Primitive Duplicate Hox Clusters in the European Eel's Genome
Citation: Henkel CV, Burgerhout E, de Wijze DL, Dirks RP, Minegishi Y, et al. (
Primitive Duplicate Hox Clusters in the European Eel's Genome
Christiaan V. Henkel 0
Erik Burgerhout 0
Danie lle L. de Wijze 0
Ron P. Dirks 0
Yuki Minegishi 0
Hans J. Jansen 0
Herman P. Spaink 0
Sylvie Dufour 0
Finn-Arne Weltzien 0
Katsumi Tsukamoto 0
Guido E. E. J. M. van den Thillart 0
Michael Schubert, Ecole Normale Superieure de Lyon, France
0 1 ZF-screens B.V. , Leiden , The Netherlands , 2 Institute of Biology, Leiden University , Leiden , The Netherlands, 3 UMR BOREA , CNRS 7208, Muse um National d'Histoire Naturelle, Paris, France, 4 Norwegian School of Veterinary Science , Oslo , Norway , 5 Atmosphere and Ocean Research Institute, The University of Tokyo , Kashiwa, Chiba, Tokyo , Japan
The enigmatic life cycle and elongated body of the European eel (Anguilla anguilla L., 1758) have long motivated scientific enquiry. Recently, eel research has gained in urgency, as the population has dwindled to the point of critical endangerment. We have assembled a draft genome in order to facilitate advances in all provinces of eel biology. Here, we use the genome to investigate the eel's complement of the Hox developmental transcription factors. We show that unlike any other teleost fish, the eel retains fully populated, duplicate Hox clusters, which originated at the teleost-specific genome duplication. Using mRNA-sequencing and in situ hybridizations, we demonstrate that all copies are expressed in early embryos. Theories of vertebrate evolution predict that the retention of functional, duplicate Hox genes can give rise to additional developmental complexity, which is not immediately apparent in the adult. However, the key morphological innovation elsewhere in the eel's life history coincides with the evolutionary origin of its Hox repertoire.
Funding: This work was supported by the Norwegian School of Veterinary Science and the Research Council of Norway (184851), by Centre National de la
Recherche Scientifique and LAgence Nationale de la Recherche (08-BLAN-0173), and by private resources from ZF-screens B.V., Leiden University and The
University of Tokyo. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have read the journals policy and have the following conflicts: HPS and GEEJMvdT are founders and shareholders of
ZFscreens B.V. CVH, EB, RPD and HJJ are employees of ZF-screens B.V. This does not alter the authors adherence to all the PLoS ONE policies on sharing data and
The life history of the European eel (Anguilla anguilla L., 1758)
involves two distinct ocean-dwelling larval stages, a protracted
juvenile phase in European continental freshwater, and finally
sexual maturation coincident with migration to spawning grounds
in the Atlantic Ocean, presumably the Sargasso Sea (Figure 1) .
The complexity and geographical range of this life cycle have long
inspired evolutionary and physiological studies, especially on the
structure of the eels single, randomly mating (panmictic)
population , interspecific hybridization with the American eel
(A. rostrata, which shares the same oceanic spawning grounds ),
its hidden migrations , and the development of fertility .
Its catadromous migratory behaviour, long life, serious habitat
reduction, pollution, and overfishing may be amongst the causes of
the catastrophic collapse of the European eel population observed
over the past decades . So far, Anguilla species have resisted
efforts directed at efficient and sustainable artificial breeding .
As knowledge on the eels genetic makeup is sparse, physiological
studies aimed at understanding maturation, reproduction and the
sustenance of successive larval stages have not been able to take
full advantage of gene expression profiling. In order to alleviate
this shortcoming, we have sequenced and assembled its genome.
While the draft genome will be an important tool in
reproduction research, it also offers new perspectives for
fundamental studies in eel biology, as well as a resource for the
comparative interpretation of model fish genomes (e.g. zebrafish
and medaka). Here, we investigate its repertoire of Hox genes in a
comparative genomics context.
