HOMEOBOX-containing genes, and the homeodomain-containing transcription factors that they encode, have crucial functions in most aspects of cellular function and embryonic development in both animals and plants (Bürglin and Affolter 2016). They were also some of the first examples discovered of invertebrate developmental genes that are highly conserved in vertebrates (Carrasco et al. 1984; Gehring 1985). One important subclass of homeodomain proteins are NK proteins. NK genes are evolutionarily ancient and are part of the ANTP megacluster, which also includes Hox and ParaHox genes. NK proteins have fundamental roles in the development of mesoderm, endoderm, the nervous system, and the heart in all bilaterian animals examined so far (Wotton et al. 2010; Holland 2013; Treffkorn et al. 2018), and they are also found in sponges, one of the most basal animals still alive, and potentially the sister group to all other animals (Larroux et al. 2007; Fortunato et al. 2014; Pisani et al. 2015; Simion et al. 2017).

Hmx proteins (H6 Family Homeodomain proteins, previously called Nk5 or Nkx5 proteins; see Supplemental Material, Table S1) are a key subfamily of NK proteins. In vertebrates there are usually three or four different Hmx genes, as Hmx4 is only found in some species (Wotton et al. 2010). Interestingly, Hmx2 and Hmx3 are usually located adjacent to each other on the same chromosome, and this is also the case for Hmx1 and Hmx4, suggesting that both pairs of genes arose from tandem duplication events rather than the two rounds of whole-genome duplication that occurred at the base of the vertebrates (Wotton et al. 2010). In teleosts, there are occasionally extra duplicates of one or more of these genes as the result of the additional genome duplication in this lineage, although interestingly, the retained genes are not consistent between different teleost species (Wotton et al. 2010). In zebrafish there are two hmx3 genes, hmx3a and hmx3b, but only one hmx1, hmx2, and hmx4 gene.

Previous research has shown that Hmx2 and Hmx3 have crucial functions in ear development in mouse, and our recent work shows that this is also the case for Hmx3a in zebrafish (Wang et al. 1998; Wang et al. 2001; Wang et al. 2004; Wang and Lufkin 2005; Hartwell et al. 2019). In mouse, both Hmx2 and Hmx3 mutants have ear defects and these are more severe in double mutants (Wang et al. 2001; Wang et al. 2004). Hmx2 and Hmx3 are also required for correct specification of the mouse hypothalamus (Wang et al. 2004), and morpholino knockdown experiments have suggested that they are required for correct lateral line development in zebrafish (Feng and Xu 2010). Hmx2 and Hmx3 are also expressed in two distinct domains in mouse spinal cord, but the spinal cord functions of these genes are unknown (Bober et al. 1994; Wang et al. 2000; Wang et al. 2004).

Here, we show that zebrafish hmx2 and hmx3a are coexpressed in spinal dI2 and V1 interneurons, whereas hmx3b, hmx1, and hmx4 are not expressed in spinal cord. Using knockdown and mutational analyses, we demonstrate that, in addition to its role in ear development, hmx3a is required for correct specification of a subset of spinal cord interneuron neurotransmitter phenotypes as well as lateral line progression and viability (survival to adulthood). Our data suggest that in the absence of functional Hmx3a protein, a subset of dI2 spinal interneurons switch their neurotransmitter phenotype from glutamatergic (excitatory) to GABAergic (inhibitory). This is important because currently very little is known about how dI2 spinal interneuron neurotransmitter phenotypes are specified, or indeed, how spinal cord excitatory neurotransmitter phenotypes in general are specified, and if neurons do not acquire the correct neurotransmitter phenotypes, they cannot function appropriately in spinal cord circuitry.

Surprisingly, despite the fact that hmx2 and hmx3a have similar expression patterns during embryonic development and both genes are required for correct ear development in mouse, our mutational analyses did not uncover any requirement for hmx2, by itself, in viability, ear development, lateral line progression, or specification of spinal cord interneuron neurotransmitter phenotypes in zebrafish. This is surprising, especially given that embryos injected with a hmx2 morpholino have reduced numbers of spinal cord glutamatergic neurons and a corresponding increase in the number of inhibitory spinal cord neurons, and that embryos injected with both hmx2 and hmx3a morpholinos have more severe spinal cord phenotypes than single knockdown (SKD) embryos. Zebrafish hmx2 mutants are viable and have no obviously abnormal phenotypes in these sensory structures and neurons that require hmx3a, even when almost all of the hmx2 locus is deleted. (In our most severe mutant allele, hmx2SU39, only 84 nucleotides of 5′ and 60 nucleotides of 3′ coding sequence remain). In addition, zebrafish embryos homozygous for deletions of both hmx2 and hmx3a have identical phenotypes to severe hmx3a single mutants. However, mutating hmx2 in hypomorphic hmx3aSU42 mutants, that usually develop normally, results in abnormal ear and lateral line progression phenotypes, suggesting that while hmx2 cannot compensate for mutations in hmx3a, it does function in these developmental processes, although to a much lesser extent than hmx3a. Our analyses of homozygous mutant phenotypes for several different hmx3a mutant alleles also suggest that Hmx3a may not require its homeodomain for its roles in viability or embryonic development. This is surprising, as homeodomain proteins usually function by binding DNA through their homeodomain and regulating gene expression. In contrast, our mutational analyses suggest that Hmx3a may only require its N-terminal domain for its vital functions in viability and sensory organ and spinal cord interneuron development.

Materials and MethodsEthics statement

All zebrafish experiments in this research were carried out in accordance with the recommendations and approval of either the UK Home Office or the Syracuse University Institutional Animal Care and Use Committee.

