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. 2022 Feb 21;12(4):jkac045. doi: 10.1093/g3journal/jkac045

Genome of the Rio Pearlfish (Nematolebias whitei), a bi-annual killifish model for Eco-Evo-Devo in extreme environments

Andrew W Thompson 1,2,✉, Harrison Wojtas 1, Myles Davoll 1,3, Ingo Braasch 1,2,✉
Editor: A Whitehead
PMCID: PMC8982402  PMID: 35188191

Abstract

The Rio Pearlfish, Nematolebias whitei, is a bi-annual killifish species inhabiting seasonal pools in the Rio de Janeiro region of Brazil that dry twice per year. Embryos enter dormant diapause stages in the soil, waiting for the inundation of the habitat which triggers hatching and commencement of a new life cycle. Rio Pearlfish represents a convergent, independent origin of annualism from other emerging killifish model species. While some transcriptomic datasets are available for Rio Pearlfish, thus far, a sequenced genome has been unavailable. Here, we present a high quality, 1.2 Gb chromosome-level genome assembly, genome annotations, and a comparative genomic investigation of the Rio Pearlfish as representative of a vertebrate clade that evolved environmentally cued hatching. We show conservation of 3D genome structure across teleost fish evolution, developmental stages, tissues, and cell types. Our analysis of mobile DNA shows that Rio Pearlfish, like other annual killifishes, possesses an expanded transposable element profile with implications for rapid aging and adaptation to harsh conditions. We use the Rio Pearlfish genome to identify its hatching enzyme gene repertoire and the location of the hatching gland, a key first step in understanding the developmental genetic control of hatching. The Rio Pearlfish genome expands the comparative genomic toolkit available to study convergent origins of seasonal life histories, diapause, and rapid aging phenotypes. We present the first set of genomic resources for this emerging model organism, critical for future functional genetic, and multiomic explorations of “Eco-Evo-Devo” phenotypes of resilience and adaptation to extreme environments.

Keywords: Nematolebias whitei, Rio Pearlfish, diapause, aging, hatching, extreme environments, Eco-Evo-Devo, teleost

Introduction

Aplocheiloid killifishes inhabit tropical freshwater habitats around the world. Some African and Neotropical species live in ephemeral waters that are subject to seasonal desiccation (Myers 1952; Simpson 1979). Desiccation kills the adults, but embryos survive inside specialized eggs (Thompson et al. 2017a) buried in the soil via 3 diapause stages (DI, DII, DIII; Wourms 1972a,1972b, 1972c). DI occurs as a migratory dispersion of blastomeres, DII occurs during somitogenesis when organs are rudimentary, and DIII occurs after organogenesis when the embryo is fully formed and poised to hatch upon habitat inundation. This seasonal life history is a remarkable example of convergent evolution (Furness et al. 2015) with 7 gains across killifish evolution (Thompson et al. 2021a). Additionally, annual killifishes show rapid aging due to relaxed selection on lifespan (Cui et al. 2019) and are an important emerging model system for the study of senescence (Valenzano et al. 2011, 2015; Harel et al. 2015; Reichwald et al. 2015).

The Rio Pearlfish, Nematolebias whitei, is a seasonal killifish endemic to the coastal plains of the Rio de Janeiro region in Brazil, inhabiting pools that dry twice annually, from July–August and February–March (Fig. 1a;Myers 1942; Costa 2002). Pearlfish represents a separate origin of seasonality from other killifish model species, i.e. Nothobranchius furzeri and Austrofundulus limnaeus (Thompson et al. 2021a). In N. whitei, DI and DII are facultative, and DIII is a “prolonged,” “deep” stasis compared to hatching delay and DIII in other killifishes, occurring just before environmentally cued hatching upon submersion in water (Wourms 1972c; Thompson and Ortí 2016; Thompson et al. 2017b). Pearlfish was suggested as a top candidate species for killifish models in the seminal work of developmental biologist John P. Wourms in 1967. They are small, prolific, and hardy, and spawn in sand (Wourms 1967), making them easily reared laboratory animals that are furthermore amenable to genetic manipulation like other killifishes (Aluru et al. 2015; Harel et al. 2015). Pearlfish has also been an emergent system to study aging (Ruijter 1987), environmental influences on development (Ruijter et al. 1984), the role of prolactin in hatching control (Schoots et al. 1983; Ruijter and Creuwels 1988), resilience to perturbations in development with the ability to develop normally from diblastomeric eggs (Carter and Wourms 1993), and the transcriptional control of diapause and hatching (Thompson and Ortí 2016).

Fig. 1.

Fig. 1.

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Rio Pearlfish evolution, ecology, development, and 3D genome structure. a) Bi-annual life cycle of the Rio Pearlfish with three developmental diapause stages following burying of eggs in soil. b) Relative position of Rio Pearlfish in the vertebrate tree of life inferred by Orthofinder based on annotated proteins. c) DNA barcode (cox1 and cytb) phylogeny inferred with RAxML of the genus Nematolebias confirming the identity of the genome specimen as N. whitei. Sequences from Costa et al. (2014) were used for comparison to the reference genome sequence. Green nodes show 100% bootstrap support for the reciprocal monophyly of N. whitei with other genera and confirms the identity of the genome specimen with high confidence. d) Hi-C contact map of the Rio Pearlfish genome showing linkage of the 24 chromosomes into chromosomal pseudomolecules. e, f) SynMap genome-wide synteny plots of Rio Pearlfish vs. medaka (e) and vs. zebrafish (f) showing genome-structure conservation across over 250 million years of teleost evolution. g) Hi-C contact maps of the syntenic region between rasa1a and mctp1a in Pearlfish liver tissue. These contact maps highlight the conserved 3D structure that include TADs conserved across teleost evolution as well as cell types and developmental stages (Nakamura et al. 2021). Species graphics Created with BioRender.com.

Here, we construct a chromosome-level genome assembly for the Rio Pearlfish, utilizing Hi-C contact maps, genome annotations, and gene expression analyses to characterize genomic evolution and hatching biology in this extremophile vertebrate.

Materials and methods

Genome sequencing and assembly

All animal work was approved by the Michigan State University Institutional Animal Care and Use Committee (PROTO202000108).