The Hox genes encode transcription factors, which throughout
the animal kingdom are involved in the developmental patterning of
the body plan. In vertebrates, Hox genes are tightly organized into
clusters which exhibit colinearity between gene position and
temporal and spatial expression along the primary body axis: genes
at the 39 ends of clusters are expressed earlier in development, and
more anterior, than genes at the 59 ends of clusters . The
organization of Hox clusters has been extensively documented for
many groups of vertebrates .
A. anguilla is a member of the superorder Elopomorpha [11,12], a
major teleost group of 856 species . As such, elopomorphs
presumably share the inferred whole-genome duplication at the base
of the teleost lineage [14,15]. This teleost-specific genome duplication
(TSGD) event is most apparent when considering the Hox genes in
extant species [10,16,17]. In tetrapods and coelacanths,
approximately forty genes are organized in four ancestral vertebrate clusters.
In theory, teleosts could have retained eight duplicate clusters.
However, whereas tetrapod Hox loci are relatively stable, teleost
genomes show dramatic gene loss, such that all species examined in
detail retain at most seven of these clusters, each with on average
about half their original gene content [9,10]. A PCR-based survey of
Figure 1. The life cycle of the European eel. After hatching, presumably in the Sargasso Sea, cylindrical larvae develop into leaf-shaped
leptocephalus larvae, which after drifting on the Gulf Stream for approximately one year metamorphose into glass eels close to the European coast.
The glass eels may stay at the coast or migrate upriver, where they stay as juveniles (elvers and yellow eel) for many years (depending on the region:
males 46 years, females 812 years). Finally, they develop into migrating silver eels; the cause and timing of silvering is not well understood. They
mature during or after migration to the spawning grounds.
the Hox clusters of the Japanese eel A. japonica found evidence for the
conservation of eight clusters and 34 genes .
As the Elopomorpha represent an early branch on the teleost
tree , the eel Hox gene complement may expose constraints on
the evolution of morphological complexity in teleost fish, and in
vertebrates in general. Furthermore, analysis of the eels Hox
clusters may shed light on the developmental mechanisms and
evolutionary history of its life cycle and body plan. In particular,
they may provide evidence regarding the evolutionary novelty of
the eels indirect development.
Genome assembly of the European eel
We have sequenced and assembled the genome of a female
juvenile A. anguilla specimen caught in the brackish Lake Veere,
the Netherlands in December 2009. Its haploid genome size was
determined to be 1.1 Gbp. Because of the impossibility of breeding
A. anguilla, no genetic linkage information is available. We
therefore employed Illumina Genome Analyzer sequencing
technology only in the assembly of a draft genome. Based on a
de novo genome assembly, we constructed 923 Mbp of scaffolds
with a length-weighed median fragment length (N50) of 78 Kbp
(Figure S1 and Table S1). An additional 179 Mbp of initial
contigs, which are either very small or highly repetitive, were
excluded from scaffolding, but included in all further analyses.
Identification of Hox transcripts and genes
To identify A. anguilla Hox genes, we used a de novo assembled
transcriptome of a 27 hours post-fertilization (hpf) embryo of the
short-finned eel, A. australis. This species is closely related to A.
anguilla , yet produces viable embryos more easily . We
compared Hox-like sequences from the transcriptome to the
genome assembly using Blast , which yielded ten genomic
scaffolds (Table S2) that were further examined for the presence of
additional genes. This resulted in the identification of 73 Hox genes
(twice as many as found in A. japonica in a previous study using PCR
fragments ), including three pseudogenes, organized in eight
clusters (Figure 2 and Table S3). The flanking regions of these eight
clusters contain an additional 24 predicted genes (Figure 2). No
further protein-coding genes were found within the Hox clusters.
Conserved microRNAs were discovered using Blast searches
with human and zebrafish homologues (Figure 2). miR-10 is
present posterior of Hox4 in six clusters (all except Aa and Ab).
miR-196 is found between Hox9 and Hox10 in five clusters (all
except Bb, Da and Db). This arrangement is consistent with that
found in other vertebrates [22,23].