Zebrafish husbandry and fish lines

Zebrafish (Danio rerio) were maintained on a 14-hr light/10-hr dark cycle at 28.5°. Embryos were obtained from natural paired and/or grouped spawnings of wild-type (WT; AB, TL, or AB/TL hybrid) fish; heterozygous or homozygous hmx2, hmx3a, or hmx2;3a mutants (created as part of this study and reported here); Tg(evx1:EGFP)SU1 or Tg(evx1:EGFP)SU2 transgenic fish (Juárez-Morales et al. 2016); Tg(UAS:mRFP) transgenic fish (Balciuniene et al. 2013); heterozygous mindbomb1 (mib1ta52b) mutants (Jiang et al. 1996); and heterozygous or homozygous hmx3asa23054 mutants (Kettleborough et al. 2013).

CRISPR mutagenesis and screening

The hmx3aSU3 allele was described previously (Hartwell et al. 2019). With the exception of hmx3asa23054 (generated in the Zebrafish Mutation Project and obtained from the Zebrafish International Resource Centre), we created all of the other hmx2, hmx3a, and hmx2;hmx3a double deletion mutants described in this paper using CRISPR mutagenesis. For all alleles, other than the hmx2MENTHU allele, we designed and synthesized single guide RNA (sgRNA) and Cas9 messenger RNA (mRNA) as in Hartwell et al. (2019). For the hmx2MENTHU allele, we designed the CRISPR RNA (crRNA) using the Microhomology-mediated End joining kNockout Target Heuristic Utility (MENTHU) tool (version 2.1.2), in the Gene Sculpt Suite (Ata et al. 2018; Mann et al. 2019). The MENTHU allele crRNA design was verified with CHOPCHOP (version 3.0.0) (Montague et al. 2014; Labun et al. 2016; Labun et al. 2019) and the CRISPR-Cas9 guide RNA design checker tool (Integrated DNA Technologies). The hmx2MENTHU crRNA was purchased together with a universal 67mer trans-activating CRISPR RNA (tracrRNA) (1072533) and Alt-R S.p. Cas9 Nuclease V3 (1081058) from Integrated DNA Technologies. See Table S2 for guide RNA sequences and Figure 4 for their genomic locations. hmx2SU35, hmx2SU36, hmx2SU37, and hmx3aSU42 alleles were all generated with a single sgRNA (sgRNA E for hmx2SU35, hmx2SU36, and hmx2SU37; sgRNA B for hmx3aSU42; Table S2; Figure 4). hmx2MENTHU was generated with a single crRNA (sgRNA D; Table S2; Figure 4). hmx2SU38, hmx2SU39, hmx2;hmx3aSU44, and hmx2;hmx3aSU45 alleles are all deletions, generated by combinatorial use of two sgRNAs. The hmx2SU38 sgRNAs (sgRNAs E and F; Table S2; Figure 4) flank the homeodomain. For hmx2SU39 we used the same 3′ sgRNA and a more 5′ sgRNA (sgRNAs C and F; Table S2; Figure 4). To make hmx2;hmx3a double deletion alleles, we designed sgRNAs that flanked the two genes, which are adjacent on chromosome 17 (sgRNAs A and F; Table S2; Figure 4). In all cases except the hmx2MENTHU mutant, we injected 2–4 nl of a mixture of 200 ng/µl of each sgRNA + 600 ng/µl nls-ZCas9-nls mRNA into the single cell of a one-cell stage AB WT embryo. To create the hmx2MENTHU mutant allele, we injected 1 nl of a 5 µM crRNA:tracrRNA:Cas9 ribonucleoprotein complex into the single cell of a very early one-cell stage embryo from an incross of heterozygous hmx3aSU42 fish. The 5 µM crRNA:tracrRNA:Cas9 ribonucleoprotein complex was synthesized as described in Hoshijima et al. (2019). hmx3aSU43 is a hsp70:Gal4 knock-in allele. We co-injected a donor template containing Gal4, under the control of a minimal hsp70 promoter (pMbait-hsp70:Gal4, a kind gift of Dr Shin-ichi Higashijima; Kimura et al. 2014) with two sgRNA molecules: one specifically targeting hmx3a (sgRNA B; see Table S2 and Figure 4), and one (Mbait sgRNA, GGCTGCTGCGGTTCCAGAGG) specifically linearizing the donor template in vivo, into the single cell of one-cell stage embryos from an incross of heterozygous Tg(UAS:mRFP) fish. For these experiments, embryos were injected with 2–4 nl of a mixture of 130 ng/µl of each sgRNA + 180 ng/µl nlz-ZCas9-nls mRNA + 66 ng/µl pMbait-hsp70:Gal4 donor DNA. We screened injected embryos for RFP fluorescence in patterns consistent with hmx3a expression (i.e., ear, lateral line primordium, and/or spinal cord) from 1 day postfertilization (d) onward, and raised injected embryos displaying appropriate expression patterns to adulthood. We then assessed germline transmission by outcrossing to heterozygous or homozygous Tg(UAS:mRFP) fish. Gal4 expression in hmx3aSU43 recapitulates hmx3a spinal expression but is not expressed in the ear or lateral line primordium (data not shown).

We identified founder fish for hmx2SU35, hmx2SU36, hmx2SU37, and hmx3aSU42 alleles using high-resolution melt analysis (HRMA), and the supermix and amplification programs described in Hartwell et al. (2019). For the PCRs described below, we used Phusion High-Fidelity DNA Polymerase (M0530L; New England BioLabs Inc.) unless otherwise stated. HRMA primers and PCR primers for sequencing are provided in Table S2.

We used the following PCR conditions to identify hmx2SU38 founder fish: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 67.0° for 30 sec, 72.0° for 40 sec; followed by a final extension at 72.0° for 5 min. We distinguished the mutant allele by gel electrophoresis on a 1% TAE agarose gel (110V for 30 min). The WT allele generated a 1098 bp product, compared with a 671 bp mutant allele product. The PCR primer sequences are provided in Table S2.

