of the 847 repeat elements were counted and tested for statistical significance by Fisher’s exact test. p-values were subjected to multiple correction by the Q-value package v1.34.046 implemented in R.


Comparison of CTCF Binding in Hox Clusters

Only “significant peaks” in CTCF ChIP-seq results were used in cross-species comparisons. The location and orientation of CTCF motifs within peak regions (summit ± 100 bp) were identified by FIMO with ‘--thresh 1e-3 and --motif-pseudo 0.01’ options. When multiple motifs were identified within a peak, only the motif with the lowest p-value was adopted for downstream analyses. The CTCF motif and the background file used here were those identified from the top 2,000 peaks for each species.


Data Availability

Nucleotide sequences of the L. camtschaticum LjCTCF and LjCTCF2 cDNAs were deposited in the NCBI GenBank database under the accession numbers KX830966 and KX830967, respectively. RNA-seq and ChIP-seq data were deposited in the DDBJ DRA under the accession DRA005605.

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References

  1. 1.

    Lobanenkov, V. V. et al. A novel sequence-specific DNA binding protein which interacts with three regularly spaced direct repeats of the CCCTC-motif in the 5′-flanking sequence of the chicken c-myc gene. Oncogene 5, 1743–53 (1990).

  2. 2.

    Ong, C. T. & Corces, V. G. CTCF: an architectural protein bridging genome topology and function. Nat. Rev. Genet. 15, 234–46 (2014).

  3. 3.

    Schmidt, D. et al. Waves of retrotransposon expansion remodel genome organization and CTCF binding in multiple mammalian lineages. Cell 148, 335–48 (2012).

  4. 4.

    Nakahashi, H. et al. A genome-wide map of CTCF multivalency redefines the CTCF code. Cell Rep 3, 1678–89 (2013).

  5. 5.

    Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–80 (2012).

  6. 6.

    Rao, S. S. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–80 (2014).

  7. 7.

    de Wit, E. et al. CTCF Binding Polarity Determines Chromatin Looping. Mol Cell 60, 676–84 (2015).

  8. 8.

    Schwalie, P. C. et al. Co-binding by YY1 identifies the transcriptionally active, highly conserved set of CTCF-bound regions in primate genomes. Genome Biol. 14, R148 (2013).

  9. 9.

    Heger, P., Marin, B., Bartkuhn, M., Schierenberg, E. & Wiehe, T. The chromatin insulator CTCF and the emergence of metazoan diversity. Proc. Natl. Acad. Sci. USA. 109, 17507–12 (2012).

  10. 10.

    Heger, P., Marin, B. & Schierenberg, E. Loss of the insulator protein CTCF during nematode evolution. BMC Mol. Biol. 10, 84 (2009).

  11. 11.

    Narendra, V. et al. CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation. Science 347, 1017–21 (2015).

  12. 12.

    Narendra, V., Bulajic, M., Dekker, J., Mazzoni, E. O. & Reinberg, D. CTCF-mediated topological boundaries during development foster appropriate gene regulation. Genes Dev. 30, 2657–2662 (2016).

  13. 13.

    Shimeld, S. M. & Donoghue, P. C. Evolutionary crossroads in developmental biology: cyclostomes (lamprey and hagfish). Development 139, 2091–9 (2012).

  14. 14.

    Smith, J. J. et al. Sequencing of the sea lamprey (Petromyzon marinus) genome provides insights into vertebrate evolution. Nat. Genet. 45, 415–21, 421e1-2 (2013).

  15. 15.

    Mehta, T. K. et al. Evidence for at least six Hox clusters in the Japanese lamprey (Lethenteron japonicum). Proc. Natl. Acad. Sci. USA. 110, 16044–9 (2013).

  16. 16.

    Manousaki, T. et al. Molecular evolution in the lamprey genomes and its relevance to the timing of whole genome duplications. in Jawless Fishes of the World, Vol. 1 (eds Orlov, A. & Beamish, R.) 2–16 (Cambridge Scholars Publishing, 2016).

  17. 17.