Hox cluster identity
We based preliminary identification of clusters on homology
between A. anguilla and Danio rerio protein sequences and
comparisons with all sequences in the NCBI non-redundant
protein database (Table S3). Whereas the two A. anguilla HoxA
clusters can easily be matched to their corresponding HoxAa and
HoxAb orthologues in D. rerio, each of the two HoxB and HoxC
clusters of A. anguilla most closely resembles D. rerio HoxBa and
HoxCa, respectively. Both A. anguilla HoxD clusters predictably
match D. rerio HoxDa only, since the zebrafish HoxDb cluster has
lost all protein-coding genes .
To more precisely assign the Hox genes to proper cluster
orthologues, we generated unrooted maximum likelihood
phylogenetic trees for paralogous group 9 (Figures 3 and S2), of which A.
Figure 2. Genomic organization of the Hox gene clusters of the European eel. Scaffolds are indicated by black lines and asterisks represent
two gaps between scaffolds. Hox genes are indicated by colored arrows that are numbered according to their paralogous groups. Three
pseudogenes are indicated by the symbol y. Neighboring genes are indicated by grey arrows. Conserved microRNA genes are indicated by triangles:
miR-196 (closed triangles) and miR-10 (open triangles).
anguilla possesses all eight copies. These confirmed the preliminary
classification in A, B, C and D paralogous groups, but failed to
validate the identity of teleost a and b cluster duplicates (with the
exception of HoxAa and HoxAb). Likewise, phylogenetic trees
based on multi-gene alignments do not conclusively indicate either
a or b cluster membership for HoxB, HoxC and HoxD (Figure 4).
In general, there appears to be a lack of sequence divergence
between eel Hox gene duplicates, which makes classification based
on coding sequence alone inaccurate.
Final orthologous relationships could only be established on the
basis of conserved local synteny between Hox clusters and flanking
genes (Figure 5). In addition to both HoxA clusters, eel HoxBa and
HoxBb appear orthologous with their respective teleost
equivalents. This identification is further supported by the absence of
miR-196 from both D. rerio and A. anguilla HoxBb clusters. The
affinities of HoxC and HoxD duplicates remain difficult to resolve
because of conserved synteny around a and b cluster duplicates,
and extensive cluster reduction and deletion in other teleosts
(Figure 5c, d).
Hox gene expression
In order to confirm the transcriptional activity of the Hox genes,
we determined relative expression levels by aligning transcriptomic
reads of the 27 hpf embryo against the Hox protein-coding regions
(Figure 6a). Transcriptome reads mapped unambiguously to 71
out of 73 Hox genes, including one pseudogene (yHoxD3b),
suggesting that all A. anguilla Hox protein-coding genes are
functional. The relative expression levels vary over five orders of
magnitude with the lowest expression observed for the posterior
paralogous groups 12 and 13, and the highest expression for
paralogous groups 79, but with particularly high expression levels
for HoxB1a, HoxB1b, HoxB4b and HoxD1a.
Full mRNA-seq read alignment to the entire Hox clusters
indicated that transcriptional activity is not restricted to protein
coding regions (Figure S3). In fact, intergenic expression
sometimes exceeds intragenic levels, supporting the observation
that complete Hox clusters function as meta-genes [9,25].
At 27 hpf, expression of posterior Hox genes is very low
(Figure 6a). We therefore confirmed transcriptional activity of
posterior Hox paralogues by whole mount in situ hybridizations
(Figure 4b). HoxB9a is expressed at 26 and 48 hpf, with an anterior
expression boundary coinciding with somite number 5/6.
Expression of HoxD12b and HoxC13a is not yet detectable at 48
hpf, but clearly visible at 96 hpf with anterior expression
boundaries located at somite numbers 65/70 and 90/95 for
HoxD12b and HoxC13a, respectively. For these Hox genes,
expression in the eel embryo appears to conform to the expected
spatio-temporal pattern (colinearity between cluster organization
and developmental timing and localization), with expression of
Hox12 and Hox13 paralogues appearing later in development,
and more posterior than Hox9.