We used nested PCR to identify hmx2SU39 founder fish, with the following conditions: nested PCR 1: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 69.0° for 20 sec, 72.0° for 75 sec; followed by a final extension at 72.0° for 5 min. The mutant allele was distinguished by gel electrophoresis on a 1% TAE agarose gel (110V for 30 min). The WT allele generated a 2012 bp product (which may or may not be detected on the gel), compared with a 576 bp mutant allele product. We then diluted the nested 1 PCR product 1:5 in sterile distilled water and performed the nested PCR 2 reaction using the following conditions: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 66.0° for 20 sec, 72.0° for 60 sec; followed by a final extension at 72.0° for 5 min. The mutant allele was distinguished by gel electrophoresis on a 1% TAE agarose gel (110V for 30 min). The WT allele generated a 1758 bp product compared with a 322 bp mutant allele product. All PCR primer sequences are provided in Table S2.

We identified hmx2MENTHU F0 embryos by PCR, followed by sequencing with the forward primer that generated the amplicon (Table S2). The PCR was performed on DNA extracted from individual embryos using the following conditions: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 64.0° for 20 sec, 72.0° for 15 sec; followed by a final extension at 72.0° for 5 min. We assayed that the PCR was successful by gel electrophoresis on a 2.5% TBE agarose gel (100V for 40 min). The PCR generates a 155 bp product. The PCR product was purified using EZ-10 Spin Column PCR Products Purification Kit (BS664; Bio Basic) and eluted in 30 µl sterile water before sequencing.

We used either assessment of germline transmission, as described above, or PCR to identify hmx3aSU43 founder fish. PCR conditions were 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 69.0° for 20 sec, 72.0° for 60 sec; followed by a final extension at 72.0° for 5 min. The mutant allele was distinguished by gel electrophoresis on a 1% TAE agarose gel (110V for 30 min). A 1471 bp PCR product was only generated by fish heterozygous for the allele. It was not produced from WT animals since the reverse primer only recognizes the inserted donor DNA sequence. The PCR primer sequences are provided in Table S2.

We identified hmx2;hmx3aSU44 and hmx2;hmx3aSU45 founder fish by nested PCR, using the following conditions: nested PCR 1: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 67.0° for 20 sec, 72.0° for 30 sec; followed by a final extension at 72.0° for 5 min. The mutant allele was distinguished by gel electrophoresis on a 1% TAE agarose gel (110V for 30 min). The WT product was too large to be generated by these PCR conditions, so only heterozygous animals are detected by the presence of a 514 bp product on the gel. We then diluted the nested 1 PCR 1:5 in sterile distilled water and performed the nested PCR 2 reaction using the following conditions: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 66.0° for 20 sec, 72.0° for 30 sec; followed by a final extension at 72.0° for 5 min. The mutant allele was distinguished by gel electrophoresis on a 1% TAE agarose gel (110V for 30 min). Again, the WT product was too large to be generated by these PCR conditions, so only heterozygous animals were detected by the presence of a 445 bp product.

Once stable lines were established, we identified hmx2SU35 fish by PCR, followed by sequencing (the mutation introduces a 1 bp insertion that cannot be resolved by restriction digestion, and we cannot distinguish heterozygotes from homozygotes using HRMA). We performed this PCR using Taq DNA Polymerase (M0320S; New England BioLabs Inc.) and the following conditions: 95.0° for 30 sec; 35 cycles of 95.0° for 20 sec, 52.0° for 30 sec, 68.0° for 45 sec; followed by a final extension at 68.0° for 5 min. The PCR primer sequences are provided in Table S2. We used HRMA and the conditions described above to identify hmx2SU36 stable mutants. Homozygous mutants segregate from heterozygous animals by the scale of their deflection in the HRMA plot. We identified hmx2SU37 mutants by performing the PCR used to sequence hmx2SU35 stable mutants (see above and Table S2). When we analyzed the products on a 1% TAE gel (110V for 30 min), the WT allele generated a 580 bp product, compared with a 528 bp mutant product. We identified hmx2SU38 stable mutants using the same PCR conditions initially used to identify founders (see above and Table S2). We used the same nested PCR conditions to identify hmx2SU39 mutants. However, the WT product was not always visible on the gel. Therefore, we also performed a WT amplicon PCR identical to that described above for identifying stable hmx2SU35 fish, as this genomic region is only present in WT and heterozygous animals (see also Table S2).

We identified stable hmx3aSU42 mutants by PCR, using Taq DNA Polymerase (M0320S; New England BioLabs Inc.) and the following conditions: 94.0° for 2 min; 35 cycles of 94.0° for 30 sec, 64.9° for 30 sec, 72.0° for 30 sec; followed by a final extension at 72.0° for 2 min. The PCR primer sequences are provided in Table S2. While the mutant PCR product (321 bp) could sometimes be distinguished from the WT product (331 bp) by running on a 2% TBE gel (100V for 55 min), the mutation also deletes a BanI restriction site. Following digestion with BanI (R0118S; New England BioLabs Inc.), the products were run on a 2% TBE gel (100V for 40 min). The WT amplicon digested to completion, producing 120 bp + 211 bp bands, whereas the mutant product did not cut. We identified stable hmx3aSU3 mutants by running the same PCR used to identify hmx3aSU42 mutants (see above and Table S2). The insertion in hmx3aSU3 was easily visualized on a 2% TBE gel. The WT product was 331 bp, compared to a mutant product of 400 bp. Since the PCR used to detect hmx3aSU43 mutants was specific to the inserted donor DNA, and the WT amplicon in hmx2;hmx3aSU44 and hmx2;hmx3aSU45 mutants was too large to detect using the nested PCR conditions, for these alleles we also performed a WT amplicon PCR to distinguish WTs from heterozygotes. The WT amplicon PCR was identical to that performed before BanI digestion on hmx3aSU42 mutants (see above and Table S2). For hmx3aSU43, the WT amplicon PCR results were compared to the PCR results (identical PCR to that first used to identify founders, see above), and for hmx2;hmx3aSU44 and hmx2;hmx3aSU45, the WT amplicon PCR results were compared to the nested 2 PCR results (identical nested 2 PCR to that first used to identify founders, see above and Table S2).