    Loukinov, D. I. et al. BORIS, a novel male germ-line-specific protein associated with epigenetic reprogramming events, shares the same 11-zinc-finger domain with CTCF, the insulator protein involved in reading imprinting marks in the soma. Proc. Natl. Acad. Sci. USA. 99, 6806–11 (2002).

  18. 18.

    Stanke, M., Steinkamp, R., Waack, S. & Morgenstern, B. AUGUSTUS: a web server for gene finding in eukaryotes. Nucleic Acids Res 32, W309–12 (2004).

  19. 19.

    Pugacheva, E. M. et al. Cloning and characterization of zebrafish CTCF: Developmental expression patterns, regulation of the promoter region, and evolutionary aspects of gene organization. Gene 375, 26–36 (2006).

  20. 20.

    Burke, L. J., Hollemann, T., Pieler, T. & Renkawitz, R. Molecular cloning and expression of the chromatin insulator protein CTCF in Xenopus laevis. Mech. Dev. 113, 95–8 (2002).

  21. 21.

    Delgado-Olguin, P. et al. CTCF promotes muscle differentiation by modulating the activity of myogenic regulatory factors. J. Biol. Chem. 286, 12483–94 (2011).

  22. 22.

    Hore, T. A., Deakin, J. E. & Marshall Graves, J. A. The evolution of epigenetic regulators CTCF and BORIS/CTCFL in amniotes. PLoS Genet. 4, e1000169 (2008).

  23. 23.

    Zhang, Y. et al. Model-based analysis of ChIP-Seq (MACS). Genome Biol. 9, R137 (2008).

  24. 24.

    Kim, T. H. et al. Analysis of the vertebrate insulator protein CTCF-binding sites in the human genome. Cell 128, 1231–45 (2007).

  25. 25.

    Cuddapah, S. et al. Global analysis of the insulator binding protein CTCF in chromatin barrier regions reveals demarcation of active and repressive domains. Genome Res. 19, 24–32 (2009).

  26. 26.

    Bailey, T. L., Johnson, J., Grant, C. E. & Noble, W. S. The MEME Suite. Nucleic Acids Res 43, W39–49 (2015).

  27. 27.

    Kuraku, S. & Kuratani, S. Time scale for cyclostome evolution inferred with a phylogenetic diagnosis of hagfish and lamprey cDNA sequences. Zool. Sci. 23, 1053–64 (2006).

  28. 28.

    Acemel, R. D. et al. A single three-dimensional chromatin compartment in amphioxus indicates a stepwise evolution of vertebrate Hox bimodal regulation. Nat. Genet. 48, 336–41 (2016).

  29. 29.

    Murakami, Y. et al. Segmental development of reticulospinal and branchiomotor neurons in lamprey: insights into the evolution of the vertebrate hindbrain. Development 131, 983–95 (2004).

  30. 30.

    Parker, H. J., Bronner, M. E. & Krumlauf, R. A Hox regulatory network of hindbrain segmentation is conserved to the base of vertebrates. Nature 514, 490–3 (2014).

  31. 31.

    Cerny, R. et al. Evidence for the prepattern/cooption model of vertebrate jaw evolution. Proc. Natl. Acad. Sci. USA. 107, 17262–7 (2010).

  32. 32.

    Takio, Y. et al. Hox gene expression patterns in Lethenteron japonicum embryos–insights into the evolution of the vertebrate Hox code. Dev. Biol. 308, 606–20 (2007).

  33. 33.

    Tahara, Y. Normal Stages of Development in the Lamprey, Lampetra-Reissneri (Dybowski). Zool. Sci. 5, 109–118 (1988).

  34. 34.

    Hamburger, V. & Hamilton, H. L. A Series of Normal Stages in the Development of the Chick-Embryo, (Reprinted from Journal of Morphology, Vol. 88, 1951). Dev. Dynam. 195, 231–272 (1992).

  35. 35.

    Sugahara, F., Murakami, Y. & Kuratani, S. Gene Expression Analysis of Lamprey Embryos. in In Situ Hybridization Methods (ed. Hauptmann, G.) 263–278 (Springer New York, New York, NY, 2015).