The evolution of Hox cluster organization
The early branching of the Elopomorpha from the main teleost
trunk allows a new reconstruction of ancestral Hox cluster
architectures (Figure 7), which are strongly constrained by the limited
organizational divergence between eel HoxB, C and D duplicates.
Since teleost fish are believed to have experienced the TSGD
event early in their evolutionary history [14,15], their genomes
should in theory possess up to eight cluster duplicates. However,
all teleosts examined in detail retain at most seven clusters of
protein-coding genes : a HoxC duplicate was lost in the lineage
leading to medaka and pufferfish, a HoxD duplicate in the lineage
represented by zebrafish. The high number of clusters in salmon is
the result of relatively recent further duplications .
The main teleost lineages diverged briefly after the TSGD .
The reconstruction in Figure 7 demonstrates that nearly all
postduplication gene loss events in the eels ancestry occurred within
this interval, followed by millions of years of stasis. Only the
HoxAb cluster appears to have accumulated major changes in
prebranching, post-genome duplication teleosts. Alternative
hypotheses, in which a whole-genome duplication is not shared between
elopomorphs and advanced teleosts, or in which the genome
duplication is followed by successive deletion and duplication of
specific clusters in the eel, are less parsimonious and not consistent
with local conservation of synteny (Figure 5).
Two rounds of Hox cluster duplications in chordates are
believed to be responsible for important vertebrate novelties (e.g.
brains, heads, jaws) and increases in complexity . A plausible
mechanism invokes a temporary relaxation of meta-genic cluster
constraints after duplication, paving the way for innovation
Figure 4. Phylogeny of Hox clusters of the European eel. Unrooted phylogenetic trees based on alignments combining multiple Hox genes
per cluster. A) Cluster A relationships, based on HoxA9, HoxA11 and HoxA13 genes. B) Cluster B relationships, based on HoxB1, HoxB5 and HoxB6
genes. C) Cluster C relationships, based on HoxC6, HoxC11, HoxC12 and HoxC13 genes. D) Cluster D relationships, based on HoxD4 and HoxD9 genes.
Species included: A. anguilla, Salmo salar (Atlantic salmon), Danio rerio (zebrafish), Oryzias latipes (medaka), and Tetraodon nigroviridis (green spotted
puffer). Asterisks indicate bootstrap support .90%.
Figure 5. Synteny around Hox clusters. Conservation of flanking genes supports the classification of A. anguilla clusters into different
orthologous subgroups. The eel clusters and up to seven flanking genes are compared with the genomic organization in zebrafish (Danio rerio) and
medaka (Oryzias latipes). Coloured box outlines indicate preserved synteny between eel and the two other species, dotted outlines denote flanking
genes found on extended eel scaffolds (see Methods). Interpretation should take into account residual synteny between a and b paralogous clusters.
Limited data is available on HoxCb (lost in O. latipes, possibly misassembled in D. rerio) and HoxDb (lost in D. rerio) clusters.
[28,29]. In contrast, the TSGD-associated third duplication of
vertebrate Hox clusters theoretically endowed teleost fish not with
additional complexity within individuals, but with increased
prospects for morphological diversification between individuals
and species [9,10]. In support of this hypothesis, advanced teleosts
have extensively pruned their Hox surplus, leading to significant
diversity in cluster structure (Figure 7). In all examined
representatives (with the exception of salmon ), the residual
number of Hox genes is not much higher than the non-duplicated
count in tetrapods. The resulting teleost Hox cluster architectures
have been interpreted as an evolutionary choice for developmental
flexibility in a trade-off with robustness . By proving that it is
possible for a vertebrate to stably preserve eight densely populated
(Figure 2) and functional (Figure 6) Hox clusters, the eel genome
presents an exception to these models, and a third alternative in
the evolution of vertebrate complexity.
For hundreds of millions of years, A. anguilla and its ancestors have
maintained the highest ontogenic potential of any vertebrate,
indicative of continuous selective pressure. However, as adults, they
do not display markedly more complex bodies than other fish or
tetrapods. The eels distinctive life cycle and body plan suggest three
(not mutually exclusive) explanations for this cryptic complexity.