In all cases, stable F1 heterozygous fish were confirmed by sequencing. To further confirm the mutant sequences of hmx2SU39 and hmx3aSU42, we extracted total RNA from embryos produced by incrosses of homozygous viable adults using TRIzol Reagent (15596018; Thermo Fisher Scientific) and the RNeasy Mini Kit (74104; QIAGEN, Valencia, CA). Total RNA was converted to complementary DNA (cDNA) using the iScript cDNA synthesis kit (1708891; Bio-Rad, Hercules, CA). We performed transcript-specific PCRs using the following primers and conditions: hmx2-forward: TGAACTGTTATGAGACGAGAATGAA and hmx2-reverse: GTGTATTTTGTACGTCTTAGTGTGTGT (PCR: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 64.2° for 20 sec, 72.0° for 30 sec; followed by final extension at 72.0° for 5 min); or hmx3a-forward: AACCGCGTTTAAGTTCCCATTG and hmx3a-reverse: GTGCGAGTAGTAAACCGGATGAG (PCR: 98.0° for 30 sec; 35 cycles of 98.0° for 10 sec, 71.0° for 20 sec, 72.0° for 30 sec; followed by final extension at 72.0° for 5 min). We then confirmed these homozygous mutant transcript sequences by sequencing.

Morpholino injections

For SKD translation-blocking experiments, 3.5 nl of a mixture containing either 2 ng/nl of a translation-blocking hmx2 morpholino (MO) (5′ TTCCGCTGTCCTCCGAATTATTCAT) or 2 ng/nl of a translation-blocking hmx3a morpholino (5′ ACGTATCCTGTGTTGTTTCGGGCAT) plus 5 ng/nl of a control zebrafish p53 morpholino (5′ GCGCCATTGCTTTGCAAGAATTG) was injected into the single cell of a one-cell stage WT embryo. For double knockdown (DKD) experiments with translation-blocking morpholinos, 3.5 nl of a mixture containing 2 ng/nl of both translation-blocking hmx morpholinos plus 5 ng/nl of the control zebrafish p53 morpholino was injected. For DKD splice-blocking experiments, 4 nl of a mixture containing 5 ng/nl of both a splice-blocking hmx2 morpholino (5′ GGCACCTGCAACCAATGCGACACAC) and a splice-blocking hmx3a morpholino (5′ TGCTGCTACAGTAATAGAGGCCAAA), plus 7 ng/nl of the control zebrafish p53 morpholino was injected (all morpholinos obtained from Gene Tools). DKD but not SKD embryos exhibit delayed development from somitogenesis stages onward when compared to uninjected controls. To circumvent this, they were incubated at 32° from 9 hours post fertilization (hpf) onward, whereas the uninjected controls remained at 28.5°. This ensured that control and injected embryos reached the desired developmental stage of 27 hpf at approximately the same time. The lateral line primordium does not migrate in DKD animals, so this could not be used to stage injected embryos. Instead, these embryos were visually inspected and fixed when they displayed the same head-trunk angle, head size, and eye size as prim-staged uninjected control embryos (Kimmel 1995). Migration of the lateral line primordium is unaffected in SKD embryos, so these were prim-staged before fixing for experiments (Kimmel 1995). Morpholino injections always produce a spectrum of phenotypes, since it is hard to ensure that every cell receives the same dose. Therefore, before fixing at 27 hpf, we removed any embryos with severely abnormal morphology (stunted length and/or severely developmentally delayed, likely caused by receiving too much morpholino). Embryos injected with hmx2/3a morpholinos (SKD and DKD) display a slight curled-tail-down morphology. Embryos that lacked this morphology (and may therefore not have received any or sufficient morpholino) were also removed before fixing.

For the mRNA + morpholino rescue experiments, we co-injected either each individual or both translation-blocking hmx morpholinos (at the same volume and dose described above), together with a total dose of up to 500 pg of morpholino-resistant (MOR) full-length hmx2 or hmx3a mRNA. Both hmx mRNAs had seven nucleotides altered in the morpholino recognition sequence. Each change was in the third nucleotide of a codon. This codon wobble was used so that the same amino acid was encoded in each case, but the mRNA would not be recognized by the morpholino. The protein encoded by the injected mRNA is therefore the same as either endogenous Hmx2 or Hmx3a.

WT hmx2: ATG AAT AAT TCG GAG GAC AGC (Met, Asn, Asn, Ser, Glu, Asp, Ser)

MOR-hmx2: ATG AAC AAC TCC GAA GAT AGT (Met, Asn, Asn, Ser, Glu, Asp, Ser)

WT hmx3a: ATG CCC GAA ACA ACA CAG GAT ACG (Met, Pro, Glu, Thr, Thr, Gln, Asp, Thr)

MOR-hmx3a: ATG CCG GAG ACT ACT CAA GAC ACC (Met, Pro, Glu, Thr, Thr, Gln, Asp, Thr)