A total of 1.25 ng of template genomic DNA extracted from the liver of a single adult female N. whitei was loaded on a Chromium Genome Chip. Whole-genome sequencing libraries were prepared using 10× Genomics Chromium Genome Library & Gel Bead Kit v.2, Chromium Genome Chip Kit v.2, Chromium i7 Multiplex Kit, and Chromium controller according to the manufacturer’s instructions with one modification. Briefly, gDNA was combined with Master Mix, a library of Genome Gel Beads, and partitioning oil to create Gel Bead-in-Emulsions (GEMs) on a Chromium Genome Chip. The GEMs were isothermally amplified. Prior to Illumina library construction, the GEM amplification product was sheared on a Covaris E220 Focused Ultrasonicator to ∼350 bp and then converted to a sequencing library following the 10× standard operating procedure. A total of 679.43 million read pairs were sequenced on an Illumina HiSeqX sequencer, and a de novo assembly was constructed with Supernova 2.1.1 (Weisenfeld et al. 2018).

A Chicago library was prepared as described previously (Putnam et al. 2016). Briefly, ∼500 ng of HMW gDNA was reconstituted into chromatin in vitro and fixed with formaldehyde. Fixed chromatin was digested with DpnII, the 5′ overhangs were filled in with biotinylated nucleotides, and then free blunt ends were ligated. After ligation, crosslinks were reversed, and the DNA was purified from protein. Purified DNA was treated to remove biotin that was not internal to ligated fragments. The DNA was then sheared to ∼350 bp mean fragment size and sequencing libraries were generated using NEBNext Ultra enzymes and Illumina-compatible adapters. Biotin-containing fragments were isolated using streptavidin beads before PCR enrichment of each library. The libraries were sequenced on an Illumina HiSeqX to produce 242 million 2× 150 bp paired end reads.

A Dovetail Hi-C library was prepared in a similar manner as described previously (Lieberman-Aiden et al. 2009). For each library, chromatin was fixed in place with formaldehyde in the nucleus and then extracted. Fixed chromatin was digested with DpnII, the 5′ overhangs filled in with biotinylated nucleotides, and then free blunt ends were ligated. After ligation, crosslinks were reversed, and the DNA purified from protein. Purified DNA was treated to remove biotin that was not internal to ligated fragments. The DNA was then sheared to ∼350 bp mean fragment size and sequencing libraries were generated using NEBNext Ultra enzymes and Illumina-compatible adapters. Biotin-containing fragments were isolated using streptavidin beads before PCR enrichment of each library. The libraries were sequenced on an Illumina HiSeqX to produce 179 million 2 × 150 bp paired end reads.

The Supernova de novo assembly built from 10× Chromium data, Chicago library reads, and Dovetail Hi-C library reads were used as input data for assembly scaffolding with HiRise v1 (Putnam et al. 2016). An iterative analysis was conducted. First, Chicago library sequences were aligned to the draft input assembly using a modified SNAP read mapper (http://snap.cs.berkeley.edu). The separations of Chicago read pairs mapped within draft scaffolds were analyzed by HiRise v1 to produce a likelihood model for genomic distance between read pairs, and the model was used to identify and break putative misjoins, to score prospective joins, and make joins above a threshold. After aligning and scaffolding Chicago data, Dovetail Hi-C library sequences were aligned, and scaffolds were generated following the same approach.

Genome annotation

The Rio Pearlfish genome was annotated with the NCBI Euakryotic genome annotation pipeline v9.0 (Thibaud-Nissen et al. 2016) and with MAKER 2.31 (Cantarel et al. 2008; Campbell et al. 2014; Bowman et al. 2017) using protein evidence from 15 fish species (Supplementary Table1) and transcriptome evidence from Rio Pearlfish DIII embryos and hatched larvae (Thompson and Ortí 2016). Genome assembly and annotation completeness (Supplementary Table2) were analyzed with BUSCO v5 (Simão et al. 2015) and CEGMA 2.4 (Parra et al. 2007) via the gVolante server (Nishimura et al. 2017, https://gvolante.riken.jp).

Phylogenetics and orthology

To confirm species identification, we extracted and concatenated the barcoding marker genes cox1 and cytb from our genome assembly, aligned them with orthologous sequences from all 3 described Nematolebias species (Costa et al. 2014) and inferred a phylogeny partitioned by codon and gene with RAxML (Stamatakis 2006, 2014) with the following parameters: -T 4 -N autoMRE -m GTRCAT -c 25 -p 12345 -f a -x 12345 –asc-corr lewis. We used Orthofinder v2.4.1 (Emms and Kelly 2015) to identify orthologous protein sequences between N. whitei and 35 other vertebrates genomes (Supplementary Table3) as well as protein sequences obtained from Cui et al. (2019), Hara et al. (2018), and the longest isoforms of other species available on NCBI RefSeq (last accessed 2021 September 22) downloaded with ortholog’s retrieve longest isoforms function (Drost et al. 2015). The output of Orthofinder (Supplementary Table4) was examined to identify Pearlfish-specific orthogroups. Genes in these orthogroups were used as queries in BLAST searches (e value cutoff of e−3) against Japanese medaka (HdrR strain, assembly ASM223467v1) protein sequences downloaded from Ensembl (last accessed 2022 January 17, Supplementary Table5). We performed a statistical overrepresentation test with a Fisher’s exact test and a false discovery rate correction on the Gene Ontologies (GOs) of these medaka genes using Panther v.16.0 (Mi et al. 2021) with the GO biological processes complete database (Supplementary Table6).

Synteny and genome 3D structure

We examined conservation of synteny using genome assemblies and NCBI annotations for Rio Pearlfish, medaka (oryLat2, UCSC), and zebrafish (GCF_000002035.5_GRCz10, NCBI) as input for SynMap in the online CoGe database and toolkit (Lyons and Freeling 2008; last accessed October 14, 2021). Bwa v 0.7.17 (Li and Durbin 2009) was used to independently map Rio Pearlfish Hi-C read pairs to the genome assembly with the following parameters: bwa mem -A1 -B4 -E50 -L0, and HiCExplorer 3.6 (Ramírez et al. 2018) was used to construct a Hi-C matrix with the resulting bam files as follows: hicBuildMatrix –binSize 10000 –restrictionSequence GATC –danglingSequence GATC. The matrix was corrected via hicCorrectMatrix correct –filterThreshold -1.5 5. The matrix was binned depending on preferred resolution for viewing. Contact maps were visualized with hicPlotMatrix –log1p, and compared with contact maps of syntenic regions in medaka and zebrafish (Nakamura et al. 2021).