Hox genes are involved in the primary patterning of the body
axis, which implies a functional role for A. anguillas Hox surplus in
axial elongation. Alterations in Hox genes have been associated
with elongated body plans [30,31], however the changes observed
are of a regulatory nature, and do not involve extra genes. For
example, elongation of the body axis in snakes has been linked to a
spatial relaxation in the posterior end of Hox clusters facilitated by
the insertion of transposable elements between genes . In
addition, even the elongate members of the Elopomorpha (which
also includes non-elongated tarpons, bonefish and others) display
considerable diversity in the developmental mechanisms resulting
in axial lengthening . Hence, the eels adult body plan cannot
explain the preservation of primitive Hox clusters between the
TSGD (226316 million years ago ) and the origin of the
genus Anguilla, estimated at 2050 million years ago .
Similarly, if the European eel may at present experience singular
evolutionary forces because of its panmictic population , any
explanation these offer does not extend beyond the genus Anguilla
of freshwater eels .
Even if for most of their lives eels are eel-shaped, as
oceandwelling larvae  their body plan is radically different (Figure 1).
In fact, until the late nineteenth century, these large, long-lived,
laterally compressed leptocephali were considered to be
autonomous pelagic species . Fully transparent and slowly
metabolizing, a leptocephalus provides considerable survival benefits
[37,38]. After approximately one year, they undergo a dramatic
metamorphosis , including extensive tissue remodelling and
shortening of the body, resulting in cylindrical juveniles. In the
early embryos investigated here (Figure 6), nearly all Hox genes
are expressed and presumably functionally involved in
determining cell fate. Logically, a high gene and cluster count can be
explained by the assumption that the eels two body plans are
simultaneously outlined at this stage.
Leptocephali are the fundamental developmental innovation
shared by all Elopomorpha , and therefore arose either
before or soon after the TSGD, or at the base of the lineage (arrows
in Figure 7). The last alternative is the most parsimonious (no loss of
developmental complexity in advanced teleosts), especially since no
member of the Elopomorpha is known to have ever discarded the
leptocephalous larval stage [11,13]. Regardless, either of the
postTSGD origins is compatible with an intercalation model of indirect
development , in which a temporary excess of developmental
potential was permanently recruited for the conception of an
additional body plan. Although speculative, an explanation
invoking the morphological challenges associated with a complex
life history is consistent with the stable high Hox gene and cluster
count found in the anadromous Atlantic salmon .
Further functional studies on eel development will become
possible once A. anguillas life cycle can be completed in captivity. In
particular, there exists considerable variation in development
(timing, number of somites) between leptocephali of related and
interbreeding Anguilla species [1,35], which can only be studied
when these larvae can be raised under controlled conditions [8,41].
Wild female and male silver short-finned silver eels (A. australis)
from Lake Ellesmere, New Zealand, were held together in a 2,300 L
recirculation system with seawater (30 ppt salinity) at 21uC. Sexual
maturation was induced as described . Briefly, males received
nine weekly injections with 250 IU human chorionic gonadotropin
and females were injected once a week with 20 mg salmon pituitary
extract. Eggs and milt were stripped and the eggs were dry fertilized.
Embryos were reared in glass beakers with UV-sterilized seawater
(35 ppt) at 21uC. At 26, 48 and 96 hpf embryos were fixed in 4%
paraformaldehyde and stored in 100% methanol.
Total RNA was isolated from 27 hpf embryos using the Qiagen
miRNeasy kit according to the manufacturers instructions (Qiagen
GmbH, Hilden, Germany), and analyzed with an Agilent
Bioanalyzer 2100 total RNA Nano series II chip (Agilent, Santa Clara). A
transcriptome library was prepared from 10 mg total RNA, using the
Illumina mRNA-Seq Sample Preparation Kit according to the
manufacturers instructions (Illumina Inc., San Diego, USA).
Genome size determination
Blood samples taken from two eels (A. anguilla and A. australis)
were washed with physiological salt and fixed in cold ethanol.