RT-PCR was performed to assess the efficiency of hmx2;hmx3a DKD by splice-blocking morpholinos (Figure S1, A and C). At 27 hpf, separate pools of 25 injected embryos (injected at the one-cell stage with the morpholino dose and volume described above) and 25 uninjected control embryos were homogenized in 200 µl of Tri Reagent Solution (AM9738; Thermo Fisher Scientific). Total RNA was extracted and purified as per the manufacturer’s instructions, before resuspending in 20 µl of sterile distilled water. To remove genomic DNA, 2.4 µl of RQ1 DNase Buffer and 2 µl of RQ1 RNase-Free DNase (M6101; Promega, Madison, WI) was added to each RNA sample and incubated for 15 min at 37°. Heat inactivation of the DNase was performed for 10 min at 65°. 20 µl RT-PCRs were performed as per the manufacturer’s instructions using the Qiagen One-Step RT-PCR kit (210210; Qiagen) and the following primers: hmx2 RT-PCR E1-2 forward: TCAAGTTTCACGATCCAGTCTA and hmx2 RT-PCR E1-2 reverse: ATAAACCTGACTCCGAGAGAAA; hmx3a RT-PCR E1-2 forward: GTCAAAGCCTAAGCCTATTTTG and hmx3a RT-PCR E1-2 reverse: TCACTCTTCTTCCAGTCGTCTA; and actb1 RT-PCR E3-4 forward: GAGGTATCCTGACCCTCAAATA and actb1 RT-PCR E3-4 reverse: TCATCAGGTAGTCTGTCAGGTC (universal PCR program: 50° for 30 min; 95° for 15 min; 35 cycles of 95° for 30 sec, 57° for 45 sec, and 72° for 1 min; followed by a final extension for 10 min at 72°). Parallel reactions, omitting reverse transcriptase and performed on non-DNase-treated samples, were used to verify the nonspliced (genomic) PCR product. 10 µl of each RT-PCR product was assessed by electrophoresing for 40 min at 100 V on a 2% TBE agarose gel. The hmx2 RT-PCR E1-2 primers generate either a 1204 bp genomic (unspliced) or 426 bp spliced product. The hmx3a RT-PCR E1-2 primers generate either a 779 bp genomic (unspliced) or 393 bp spliced product. The actb1 RT-PCR E3-4 primers generate either a 697 bp genomic (unspliced) or 387 bp spliced product (Figure S1, A–D).

To assess whether genetic compensation occurs in either hmx2SU39 or hmx3aSU42 mutants, which lack obvious phenotypes, or hmx3aSU3 mutants, which have milder spinal cord phenotypes than hmx2;hmx3a DKD embryos, we injected the same dose of either hmx2 + p53 MOs (hmx2SU39) or hmx3a + p53 MOs (hmx3aSU42, hmx3aSU3) as described above, into the single cell of one-cell stage embryos generated from incrosses of heterozygous hmx2SU39, hmx3aSU42, or hmx3aSU3 parents, respectively. If genetic compensation is occurring, the upregulated compensating gene(s) will not be knocked down by the hmx morpholino and the phenotype of homozygous mutants should be unchanged. In contrast, WT and heterozygous animals, which contain at least one WT copy of the respective hmx gene will be susceptible to the hmx morpholino and should exhibit stronger, morphant-like phenotypes. For these experiments, while we removed any embryos with severely abnormal morphology, we did not remove embryos that lacked the curled-tail-down morphology, in case these were morpholino-resistant mutant embryos. After fixing, we performed an in situ hybridization for the glutamatergic marker, slc17a6a/b. We visually inspected the embryos on a dissecting microscope and categorized them as either the stronger, morphant-like phenotype (large reduction in the number of slc17a6a/b-expressing cells) or a more subtle phenotype (WT-like in the case of hmx2SU39 and hmx3aSU42, or a smaller reduction in the number of slc17a6a/b-expressing cells in the case of hmx3aSU3 embryos). Embryos within each class were then genotyped as described in the CRISPR mutagenesis section above.

Genotyping

We isolated DNA for genotyping from both anesthetized adult fish and fixed embryos via fin biopsy or head dissections, respectively. For assaying ear phenotypes, we dissected tail tips instead. We genotyped the hmx CRISPR mutants as described above. For mib1ta52b and hmx3asa23054 mutants, we used KASP assays designed by LGC Biosearch Technologies. KASP assays use allele-specific PCR primers, which differentially bind the fluorescent dyes that we quantified with a Bio-Rad CFX96 real-time PCR machine to distinguish genotypes. The proprietary primers used were mib_ta52b and hmx3a_sa23054. Heads or tail tips of fixed embryos were dissected in 70% glycerol/30% distilled water with insect pins. Embryo trunks were stored in 70% glycerol/30% distilled water at 4° for later analysis. For all experiments except phalloidin-staining experiments, DNA was extracted via the HotSHOT method (Truett et al. 2000) using 10 μl of 50 mM NaOH and 1 μl of 1M Tris-HCl (pH 7.4). For phalloidin-staining experiments, the tail up until the end of the yolk extension was dissected in 70% glycerol/30% distilled water as described above and transferred to PBS with 0.1% Tween-20 (PBST). The PBST was then replaced with 50 µl of DNA extraction buffer [10 mM Tris, pH 8.0, 10 mM EDTA, 200 mM NaCl, 0.5% SDS, 200 µg/ml Proteinase K (Proteinase K, recombinant, PCR grade, 3115879001; Sigma Aldrich, St. Louis, MO)], before incubating for 3 hours in a 55° water bath. The samples were vortexed periodically to ensure thorough digestion of the tissue. Subsequently, the Proteinase K was inactivated by heating the samples for 10 min at 100°, before centrifuging for 20 min at 14,000 rpm at room temperature to pellet debris. The supernatant was transferred to sterile microcentrifuge tubes before adding 20 µg UltraPure Glycogen (10814010; Thermo Fisher Scientific) and 2 volumes of ice-cold RNase-free ethanol. Samples were precipitated at −20° overnight. Genomic DNA was recovered by centrifugation at 4°, followed by washing with 70% RNase-free ethanol and further centrifugation at 4°. After carefully removing the ethanolic supernatant, the pellets were air dried for 5–10 min at room temperature before resuspending in 15 µl of sterile distilled water.