Repeat content and repeat landscape

We constructed a species-specific repeat database with Repeat Modeler 2.0.1 (Smit and Hubley 2008). This library as well as vertebrate Repbase annotations (Jurka 2000) (downloaded 15 November 2017), and repeat libraries from platyfish (Schartl et al. 2013), coelacanth (Amemiya et al. 2013), bowfin (Thompson et al. 2021b), and spotted gar (Braasch et al. 2016) were combined to annotate repeat elements with RepeatMasker v4.0.5 (Smit et al. 2013). CalcDivergenceFromAlign.pl and createRepeatLandscape.pl in the RepeatMasker package were used to generate a repeat landscape. We graphically compared the repeat landscape of Rio Pearlfish to those described for other sequenced killifish species (Reichwald et al. 2015; Valenzano et al. 2015; Rhee et al. 2017; Cui et al. 2019) to identify similarities and difference in the magnitude and location of peaks at different Kimura distances in the histograms.

Hatching enzyme gene expression

Aquatic vertebrates hatch by secreting choriolytic enzymes from hatching gland cells (HGCs) that break down the egg chorion (Yamagami 1988; Hong and Saint-Jeannet 2014). Teleost fishes underwent hatching enzyme gene duplications followed by divergence and functional divergence into the high choriolytic enzyme (hce) and low choriolytic enzyme (lce) genes (Yasumasu et al. 1992; Kawaguchi et al. 2006, 2010; Sano et al. 2014). We used BLAST to search the well-studied medaka hatching enzyme paralogs (lce and hce) against the annotated Pearlfish genome. We used the Pearlfish gene sequences from these BLAST hits as well as other metalloprotease gene sequences from medaka, Austrofundulus, Kryptolebias, and Nothobranchius (accession numbers in Supplementary Table7) to infer gene trees. The identified Pearlfish lce and hce genes are orthologous to those of other teleosts (data not shown). Pearlfish hatching enzyme orthologs were examined for transcript evidence from DIII embryos (Thompson and Ortí 2016) to identify active lce and hce gene expression in Pearlfish DIII.

We generated an antisense RNA probe for the Pearfish lce.2 gene and performed whole-mount RNA in situ hybridization to identify hatching enzyme gene expression patterns as markers for the location of HGCs in Pearlfish. Total RNA was extracted from DIII Pearlfish embryos with a Qiagen RNeasy mini plus kit and reverse transcribed with a superscript IV VILO kit (ThermoFisher) according to the manufacturers’ instructions. The lce cDNA was amplified from the reverse transcribed template via PCR (Primers: Nwh_lce.1_1F: 5′-ATGGACCATAAAGCAAAAGTTTCTCTC-3′; Nwh_lce.1_792R: 5′-CTATTGCTTGTATTTTGAACACTTGT-3′; Nwh_lce.2_1F: 5′-ATGGACCATAAAGCAAAAGTTACTCTT-3′; Nwh_lce.2_825R: 5′-CTATTGCTTGTATTTTGAACAGTTGT-3′) and lce.2 was inserted into a TOPO TA cloning kit vector (Invitrogen) according to the manufacturer’s instructions. Whole mount lce.2 mRNA in situ hybridization on manually dechorionated DIII Rio Pearlfish embryos was performed following Deyts et al. (2005) with a 25 µg/ml proteinase K digestion treatment for 45 min (n = 3 embryos), 60 min (n = 3 embryos), and 90 min (n = 2 embryos).

Results and discussion

Genome sequencing and assembly

We report a high-quality, 1.2 Gb chromosome-level genome assembly of N. whitei. The Rio Pearlfish genome assembly consists of 24 chromosomal pseudomolecules represented by 24 superscaffolds that match the described karyotype (n = 24; Van Post, 1965). The assembly has an N50 of over 49.98 Mb and an L50 of 11 scaffolds (Table 1). BUSCO and CEGMA scores for different core gene databases indicate a high-quality assembly with an average of 94% complete BUSCOS and CEGs across all relevant databases (Table 1, Supplementary Table2).

Table 1.

Rio Pearlfish genome assembly (NemWhi1) and annotation statistics.

Genome assembly statistics
 No. of scaffolds 18,999
 No. of base pairs 1,218,332,341
 N50 49,984,095
 L50 11
 N90 32,525,398
 L90 22
 No. of superscaffolds 24
 No. of chromosomes (n)a 24
 GC content 41.8%
 Repeat contentb 57.3%
 BUSCO scoresc: Vertebrata, Actinopterygii 96.9%, 95.5%
 CEGMA scoresc: CEG, CVG 99.19%, 99.57%
Gene annotation statistics
 MAKER annotation
  No. of protein-coding genes 26,016
  BUSCO scoresc: Vertebrata, Actinopterygii 91.4%, 86.2%
 NCBI RefSeq annotationd
  No. of protein-coding genes 21,341
  BUSCO scoresc: Vertebrata, Actinopterygii 97.4%, 96.5%
  No. of genes in orthogroupse 21,176 (99.2%)
  No. of species-specific orthogroups (genes) 17 (42)
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b

See Supplementary Table 10 for more information.

c

See Supplementary Table 2 for more information.

d

See Supplementary Table 8 for more information.

e

See Supplementary Table 4 for more information.

Genome annotation

The NCBI N. whitei Annotation Release 100.20210725 contains 23,038 genes, with 21,341 protein-coding genes, similar to other chromosome-level killifish genome assemblies from N. furzeri and Kryptolebias marmoratus (Reichwald et al. 2015; Valenzano et al. 2015; Kelley et al. 2016; Rhee et al. 2017) (Supplementary Table8). Minor differences in gene numbers among killifish species are likely due to annotation methods and minor species-specific gene losses or expansions. The number and content of annotated genes can be influenced by evidence used for annotation, differences in gene model prediction likelihoods, and postannotation filtering (Holt and Yandell 2011; Campbell et al. 2014). MAKER annotated 26,016 protein-coding genes, on par with the NCBI annotation. See Table 1 and Supplementary Tables 2 and 8 for Rio Pearlfish genome annotation statistics. Although our BUSCO analyses show fewer genes missed by the NCBI annotation (Supplementary Table2), our additional MAKER annotation provides additional, valid gene models missed by the NCBI pipeline. For example, MAKER annotates 28 vertebrate and 27 actinopterygian BUSCOs missed by the NCBI annotation pipeline (Supplementary Table9).