Prior to analysis the cells were collected, resuspended in
physiological salt and stained with propidium iodide. After
30 minutes of incubation the cells were analyzed by FACS, using
human blood cells as a size reference (3.05 Gbp haploid). The eel
genome size was calculated by (human size)/(mean fluorescence
human)6(mean fluorescence eel). Both Anguilla genomes were
determined to be 1.1 Gbp in size (haploid).
Genomic DNA libraries
Genomic DNA was isolated from blood of a female yellow
European eel (A. anguilla, caught in Lake Veere, The Netherlands)
using the Qiagen Blood and Tissue DNeasy kit according to the
manufacturers description. Paired-end libraries were prepared
from 5 mg of isolated gDNA using the Paired-End Sequencing
Sample Prep kit according to the manufacturers description.
Either a 200 bp band or a 600 bp band was cut from the gel
(libraries PE200 and PE600, Table S1). After amplification for 10
cycles the resulting libraries were analyzed with a Bioanalyzer
2100 DNA 1000 series II chip.
Mate pair libraries were prepared from 10 mg of isolated gDNA
using the Mate Pair Library Prep Kit v2 (Illumina Inc.). Either a
3,000 bp band or a 10,000 bp band was cut from gel (libraries
MP3K and MP10K, Table S1). After the first gel purification the
fragment length was analyzed using a Agilent Bioanalyzer 2100
DNA 12000 chip. After circularization, shearing, isolation of
biotinylated fragments, and amplification, the 400 to 600 bp
fraction of the resulting fragments was isolated from gel. Finally,
the libraries were examined with an Agilent Bioanalyzer 2100
DNA 1000 series II chip.
All libraries were sequenced using an Illumina GAIIx
instrument according to the manufacturers description. Genomic
paired-end libraries were sequenced with a read length of 2676
nucleotides (to ,20-fold genome coverage), genomic mate-pair
libraries with a read length of 2651 nucleotides (to ,33-fold
genome span), and the mRNA-Seq library with a read length of
2676 nucleotides (Table S1). Image analysis and base calling was
done by the Illumina pipeline.
Sequencing reads from both paired-end libraries were used in
building the initial contigs (Figure S1). Both sets were preprocessed
to eliminate low quality and adapter contamination. Whenever
possible, PE200 pairs were merged into longer single reads. For
initial contig assembly, we employed the De Bruijn graph-based de
novo assembler implemented in the CLC bio Genomics
Workbench version 3.6.5 (CLC bio, Aarhus, Denmark). A run with a
kmer length of 25 nt resulted in an assembly a total length of
969 Mbp and a contig N50 of 1672 bp.
Initial contigs were oriented in larger supercontigs (scaffolds)
using SSPACE . In scaffolding the contigs, we decided to
exclude low-quality and highly repetitive contigs as much as
possible. SSPACE was used in a hierarchical fashion, employing
first links obtained from the PE600 library to generate
intermediate supercontigs, which were used as input for subsequent runs
with links from the MP3K and MP10K libraries, respectively. At
each stage, a minimum of three non-redundant links was required
to join two contigs. This procedure resulted in a final scaffold set
with a total length of 923 Mbp and an N50 of 77.8 Kbp (Table
S1). AUGUSTUS (version 2.4) was used to predict genes ,
which were provisionally annotated using Blast2GO (version 2.4.8)
. The draft assembly is available at www.eelgenome.org.
In order to obtain more information on flanking genes for the
analysis of conserved synteny (Figure 5), scaffolds were subjected to
a further round of linking by SSPACE using reduced stringency
(two instead of three non-redundant links required to join scaffolds).
This resulted in extended scaffolds with an N50 of 169 Kbp.
Hox contigs in the short-finned eel embryonic transcriptome
(generated using CLC bios de novo assembler) were identified via
Blast  searches at the NCBI website (www.ncbi.nlm.nih.gov).
European eel genomic scaffolds were annotated using CLC bios
DNA Workbench. Remaining Hox genes and genes flanking the
Hox clusters were identified using Blast, based on AUGUSTUS/
Blast2GO predictions. Annotated Hox scaffolds have been
submitted to GenBank (accession numbers JF891391JF891400).