In situ hybridization and immunohistochemistry

We fixed embryos in 4% paraformaldehyde/PBS and performed single and double in situ hybridizations and immunohistochemistry plus in situ hybridization double-labeling experiments as previously described (Concordet et al. 1996; Batista et al. 2008). Sources of in situ hybridization probes are provided in Table S1. To amplify in situ hybridization probe templates for hmx1 and hmx3b, we created cDNA from 27 hpf WT zebrafish embryos. We extracted total RNA by homogenizing 50–100 mg of embryos in 1 ml of TRIzol reagent (15596-026; Ambion). We confirmed RNA integrity (2:1 ratio of 28S:18S ribosomal RNA bands) and quality (A260/A280 ratio of ∼2.0) using agarose gel electrophoresis and spectrophotometry, respectively. We synthesized cDNA using Bio-Rad iScript Reverse Transcription Supermix kit (170-8891; Bio-Rad). We amplified hmx1 sequence from the cDNA using Phusion High-Fidelity DNA Polymerase (M0530L; New England BioLabs Inc.), primers hmx1-forward: CTGGTATATTTGCTCAAGACATGC and hmx1-reverse: GCTTCTGCTGAACACAGTTCG, and PCR conditions 98.0° for 10 sec; 30 cycles of 98.0° for 60 sec, 57.0° for 30 sec, and 72.0° for 30 sec; followed by a final extension for 45 sec at 72.0°. The PCR product was assessed on a 1% TAE gel, before purifying with QIAquick PCR Purification Kit (28104; QIAGEN). We used Taq DNA Polymerase (M0320S; New England BioLabs Inc.) to add 3′A overhangs before TOPO TA-cloning (K4600-01; Invitrogen, Carlsbad, CA). We then performed colony PCR using the same PCR primers and conditions used to amplify the hmx1 sequence from cDNA. We extracted plasmid DNA from positive colonies using QIAprep Spin Miniprep Kit (27104; QIAGEN) and then verified the sequence using standard SP6 and T7 primers for Sanger sequencing. To make the antisense RNA riboprobe, we linearized DNA with HindIII-HF (R3104S; New England BioLabs Inc.) and transcribed with T7 RNA Polymerase (10881767001; Roche). We used a PCR-based DNA template to make the hmx3b ISH probe. The reverse primer contains the T3 RNA Polymerase minimal promoter sequence (underlined). We used primers hmx3b-forward: GTGTGCCCGTCATCTACCAC and hmx3b-reverse: AATTAACCCTCACTAAAGGGATGAAGATGATGAAGATGCGCAAC, 27 hpf WT cDNA, Phusion High-Fidelity DNA Polymerase (M0530L, New England BioLabs Inc.) and PCR conditions: 94.0° for 3 min; 35 cycles of 94.0° for 30 sec, 56.5° for 30 sec, and 72.0° for 1.5 min; followed by a final extension step of 72.0° for 10 min. We purified the template through phenol:chloroform:isoamyl alcohol extraction and precipitation with 0.2 M NaCl and ice-cold ethanol before in situ probe synthesis using 1 µg purified PCR product, T3 RNA Polymerase (11031171001; Roche), and DIG RNA Labeling Mix (11277073910; Roche).

Embryos older than 24 hpf were usually incubated in 0.003% 1-phenyl-2-thiourea to prevent pigment formation. For some experiments (indicated in the results) we added 5% dextran sulfate to the hybridization buffer. Dextran sulfate can increase specific staining in in situ hybridization experiments as it facilitates molecular crowding (Ku et al. 2004; Lauter et al. 2011).

In cases where we did not detect expression of a particular gene in the spinal cord, we checked for low levels of expression by exposing embryos to prolonged staining. In some cases, this produced higher background (diffuse, nonspecific staining), especially in the hindbrain, where ventricles can sometimes trap antisense riboprobes.

To determine neurotransmitter phenotypes, we used probes for genes that encode proteins that transport or synthesize specific neurotransmitters, as these are some of the most specific molecular markers of these cell fates [Higashijima et al. (2004b,c) and references therein]. A mixture of probes to slc17a6a and slc17a6b (previously called vglut), which encode glutamate transporters, was used to label glutamatergic neurons (Higashijima et al. 2004b,c). GABAergic neurons were labeled using probes to gad1b (probes previously called gad67a and gad67b) (Higashijima et al. 2004b,c). The gad1b gene encodes for a glutamic acid decarboxylase, which is necessary for the synthesis of GABA from glutamate. A mixture of probes (glyt2a and glyt2b) for slc6a5 (previously called glyt2) was used to label glycinergic cells (Higashijima et al. 2004b,c). slc6a5 encodes for a glycine transporter necessary for glycine reuptake and transport across the plasma membrane.

The antibodies that we used for fluorescence in situ hybridization were mouse anti-Dig (200-002-156; 1:5000; Jackson ImmunoResearch) and rabbit anti-Flu (A889; 1:2500; Invitrogen). These were detected using secondary antibodies goat anti-rabbit-HRP (G-21234; 1:750; Thermo Fisher Scientific) and goat anti-mouse-HRP (G-21040, 1:750; Thermo Fisher Scientific), and Tyramide SuperBoost Kits B40922 and B40915 (Thermo Fisher Scientific).

For double-fluorescence in situ hybridization and immunohistochemistry, after detection of the in situ hybridization reaction using Tyramide SuperBoost Kit B40915 (with HRP, goat anti-mouse IgG and Alexa Fluor 594 Tyramide), embryos were washed eight times for 15 min in PBST and incubated in Image-iT FX Signal Enhancer (I36933; Thermo Fisher Scientific) for 30 min at room temperature. Immunohistochemistry was performed using chicken polyclonal anti-GFP primary antibody (Ab13970; 1:500; Abcam) and a goat anti-chicken IgY (H+L), Alexa Fluor 488 secondary antibody (A-11039; 1:1000; Thermo Fisher Scientific).