  36. 36.

    Katoh, K. & Standley, D. M. MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol. Biol. Evol. 30, 772–80 (2013).

  37. 37.

    Capella-Gutierrez, S., Silla-Martinez, J. M. & Gabaldon, T. trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics 25, 1972–3 (2009).

  38. 38.

    Stamatakis, A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30, 1312–3 (2014).

  39. 39.

    Lartillot, N., Lepage, T. & Blanquart, S. PhyloBayes 3: a Bayesian software package for phylogenetic reconstruction and molecular dating. Bioinformatics 25, 2286–8 (2009).

  40. 40.

    Shimodaira, H. & Hasegawa, M. CONSEL: for assessing the confidence of phylogenetic tree selection. Bioinformatics 17, 1246–7 (2001).

  41. 41.

    Langmead, B., Trapnell, C., Pop, M. & Salzberg, S. L. Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol. 10, R25 (2009).

  42. 42.

    Quinlan, A. R. & Hall, I. M. BEDTools: a flexible suite of utilities for comparing genomic features. Bioinformatics 26, 841–2 (2010).

  43. 43.

    Grant, C. E., Bailey, T. L. & Noble, W. S. FIMO: scanning for occurrences of a given motif. Bioinformatics 27, 1017–8 (2011).

  44. 44.

    Smit, A. F. A. & Hubley, R. RepeatModeler Open-1.0. [http://www.repeatmasker.org] (2008–2015).

  45. 45.

    Smit, A. F. A., Hubley, R. & Green, P. RepeatMasker Open-4.0. [http://www.repeatmasker.org] (2013–2015).

  46. 46.

    Storey, J. D. The positive false discovery rate: A Bayesian interpretation and the q-value. Ann. Stat 31, 2013–2035 (2003).

  47. 47.

    Shimodaira, H. An approximately unbiased test of phylogenetic tree selection. Syst. Biol. 51, 492–508 (2002).

  48. 48.

    Kishino, H. & Hasegawa, M. Evaluation of the maximum likelihood estimate of the evolutionary tree topologies from DNA sequence data, and the branching order in hominoidea. J. Mol. Evol. 29, 170–9 (1989).

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Acknowledgements

We thank Reiko Nakagawa in the Mass Spectrometry Facility in Kobe RIKEN for mass spectrometry, Kana Miyagishi for assistance in ChIP assays, Koh Onimaru, Sean D. Keeley, Kenta Sumiyama and Hisashi Miura for critical reading of the manuscript and discussion, Yukiko Nakaya and Guojun Shen for harvesting chicken embryos, Rie Kusakabe, Fumiaki Sugahara, Juan Pascual-Anaya and Shigeru Kuratani for lamprey embryos and inshore hagfish sequence data.

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Author notes

  1. Yuichiro Hara, Kaori Tanaka and Wataru Takagi contributed equally to this work.

Affiliations

  1. Phyloinformatics Unit, RIKEN Center for Life Science Technologies, 2-2-3 Minatojima-minami, Chuo-ku, Kobe, 650-0047, Japan

    • Mitsutaka Kadota
    • , Yuichiro Hara
    • , Kaori Tanaka
    • , Chiharu Tanegashima
    • , Osamu Nishimura
    •  & Shigehiro Kuraku
  2. Evolutionary Morphology Laboratory, RIKEN, 2-2-3 Minatojima-minami, Chuo-ku, Kobe, 650-0047, Japan

    • Wataru Takagi

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Contributions

M.K. and S.K. designed research. M.K., Y.H., K.T., W.T., O.N., and C.T. performed the experiments and sequence analysis. M.K., Y.H., K.T., W.T., and S.K. analyzed data. M.K., Y.H. and S.K. wrote the paper.

Competing Interests

The authors declare that they have no competing interests.

Corresponding author

Correspondence to Shigehiro Kuraku.

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https://doi.org/10.1038/s41598-017-04506-x

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