MicroRNAs were identified by Blast using H. sapiens and D. rerio
miR-10 and miR-196 sequences (precursors and mature) retrieved
from miRBase release 18 (www.mirbase.org, ).
Species and Hox gene accession numbers used are listed in
Table S4. Amino acid sequences of Hox genes were aligned using
Clustal X  and checked manually. After excluding ambiguous
alignments, ProtTest 2.4  was used to choose an optimum
substitution model, based on the Akaike information criterion. The
aligned sequences were subjected to maximum likelihood analysis
using RAxML version 7.2.6  with 1000 rapid bootstrap
replicates (-f a option).
For the analysis of Hox9 genes (Figure 3), 70 aligned residues
were used and analyzed using a JTT+I+C model . All other
alignments were fitted using a JTT+C model. The multi-gene
analyses of HoxA, HoxB, HoxC and HoxD (Figure 4) were based
on alignments of 427, 493, 935 and 308 amino acid residues,
respectively. The phylogenetic trees of sarcopterygian and
actinopterygian Hox9 paralogues (Figure S2) were based on 151
(HoxA9), 210 (HoxB9), 248 (HoxC9), and 136 (HoxD9) residues.
Synteny was analyzed using D. rerio and O. latipes genomic
contexts extracted from Ensembl release 65 (www.ensembl.org),
based on the Zv9 and MEDAKA1 genome assemblies,
respectively (Table S5). Pairwise alignments were generated by NCBI
tblastx and analyzed using genoPlotR .
Whole mount in situ hybridization
Chromosomal DNA was isolated from A. australis blood using a
DNeasy Blood & Tissue Kit (Qiagen). Riboprobe template
fragments, including a T7 RNA polymerase promoter, were PCR
amplified from chromosomal DNA using the following primer sets:
HoxB9a forward (59-TGAAACCGAAGACCCGAC-39), HoxB9a
(59-GAAATTAATACGACTCACTATAGGGCTGAGGAAGACTCCAA), HoxD12b forward
(59-TAATCTTCTCAGTCCTGGCTATG-39), HoxD12b reverse
HoxC13a forward (59-CACCTTGATGTACGTGTATGAAAA-39),
riboprobes were made according to standard protocols using T7 RNA
polymerase. Whole mount in situ hybridization with labelled
riboprobes was performed at 70uC, according to a slightly modified
version of a standard protocol . Hybridizing riboprobes were
made visible using anti-Digoxigenin AP and BM Purple AP
substrate (Roche). Stained embryos were bleached using hydrogen
peroxide (Sigma-Aldrich) and photographed using a Leica M205
FA stereo microscope.
Genome assembly pipeline. See Methods section
Figure S2 Unrooted maximum likelihood phylogenetic
trees of actinopterygian and sarcopterygian Hox9 genes.
See Methods section for details. Sequences used are listed in Table S4.
Figure S3 Meta-genic expression of Hox clusters.
mRNA-seq reads of the A. australis embryo were aligned to entire
Hox-containing scaffolds, demonstrating large amounts of mRNA
production from intronic and intergenic regions.
Table S5 Hox clusters used in synteny analysis. Genomic
locations of D. rerio and O. latipes Hox clusters. HoxCb is absent
from O. latipes, and HoxDb from D. rerio. However, the genomic
loci can still be identified based on the presence of flanking gene
duplicates or conserved microRNA (D. rerio HoxDb).
We thank the following colleagues for generously contributing to our work:
Bas Brittijn for production of short-finned eel embryos, Nabila Bardine and
Tony Durston for help with in situ hybridizations, Tony Durston and Joost
Woltering for critical reading of the manuscript, Pieter Slijkerman for
project financial management, Marten Boetzer and Walter Pirovano for
help with genome assembly.
Conceived and designed the experiments: CVH EB RPD HPS SD F-AW
KT GEEJMvdT. Performed the experiments: EB DLdW HJJ. Analyzed
the data: CVH RPD YM. Wrote the paper: CVH RPD.
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