Phylogenetics and orthology

Our orthofinder analysis illustrates the phylogenetic position of Rio Pearlfish among vertebrates (Fig. 1b) and identified 31,317 orthogroups across 36 vertebrate species with 99.2% of Rio Pearlfish genes within orthogroups (Table 1; Supplementary Table 4). We identified 7,287 orthogroups present across all species from sharks to human to Rio Pearlfish, highlighting the utility of the Rio Pearlfish genome to connect species with extreme developmental phenotypes to other vertebrates, including traditional vertebrate model species such as mouse, Xenopus, zebrafish, etc. We confirmed the identity of our genome specimen with barcoding and a molecular phylogeny of cox1 and cytb with its position located within the N. whitei clade of Nematolebias killifishes (Fig. 1c). We found a total of 17 Pearlfish-specific orthogroups comprising a total of 42 protein sequences. For 39 of these, we established homology to a medaka gene class by BLAST (Supplementary Table5) and found an overrepresentation for GO terms related to glycolysis (Supplementary Table6). This may indicate an adaptive expansion of metabolic genes in this species as annual killifishes tolerate anoxia (Podrabsky et al. 2007, 2012; Wagner et al. 2018), severely depress metabolic rate during diapause (Podrabsky and Hand 1999), and drastically increase metabolic rate during fast maturation (Vrtílek et al. 2018) necessary for an annual life cycle. In a separate study, we have also found that genes involved in cell respiration, specifically oxidative phosphorylation, show higher ratios of nonsynonymous/synonymous codon changes in annual killifishes compared to their nonannual counterparts (Thompson et al. 2021a). Together, these observations points to potential positive selection on genes involved in cell respiration in annual killifishes.

Synteny and genome 3D structure

Three-dimensional chromatin structure impacts gene regulation and can manifest as topologically associated domains (TADs) that could represent higher order gene regulatory regions conserved across evolution (Krefting et al. 2018). However, 3D genome structure has thus far remained uncharacterized in annual killifishes. To confirm the quality of the genome assembly and assess the utility of the chromatin conformation data to interrogate 3D genome structure and gene regulation, we constructed a Hi-C contact map showing higher contact frequency within the 24 pearlfish chromosomes (Fig. 1d) than between chromosomes. Using the genome sequence and gene annotations for Rio Pearlfish in synteny comparisons to another atherinomorph teleost, the medaka (separated by ∼85 million years), and the ostariophysian teleost zebrafish (separated by ∼224 million years), we reveal largely conserved synteny with these species (Fig. 2, e and f) across millions of years of teleost evolution (Hughes et al. 2018; Thompson et al. 2021a). We examined a TAD previously shown to be conserved from zebrafish to medaka (Nakamura et al. 2021) and found high frequency of contacts in the syntenic region between rasa1a and mctp1a in Rio Pearlfish liver tissue that resembles contact maps both in a medaka fibroblast cell line and zebrafish whole embryos (Fig. 1g). Hi-C analyses thus confirm the high-quality of our genome assembly as well as the strikingly conserved nature of 3D genome interactions across teleost evolution, developmental stages, and among cell and tissue types.

Fig. 2.

Fig. 2.

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Rio Pearlfish repeat landscape, hatching enzyme genes, and hatching gland location. a) Repeat landscape of mobile genetic elements in Rio Pearlfish showing a high repeat content with 2 peaks at Kimura distance 4 and 21. Insert: Total TE landscape among killifishes with independent, recent expansions in the convergently annual Nothobranchius (Cui et al. 2019) and Nematolebias (this study) compared to the nonannual Kryptolebias (Choi et al. 2020). b) Locations of 5 hatching enzyme genes in the Rio Pearlfish genome expressed during DIII. c, d) Wholemount RNA in situ hybridization for lce.2 in DIII Rio Pearlfish embryos marking hatching gland cells (HGCs) identified in the buccal (BHGCs, red arrows) and pharyngeal (PHGCs, white arrow) cavities.

Repeat content and transposable element landscape

Transposable elements (TEs) are hypothesized to generate novel genetic substrate for adaptations (Casacuberta and González 2013; Feiner 2016; Esnault et al. 2019). Some annual killifish species have expanded TE content compared to nonannual relatives (Cui et al. 2019), and the link between TEs, aging, and human diseases (Bravo et al. 2020) coupled with the rapid senescence of annual killifishes highlights the importance of examining the Pearlfish “mobilome.” We found that the Rio Pearlfish genome is highly repetitive with a repeat content of ∼57% (Fig. 2a and Table 1; Supplementary Table 10) which is substantially elevated compared to a nonannual member of the same South American family, K. marmoratus, with a repeat content of around ∼27% (Rhee et al. 2017; Choi et al. 2020). Similarly, African annual Nothobranchius killifishes have higher TE repeat content than their nonannual relatives (Cui et al. 2019). This pattern might be the result of adaptation to extreme environments as animals, fungi, and plants have co-opted TEs for environmental adaptations to harsh conditions (Casacuberta and González 2013; Esnault et al. 2019) and TEs may play roles in vertebrate adaptive radiations (Feiner 2016). Our findings further highlight the expanded repeat content in annual killifish genomes and the Pearlfish genome provides novel resources to study the role of mobile DNA in extremophiles.

Hatching enzyme gene expression and hatching gland location

While hatching from the egg is a critical time point during animal development, little is known about its genetic regulation and the integration of environmental cues. Additionally, development of HGCs is dynamic among fishes (Inohaya et al. 1995; 1997) as they migrate and localize in different anatomical locations in different species (Korwin-Kossakowski 2012; Shimizu et al. 2014; Nagasawa et al. 2016). Pinpointing HGC location in seasonal killifishes is necessary for understanding the regulation of hatching in extreme environments.