Phalloidin staining

Four-day-old embryos generated from incrosses of heterozygous hmx2SU39 or hmx2;hmx3aSU44 parents were fixed and processed for phalloidin staining as described in Hartwell et al. (2019). Stained embryos were stored in DABCO [2% w/v 1,4-Diazabicyclo[2.2.2]octane (D27802; Sigma Aldrich) in 80% glycerol in sterile distilled water].

Quantitative PCR analyses

We collected embryos from incrosses of AB WT parents and flash-froze them at 16-cell, 6, 14, 27, and 48 hpf stages. We collected 40–50 embryos per biological replicate per developmental stage and performed duplicate biological replicates. We isolated total RNA by homogenizing each sample in 1 ml of TRIzol reagent (15596-026; Ambion). Following chloroform extraction, we added 20 µg UltraPure Glycogen (10814010; Thermo Fisher Scientific) to the aqueous phase followed by one volume of RNase-free ethanol. We performed RNA purification and genomic DNA removal using the Monarch Total RNA Miniprep Kit (T2010S; New England BioLabs Inc.), following manufacturer’s instructions for purifying TRIzol-extracted samples. RNA concentration was measured using Nanodrop 2000 (ND2000; Thermo Fisher Scientific), before synthesizing cDNA using the Bio-Rad iScript Reverse Transcription Supermix kit (170-8891; Bio-Rad). We also included controls lacking reverse-transcriptase to assay for the presence of genomic DNA contamination. Quantitative PCR (qPCR) was performed in triplicate for each sample using iTaq Universal SYBR Green Supermix (1725121; Bio-Rad) and a Bio-Rad CFX96 real-time PCR machine. The following qPCR primers were used: hmx2-qPCR-forward: CCCATTTCAAGTTTCACGATCCAGTC and hmx2-qPCR-reverse: TGCTCCTCTTTGTAATCCGGTAG; hmx3a-qPCR-forward: TTGATGGCAGCTTCTCCCTTTC and hmx3a-qPCR-reverse: ACTCTTCTTCCAGTCGTCTATGC; and mob4-qPCR-forward: CACCCGTTTCGTGATGAAGTACAA and mob4-qPCR-reverse: GTTAAGCAGGATTTACAATGGAG.

The hmx2 and hmx3a primers were generated in this study. The mob4 primers were generated by Hu et al. (2016). They demonstrated that mob4 is a more effective reference gene than actb2 across a broad range of zebrafish developmental stages, including early stages where only maternal mRNAs should be present (Hu et al. 2016). To generate amplification data the program used was 95.0° for 30 sec; 40 cycles of 95.0° for 5 sec and 63.3° (hmx2)/64.5° (hmx3a)/60.0° (mob4) for 30 sec; with imaging after each cycle. To assay amplification specificity and exclude false positives from primer dimers we then generated melt data using 65.0° for 30 sec; 40 cycles of 65.0°–95.0°, +0.5°/second increment, with each increment held for 5 sec before imaging; 95.0° for 15 sec.

Screening lateral line and otolith phenotypes

We examined whether any of the hmx mutants generated in this study had lateral line and/or fused otolith phenotypes, as reported for hmx2;hmx3a DKD embryos (Feng and Xu 2010). To assay live lateral line phenotypes, we anesthetized embryos from incrosses of heterozygous mutant fish in 0.016% tricaine (A5040; Sigma Aldrich) in embryo medium [5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2.2H2O, 0.33 mM MgSO4.7H2O, 0.017% w/v (0.7 mM) HEPES pH 7.8 and 0.00004% methylene blue in autoclaved reverse osmosis water] and mounted them on coverslip bridges [2 × 22 mm square glass coverslips (16004-094; VWR) glued together on either side of a 24 × 60 mm glass cover slip (12460S; Thermo Fisher Scientific), overlaid with a third 22 mm square glass coverslip]. Using a Zeiss Axio Imager M1 compound microscope, we located the tip of the lateral line primordium and counted the somite number adjacent to this position. We also used this method routinely to determine the developmental stage of embryos before fixing for in situ hybridization. To assay lateral line phenotypes in fixed embryos, we performed in situ hybridizations for hmx3a or krt15 (both of which label the migrating primordium and neuromasts) and then determined the lateral line position as in live embryos. To examine live otolith phenotypes, embryos were raised until 3 d, before anesthetizing (as for assessing live lateral line phenotypes) and examining the spatial location of otoliths in both ears. WT embryos have two otoliths in each ear: one smaller, anterior (utricular) otolith, and one larger, posterior (saccular) otolith. These are separate from each other and spatially distinct. We classified otoliths as fused if only one large, amalgamated otolith was visible in a midventral position within the otic vesicle.

Imaging

Embryos were mounted in 70% glycerol:30% distilled water and differential interference contrast (DIC) pictures were taken using an AxioCam MRc5 camera mounted on a Zeiss Axio Imager M1 compound microscope. Fluorescence images were taken on a Zeiss LSM 710 confocal microscope. Images were processed using Adobe Photoshop software (Adobe, Inc) and Image J software (Abramoff et al. 2004). In some cases, different focal planes were merged to show labeled cells at different medio-lateral positions in the spinal cord. All images were processed for brightness, contrast and color balance using Adobe Photoshop software (Adobe, Inc.). Images of control and mutant embryos from the same experiment were processed identically. Figures were assembled using Adobe Photoshop and Adobe Illustrator (Adobe, Inc.).