Rio Pearlfish is a tractable model for studying hatching regulation since hatching is easily induced in this species by exposing DIII embryos to water (Thompson and Ortí 2016). Thus, we examined the hatching enzyme gene repertoire and HGC locations in Pearlfish. We identified five expressed hatching enzyme genes (Fig. 2b, 3 hce and 2 lce genes) upon mapping DIII mRNA reads from Thompson and Ortí (2016) to our reference genome assembly. We annotated hce1 and hce2 on chromosome 2 (corresponding to NCBI genes LOC119423801, LOC119423789), and hce3 on chromosome 20 (LOC119426643) and the adjacent lce.1 and lce.2 genes (LOC119418488, LOC119418489) on chromosome 12 that are species-specific tandem duplicates (Fig. 2b) supported by transcript evidence (Thompson and Ortí 2016). Using whole mount RNA in situ hybridization for lce.2 in DIII embryos, we identified HGC locations in the buccal and pharyngeal cavity in Rio Pearlfish (Fig. 2, c and d) similar to HGC localization in the related mummichog or Atlantic killifish (Fundulus heteroclitus) (Kawaguchi et al. 2005) and in medaka (Inohaya et al. 1995).

A pattern of expanded hce genes is also found in other Atherinomorph fishes like medaka. High choriolytic enzyme genes in this clade of teleosts have lost introns (Kawaguchi et al. 2010) and subfunctionalized postduplication with some hce genes performing better at higher or lower salinities in the euryhaline medaka Oryzias javanicus (Takehana et al. 2020) and the Atlantic killifish F. heteroclitus (Kawaguchi et al. 2013b). Furthermore, the duplication of the lce gene in Rio Pearlfish is an example of convergent evolution within teleosts with another lce duplication in stickleback fishes (Kawaguchi et al. 2013a). These findings underscore the commonality of hatching enzyme gene duplications in teleost fishes that provides a model system for studying convergent gene duplications and functional divergence by sub- and neofunctionalizations.

Conclusions

Our chromosome-level, dually annotated genome assembly of the Rio Pearlfish provides a valuable comparative genomics resource strengthening the utility of killifishes for studying aging, suspended animation, and response to environmental stress. The Rio Pearlfish is an emerging “Extremo” Eco-Evo-Devo research organism, and this reference genome will be a substrate for future functional genetic and multiomic approaches exploring how organisms integrate developmental and environmental cues to adapt to extreme environmental conditions in a changing world.

Data availability

The genome sequence, annotation, and sequence read data are available on NCBI under accession GCA_014905685.2 and Bioproject PRJNA560526. The genome assembly and annotation has also been integrated to the University of California Santa Cruz Genome Browser (https://hgdownload.soe.ucsc.edu/hubs/fish/index.html). The MAKER genome annotation is available on github (https://github.com/AndrewWT/NematolebiasGenomics).

Supplemental material is available at G3 online.

Supplementary Material

jkac045_Supplementary_Data
Click here for additional data file. (131.8KB, zip)

Acknowledgments

We thank Camilla Peabody for guidance with RNA in situ hybridization, Kevin Childs for computational resources, and Françoise Thibaud-Nissen for help integrating the genome into NCBI’s Eukaryotic Annotation Pipeline.

AWT and IB conceived the project, wrote the manuscript, and acquired funding; genome sequencing and assembly was performed with Dovetail Genomics; MD, HW, AWT, and IB analyzed hatching enzyme genes; HW and AWT performed RNA in situ hybridization; AWT performed comparative genomic analyses, and genome structure analyses; AWT and IB analyzed the repeat content.

Funding

This work was supported by a National Science Foundation Bio/computational Evolution in Action CONsortium grant to AWT and IB (NSF BEACON, Cooperative Agreement No. DBI-0939454, project #1233) and a National Institutes of Health Office of Research Infrastructure Programs grant to IB (NIH ORIP R01OD011116).

Conflicts of interest

None declared.

Literature cited

  1. Aluru N, Karchner SI, Franks DG, Nacci D, Champlin D, Hahn ME.. Targeted mutagenesis of aryl hydrocarbon receptor 2a and 2b genes in Atlantic killifish (Fundulus heteroclitus). Aquat Toxicol. 2015;158:192–201. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Amemiya CT, Alföldi J, Lee AP, Fan S, Philippe H, Maccallum I, Braasch I, Manousaki T, Schneider I, Rohner N, et al. The African coelacanth genome provides insights into tetrapod evolution. Nature. 2013;496(7445):311–316. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bowman MJ, Pulman JA, Liu TL, Childs KL.. A modified GC-specific MAKER gene annotation method reveals improved and novel gene predictions of high and low GC content in Oryza sativa. BMC Bioinformatics. 2017;18(1):15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Braasch I, Gehrke AR, Smith JJ, Kawasaki K, Manousaki T, Pasquier J, Amores A, Desvignes T, Batzel P, Catchen J, et al. The spotted gar genome illuminates vertebrate evolution and facilitates human-teleost comparisons. Nat Genet. 2016;48(4):427–437. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bravo JI, Nozownik S, Danthi PS, Benayoun BA.. Transposable elements, circular RNAs and mitochondrial transcription in age-related genomic regulation. Development. 2020;147(11):1–18. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Campbell, MS, Holt, C, Moore, B, Yandell, M. Genome annotation and curation using MAKER and MAKER‐P. Curr Protoc Bioinform. 2014;48(1):4–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Cantarel BL, Korf I, Robb SMC, Parra G, Ross E, Moore B, Holt C, Sánchez Alvarado A, Yandell M.. MAKER: an easy-to-use annotation pipeline designed for emerging model organism genomes. Genome Res. 2008;18(1):188–196. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Carter CA, Wourms JP.. Naturally occurring diblastodermic eggs in the annual fish Cynolebias: implications for developmental regulation and determination. J Morphol. 1993;215(3):301–312. [DOI] [PubMed] [Google Scholar]
  9. Casacuberta E, González J.. The impact of transposable elements in environmental adaptation. Mol Ecol. 2013;22(6):1503–1517. [DOI] [PubMed] [Google Scholar]
  10. Choi BS, Park JC, Kim MS, Han J, Kim DH, Hagiwara A, Sakakura Y, Hwang UK, Lee BY, Lee JS. The reference genome of the selfing fish Kryptolebias hermaphroditus: identification of phases I and II detoxification genes. Comp Biochem Physiol Part D Genomics Proteomics. 2020;35:100684. [DOI] [PubMed] [Google Scholar]
  11. Costa WJEM, Amorim PF, Aranha GN.. Species limits and DNA barcodes in Nematolebias, a genus of seasonal Killifishes threatened with extinction from the Atlantic Forest of South-Eastern Brazil, with description of a New Species (Teleostei Rivulidae). Ichthyol Explor Freshwaters. 2014;24:225–236. [Google Scholar]
  12. Costa WJEM. The neotropical seasonal fish genus Nematolebias (Cyprinodontiformes Rivulidae Cynolebiatinae) taxonomic revision with description of a new species. Ichthyol Explor Freshwaters. 2002;1:41–52. [Google Scholar]
  13. Cui R, Medeiros T, Willemsen D, Iasi LNM, Collier GE, Graef M, Reichard M, Valenzano DR.. Relaxed selection limits lifespan by increasing mutation load. Cell. 2019;178(2):385–399. [DOI] [PubMed] [Google Scholar]
  14. Deyts C, Candal E, Joly JS, Bourrat F.. An automated in situ hybridization screen in the medaka to identify unknown neural genes. Dev Dyn. 2005;234(3):698–708. [DOI] [PubMed] [Google Scholar]
  15. Drost HG, Gabel A, Grosse I, Quint M.. Evidence for active maintenance of phylotranscriptomic hourglass patterns in animal and plant embryogenesis. Mol Biol Evol. 2015;32(5):1221–1231. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Emms DM, Kelly S.. OrthoFinder: solving fundamental biases in whole genome comparisons dramatically improves orthogroup inference accuracy. Genome Biol. 2015;16(1):14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Esnault C, Lee M, Ham C, Levin HL.. Transposable element insertions in fission yeast drive adaptation to environmental stress. Genome Res. 2019;29(1):85–95. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Feiner N. Accumulation of transposable elements in hox gene clusters during adaptive radiation of anolis lizards. Proc R Soc B Biol Sci. 2016;283(1840):20161555. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Furness AI, Reznick DN, Springer MS, Meredith RW.. Convergent evolution of alternative developmental trajectories associated with diapause in African and South American killifish. Proc R Soc B. 2015;282(1802):20142189. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Hara Y, Yamaguchi K, Onimaru K, Kadota M, Koyanagi M, Keeley SD, Tatsumi K, Tanaka K, Motone F, Kageyama Y, et al. Shark genomes provide insights into elasmobranch evolution and the origin of vertebrates. Nat Ecol Evol. 2018;2(11):1761–1771. [DOI] [PubMed] [Google Scholar]
  21. Harel I, Benayoun BA, Machado B, Singh PP, Hu C-K, Pech MF, Valenzano DR, Zhang E, Sharp SC, Artandi SE, et al. A platform for rapid exploration of aging and diseases in a naturally short-lived vertebrate. Cell. 2015;160(5):1013–1026. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Holt C, Yandell M.. MAKER2: an annotation pipeline and genome-database management tool for second-generation genome projects. BMC Bioinformatics. 2011;12:491. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Hong CS, Saint-Jeannet JP.. Xhe2 is a member of the astacin family of metalloproteases that promotes Xenopus hatching. Genesis. 2014;52(12):946–951. [DOI] [PubMed] [Google Scholar]
  24. Hughes LC, Ortí G, Huang Y, Sun Y, Baldwin CC, Thompson AW, Arcila D, Betancur-R R, Li C, Becker L, et al. Comprehensive phylogeny of ray-finned fishes (Actinopterygii) based on transcriptomic and genomic data. Proc Natl Acad Sci U S A. 2018;115(24):6249–6252. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Inohaya K, Yasumasu S, Araki K, Naruse K, Yamazaki K, Yasumasu I, Iuchi I, Yamagami K.. Species-dependent migration of fish hatcing gland cells that commonly express astacin-like proteases in common. Dev Growth Differ. 1997;39(2):191–197. [DOI] [PubMed] [Google Scholar]
  26. Inohaya K, Yasumasu S, Ishimaru M, Ohyama A, Iuchi I, Yamagami K.. Temproral and spatial patterns of gene expression for the hatching enzyme in the teleost embryo. Dev Biol. 1995;171(2):374–385. [DOI] [PubMed] [Google Scholar]
  27. Jurka J. Repbase update: a database and an electronic journal of repetitive elements. Trends Genet. 2000;16(9):418–420. [DOI] [PubMed] [Google Scholar]