Cell counts and statistics

In all cases except where noted to the contrary, cell counts are for both sides of a 5-somite length of spinal cord adjacent to somites 6–10. Embryos were mounted laterally with the somite boundaries on each side of the embryo exactly aligned and the apex of the somite over the middle of the notochord. This ensures that the spinal cord is straight along its dorsal-ventral axis and that cells in the same dorsal/ventral position on opposite sides of the spinal cord will be directly above and below each other. Embryos from mutant crosses were counted blind to genotype. Labeled cells in embryos analyzed by DIC were counted while examining embryos on a Zeiss Axio Imager M1 compound microscope. We identified somites 6–10 in each embryo and counted the number of labeled cells in that stretch of the spinal cord. We adjusted the focal plane as we examined the embryo to count cells at all medio-lateral positions (both sides of the spinal cord; Batista et al. 2008; Batista and Lewis 2008; England et al. 2011; Hilinski et al. 2016; Juárez-Morales et al. 2016).

In some cases, cell count data were pooled from different experiments. Before pooling, all pairwise combinations of data sets were tested to determine if there were any statistically significant differences between them, as described below. Data were only pooled if none of the pairwise comparisons were statistically significantly different from each other. In addition, as in situ hybridization staining can vary slightly between experiments, we only compared different mutant results when the counts from their corresponding WT sibling embryos were not statistically significantly different from each other.

To determine whether differences in values are statistically significant, data were first analyzed for normality using the Shapiro–Wilk test. Data sets with nonnormal distributions were subsequently analyzed using the Wilcoxon–Mann–Whitney test (also called the Mann–Whitney U-test). For data sets with normal distributions, the F-test for equal variances was performed, before conducting either a type 2 (for equal variances) or type 3 (for unequal variances) Student’s t-test. P-values generated by Wilcoxon–Mann–Whitney, type 2 Student’s t-test and type 3 student’s t-test are indicated by ^, +, and §, respectively. To control for type 1 errors, when comparing three or more experimental conditions, a one-way ANOVA test was performed. Before conducting ANOVA tests, data were first analyzed for normality using the Shapiro–Wilk test, as described above. All data sets for ANOVA analysis had normal distributions and so were subsequently assessed for homogeneity of variances using Bartlett’s test. All of the data sets also had homogeneous (homoscedastic, Bartlett’s test P > 0.05) variances and so standard ANOVA analysis was performed. ANOVA results are reported as F(dfn,dfd) = F-ratio, P-value = x, where F is the F-statistic, dfn is the degree of freedom for the numerator of the F-ratio, dfd is the degree of freedom for the denominator of the R-ratio, and x is the P-value. For statistically significant ANOVA, to determine which specific experimental groups or groups differed, post hoc testing was performed. Since all ANOVA data sets had homogeneous (homoscedastic) variances, Tukey’s honestly significant difference post hoc test for multiple comparisons was performed. P-values generated by Tukey’s honestly significant difference test are indicated by ‡. Data are depicted as individual value plots and the n-values for each experimental group are also shown. For each plot, the wider red horizontal bar depicts the mean and the red vertical bar depicts the SEM (SEM values are listed in Tables 1 and 2). Individual data value plots were generated using Prism version 8.4.3 (GraphPad Software, San Diego, California; www.graphpad.com). To assess whether mutant phenotypes occurred at Mendelian frequencies, we performed chi-squared tests. To test whether a small number of embryos with abnormal phenotypes was statistically significantly different from zero we performed a binomial distribution test, using the cumulative distribution function, the number of embryos without mutant phenotypes, the total number of embryos examined (n) and a probability argument of n − 1/n. P-values > 0.05 support the null hypothesis that the number of embryos with abnormal phenotypes is not statistically significantly different from zero. Shapiro–Wilk and Wilcoxon–Mann–Whitney testing was performed in R version 3.5.1 (R Development Core Team 2005). The F-test, Student’s t-test, chi-squared test, and binomial distribution test were performed in Microsoft Excel version 16.41. Bartlett’s testing, standard ANOVA, and Tukey’s honestly significant difference testing were performed in Prism version 8.4.3 (GraphPad Software).

Table 1

Statistical comparisons of numbers of cells expressing particular genes in morpholino knockdown experiments

Table 2

Statistical comparisons of numbers of cells expressing particular genes in mutant experiments

Data and reagent availability

Plasmids and zebrafish strains are available upon request. Supplemental material available at figshare: https://doi.org/10.25386/genetics.13108325. Figure S1 contains RT-PCR and cell-count data demonstrating the efficacy of hmx2;hmx3a DKD with splice-blocking morpholinos. Figure S2 shows an alignment of mouse and zebrafish Hmx2 and Hmx3(a) protein sequences. Table S1 includes gene names, ZFIN identifiers, and references for in situ hybridization probes. Table S2 contains the sgRNA and primer sequences used for hmx2, hmx3a, and hmx2;hmx3a CRISPR mutagenesis and genotyping. Microarray data have been previously deposited in the NCBI Gene Expression Omnibus under accession number GSE145916.

Resultshmx2 and hmx3a are the only hmx genes expressed in the spinal cord

While the expression and functions of zebrafish hmx genes have been analyzed during the development of sensory structures such as the eye and the ear, the expression of hmx1, hmx2, hmx3a, and hmx4 in the developing spinal cord has not been investigated and no expression data has previously been reported for hmx3b, which only appeared in more recent versions of the zebrafish genome sequence (Zv9 and above). Therefore, to determine which of the hmx genes are expressed in the spinal cord we performed in situ hybridizations for hmx1, hmx2, hmx3a, hmx3b, and hmx4 at different developmental stages (Figure 1). At all of these stages, we observed no spinal cord expression of hmx1, hmx3b, or hmx4 (Figure 1). However, consistent with previous reports, both hmx1 and hmx4 were expressed in the developing eye, ear, and anterior lateral line neuromasts (Figure 1; French et al. 2007; Feng and Xu 2010; Gongal et al. 2011; Boisset and Schorderet 2012; Marcelli et al. 2014). In contrast, the only expression of hmx3b that we observed was weak hindbrain expression at later stages of development (36–48 hpf; Figure 1, S’, X’, and AC’).