  28. Kawaguchi M, Hiroi J, Miya M, Nishida M, Iuchi I, Yasumasu S. Intron-loss evolution of hatching enzyme genes in teleostei. BMC Evol Biol. 2010;10:1–10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kawaguchi M, Takahashi H, Takehana Y, Naruse K, Nishida M, Yasumasu S.. Sub-functionalization of duplicated genes in the evolution of nine-spined stickleback hatching enzyme. J Exp Zool B Mol Dev Evol. 2013a;320(3):140–150. [DOI] [PubMed] [Google Scholar]
  30. Kawaguchi M, Yasumasu S, Hiroi J, Naruse K, Inoue M, Iuchi I.. Evolution of teleostean hatching enzyme genes and their paralogous genes. Dev Genes Evol. 2006;216(12):769–784. [DOI] [PubMed] [Google Scholar]
  31. Kawaguchi M, Yasumasu S, Shimizu A, Hiroi J, Yoshizaki N, Nagata K, Tanokura M, Iuchi I.. Purification and gene cloning of Fundulus heteroclitus hatching enzyme: a hatching enzyme system composed of high choriolytic enzyme and low choriolytic enzyme is conserved between two different teleosts, Fundulus heteroclitus and medaka Oryzias latipes. FEBS J. 2005;272(17):4315–4326. [DOI] [PubMed] [Google Scholar]
  32. Kawaguchi M, Yasumasu S, Shimizu A, Kudo N, Sano K, Iuchi I, Nishida M.. Adaptive evolution of fish hatching enzyme: one amino acid substitution results in differential salt dependency of the enzyme. J Exp Biol. 2013b;216(Pt 9):1609–1615. [DOI] [PubMed] [Google Scholar]
  33. Kelley JL, Yee MC, Brown AP, Richardson RR, Tatarenkov A.. The genome of the self-fertilizing mangrove rivulus fish, Kryptolebias marmoratus: a model for studying phenotypic plasticity and adaptations to extreme environments. Genome Biol Evol. 2016;8(7):2145–2154. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Korwin-Kossakowski M. Fish hatching strategies: a review. Rev Fish Biol Fisheries. 2012;22(1):225–240. [Google Scholar]
  35. Krefting J, Andrade-Navarro MA, Ibn-Salem J.. Evolutionary stability of topologically associating domains is associated with conserved gene regulation. BMC Biol. 2018;16(1):1–12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Li H, Durbin R.. Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics. 2009;25(14):1754–1760. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science. 2009;326(5950):289–293. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Lyons E, Freeling M.. How to usefully compare homologous plant genes and chromosomes as DNA sequences. Plant J. 2008;53(4):661–673. [DOI] [PubMed] [Google Scholar]
  39. Mi H, Ebert D, Muruganujan A, Mills C, Albou L-P, Mushayamaha T, Thomas PD.. PANTHER version 16: a revised family classification, tree-based classification tool, enhancer regions and extensive API. Nucleic Acids Res. 2021;49(D1):D394–D403. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Myers GS. Annual fishes. Aquarium J. 1952;23:125–141. [Google Scholar]
  41. Myers GS. Studies on South American freshwater fishes I. Stanford Ichthyol Bull. 1942;2:89–114. [Google Scholar]
  42. Nagasawa T, Kawaguchi M, Yano T, Sano K, Okabe M, Yasumasu S.. Evolutionary changes in the developmental origin of hatching gland cells in basal ray-finned fishes. Zool Sci. 2016;33(3):272–281. [DOI] [PubMed] [Google Scholar]
  43. Nakamura R, Motai Y, Kumagai M, Wike CL, Nishiyama H, Nakatani Y, Durand NC, Kondo K, Kondo T, Tsukahara T, et al. CTCF looping is established during gastrulation in medaka embryos. Genome Res. 2021;31(6):968–980. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Nishimura O, Hara Y, Kuraku S.. GVolante for standardizing completeness assessment of genome and transcriptome assemblies. Bioinformatics. 2017;33(22):3635–3637. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Parra G, Bradnam K, Korf I.. CEGMA: a pipeline to accurately annotate core genes in eukaryotic genomes. Bioinformatics. 2007;23(9):1061–1067. [DOI] [PubMed] [Google Scholar]
  46. Podrabsky JE, Hand SC.. The bioenergetics of embryonic diapause in an annual Killifish, Austrofundulus limneaus. J Exp Biol. 1999;202(19):2567–2580. [DOI] [PubMed] [Google Scholar]
  47. Podrabsky JE, Lopez JP, Fan TWMM, Higashi R, Somero GN.. Extreme anoxia tolerance in embryos of the annual killifish Austrofundulus limnaeus: insights from a metabolomics analysis. J Exp Biol. 2007;210(Pt 13):2253–2266. [DOI] [PubMed] [Google Scholar]
  48. Podrabsky JE, Menze MA, Hand SC.. Long-term survival of anoxia despite rapid ATP decline in embryos of the annual killifish Austrofundulus limnaeus. J Exp Zool A Ecol Genet Physiol. 2012;317(8):524–532. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Putnam NH, O'Connell BL, Stites JC, Rice BJ, Blanchette M, Calef R, Troll CJ, Fields A, Hartley PD, Sugnet CW, et al. Chromosome-scale shotgun assembly using an in vitro method for long-range linkage. Genome Res. 2016;26(3):342–350. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Ramírez F, , BhardwajV, , ArrigoniL, , LamKC, , GrüningBA, , VillavecesJ, , HabermannB, , AkhtarA, , Manke T.. High-resolution TADs reveal DNA sequences underlying genome organization in flies. Nat Commun. 2018;9(1):189. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Reichwald K, Petzold A, Koch P, Downie BR, Hartmann N, Pietsch S, Baumgart M, Chalopin D, Felder M, Bens M, et al. Insights into sex chromosome evolution and aging from the genome of a short-lived fish. Cell. 2015;163(6):1527–1538. [DOI] [PubMed] [Google Scholar]
  52. Rhee JS, Choi BS, Kim J, Kim BM, Lee YM, Kim IC, Kanamori A, Choi IY, Shartl M, Lee JS. Diversity, distribution, and significance of transposable elements in the genome of the only selfing hermaphroditic vertebrate Kryptolebias marmoratus. Sci Rep. 2017;7(1):1–10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Ruijter JM, Creuwels LAJM.. The ultrastructure of prolactin cells in the annual cyprinodont Cynolebias whitei during its life cycle. Cell Tissue Res. 1988;253(2):477–483. [DOI] [PubMed] [Google Scholar]
  54. Ruijter JM, Van Kemenade JAM, Bonga SEW.. Environmental influences on prolactin cell development in the cyprinodont fish, Cynolebias whitei. Cell Tissue Res. 1984;238(3):595–600. [DOI] [PubMed] [Google Scholar]
  55. Ruijter JM. Development and aging of the teleost pituitary: qualitative and quantitative observations in the annual cyprinodont Cynolebias whitei. Anat Embryol (Berl). 1987;175(3):379–386. [DOI] [PubMed] [Google Scholar]
  56. Sano K, Kawaguchi M, Watanabe S, Yasumasu S.. Neofunctionalization of a duplicate hatching enzyme gene during the evolution of teleost fishes. BMC Evol Biol. 2014;14:221. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Schartl M, Walter RB, Shen Y, Garcia T, Catchen J, Amores A, Braasch I, Chalopin D, Volff J-N, Lesch K-P, et al. The genome of the platyfish, Xiphophorus maculatus, provides insights into evolutionary adaptation and several complex traits. Nat Genet. 2013;45(5):567–572. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Schoots AFM, Ruijter JM, van Kemenade JAM, Denucé JM.. Immunoreactive prolactin in the pituitary gland of cyprinodont fish at the time of hatching. Cell Tissue Res. 1983;233(3):611–618. [DOI] [PubMed] [Google Scholar]
  59. Shimizu D, Kawaguchi M, Sano K, Yoshizaki N, Shimizu D, Fujinami Y, Noda T, Yasumasu S.. Comparison of hatching mode in pelagic and demersal eggs of two closely related species in the order Pleuronectiformes. Zoolog Sci. 2014;31(11):709–715. [DOI] [PubMed] [Google Scholar]
  60. Simão FA, Waterhouse RM, Ioannidis P, Kriventseva EV, Zdobnov EM.. BUSCO: assessing genome assembly and annotation completeness with single-copy orthologs. Bioinformatics. 2015;31(19):3210–3212. [DOI] [PubMed] [Google Scholar]
  61. Simpson BRC. The phenology of annual killifish. Symp Zool Soc Lon. 1979;44:243–261. [Google Scholar]
  62. Smit AFA, Hubley R. RepeatModeler Open-1.0; 2008. http://www.repeatmasker.org.
  63. Smit AFA, Hubley R, Green P. RepeatMasker Open-4.0; 2013.http://www.repeatmasker.org.
  64. Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30(9):1312–1313. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Stamatakis A. RAxML-VI-HPC: maximum likelihood-based phylogenetic analyses with thousands of taxa and mixed models. Bioinformatics. 2006;22(21):2688–2690. [DOI] [PubMed] [Google Scholar]
  66. Takehana Y, Zahm M, Cabau C, Klopp C, Roques C, Bouchez O, Donnadieu C, Barrachina C, Journot L, Kawaguchi M, et al. Genome sequence of the euryhaline Javafish Medaka, Oryzias javanicus: a small aquarium fish model for studies on adaptation to salinity. G3 (Bethesda). 2020;10(3):907–915. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Thibaud-Nissen F, DiCuccio M, Hlavina W, Kimchi A, Kitts PA, Murphy TD, Pruitt KD, Souvorov A.. P8008 the NCBI eukaryotic genome annotation pipeline. J Anim Sci. 2016;94(Suppl 4):184. [Google Scholar]
  68. Thompson AW, Black AC, Huang Y, Shi Q, Furness AI, et al. Deterministic shifts in molecular evolution correlate with convergence to annualism in killifishes. BioRxiv, 2021. a, https://doi.org/10.1101/2021.08.09.455723 [Google Scholar]
  69. Thompson AW, Furness AI, Stone C, Rade C, Ortí G.. Microanatomical diversification of the zona pellucida in aplochelioid killifishes. J Fish Biol. 2017a;91(1):126–143. [DOI] [PubMed] [Google Scholar]
  70. Thompson AW, Hawkins MB, Parey E, Wcisel DJ, Ota T, Kawasaki K, Funk E, Losilla M, Fitch OE, Pan Q, et al. The bowfin genome illuminates the developmental evolution of ray-finned fishes. Nat Genet. 2021b;53(9):1373–1384. [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Thompson AW, Hayes A, Podrabsky JE, Ortí G.. Gene expression during delayed hatching in fish-out-of-water. Ecol. Genet. Genomics. 2017b;3–5:52–59. [Google Scholar]
  72. Thompson AW, Ortí G.. Annual killifish transcriptomics and candidate genes for metazoan diapause. Mol Biol Evol. 2016;33(9):2391–2395. [DOI] [PubMed] [Google Scholar]
  73. Valenzano DR, Benayoun BA, Singh PP, Zhang E, Etter PD, Hu C-K, Clément-Ziza M, Willemsen D, Cui R, Harel I, et al. The African Turquoise Killifish genome provides insights into evolution and genetic architecture of lifespan. Cell. 2015;163(6):1539–1554. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Valenzano DR, Sharp SC, Brunet A.. Transposon-mediated transgenesis in the short-lived African Killifish Nothobranchius furzeri, a vertebrate model for aging. G3 (Bethesda). 2011;1(7):531–538. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Van Post A. Vergleichende Untersuchungen der Chromosomenzahlen bei Süßwasser Teleosteern. Zeitschrift Für Zool Syst Evol. 1965;3(1–2):47–93. [Google Scholar]
  76. Vrtílek M, Žák J, Pšenička M, Reichard M.. Extremely rapid maturation of a wild African annual fish. Curr Biol. 2018;28(15):R822–R824. [DOI] [PubMed] [Google Scholar]
  77. Wagner JT, Singh PP, Romney AL, Riggs CL, Minx P, Woll SC, Roush J, Warren WC, Brunet A, Podrabsky JE, et al. The genome of Austrofundulus limnaeus offers insights into extreme vertebrate stress tolerance and embryonic development. BMC Genomics. 2018;19(1):21. [DOI] [PMC free article] [PubMed] [Google Scholar]
  78. Weisenfeld NI, Kumar V, Shah P, Church DM, Jaffe DB.. Corrigendum: direct determination of diploid genome sequences. Genome Res. 2018;28(4):606.1–767. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Wourms JP. Annual fishes. In: Methods in Developmental Biology. New York: Thomas and Crowell Company; 1967. p. 123–137. [Google Scholar]
  80. Wourms JP. The developmental biology of annual fishes I. Stages in the normal development of Austrofundulus myersi Dahl. J Exp Zool. 1972a;182(2):143–168. [DOI] [PubMed] [Google Scholar]
  81. Wourms JP. The developmental biology of annual fishes II. Naturally occurring dispersion and reaggregation of blastomeres during the development of annual fish eggs. J Exp Zool. 1972b;182(2):169–200. [DOI] [PubMed] [Google Scholar]
  82. Wourms JP. The developmental biology of annual fishes III. Pre-embryonic and embryonic diapause of variable duration in the eggs of annual fishes. J Exp Zool. 1972c;182(3):389–414. [DOI] [PubMed] [Google Scholar]
  83. Yamagami K. Mechanisms of hatching in fish. In: Fish Physiology. Cambridge, Massachusetts: Academic Press; 1988. p. 447–499. [Google Scholar]
  84. Yasumasu S, Yamada K, Akasaka K, Mitsunaga K, Iuchi I, Shimada H, Yamagami K.. Isolation of cDNAs for LCE and HCE, two constituent proteases of the hatching enzyme of Oryzias latipes, and concurrent expression of their mRNAs during development. Dev Biol. 1992;153(2):250–258. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

jkac045_Supplementary_Data
Click here for additional data file. (131.8KB, zip)

Data Availability Statement

The genome sequence, annotation, and sequence read data are available on NCBI under accession GCA_014905685.2 and Bioproject PRJNA560526. The genome assembly and annotation has also been integrated to the University of California Santa Cruz Genome Browser (https://hgdownload.soe.ucsc.edu/hubs/fish/index.html). The MAKER genome annotation is available on github (https://github.com/AndrewWT/NematolebiasGenomics).

Supplemental material is available at G3 online.

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