Ethical statement

All animal experimental procedures were approved by the local ethics committee for animal research, Uppsala djurförsöksetiska nämnd (permit numbers C161/4 and 5.8.18-18096/2019). All procedures for the experiments were performed in accordance with the Swedish Board of Agriculture’s Regulations and General Advice about Laboratory Animals.

CRISPR/Cas9 target design

Two sgRNAs targeting the single zebrafish nkx3.2 gene with no predicted off-target effects were designed using the online software CHOPCHOP [29], both targeting the first exon: 5´-GATCAGGAATCCGCGGCCAA-3´ and 5´-GTCGTTGTCCTCGCTCAGCC-3´. The second base of each target was modified to "G" in order to allow T7 transcription without modifications. The sgRNAs were prepared as previously described [30], creating a fragment consisting of the T7 promotor, the targeted gene-specific sequence, and the guide core sequence. The sgRNAs were synthesised by in vitro transcription using the HiScribe T7 High Yield RNA Synthesis Kit (New England Biolabs, Ipswich, MA). Cas9 mRNA was prepared by in vitro transcription with the mMESSAGE mMACHINE T3 Transcription Kit (Life Technologies, Carlsbad, CA) using 500 ng of linearised plasmid that was retrieved from 5 μg of p-T3TS-nCas9n plasmid (plasmid #46757; Addgene, Cambridge, MA) digested with XbaI (New England Biolabs, Ipswich, MA). The gRNAs were purified with mirVana miRNA isolation kit (Life technologies, Carlsbad, CA) and Cas9 mRNA was purified with RNeasy Mini Kit (Qiagen, Maryland; MD), and their integrity was assessed using a denaturation gel.

Generation of zebrafish mutant line

Fertilised zebrafish (Danio rerio) eggs were obtained by natural spawning of Tg(sox10:egfp) line [31]. Embryos were injected at the one-cell stage with 150 pg of Cas9 mRNA and 50 pg of each sgRNA in RNase-free water as previously described [30], and maintained at 28.5°C in E3 medium [32]. The efficiency of the targets was estimated by the CRISPR-Somatic Tissue Activity Test (STAT) methodology in eight embryos at two days post-injection, as previously described [33]. The injected founder zebrafish (F0) were raised and incrossed. For genotyping the F1 zebrafish, DNA was extracted from a 1–3 mm amputation of the adult zebrafish caudal fin by lysing the tissue in 30 μl of 50 mM NaOH for 20 min at 95°C, adding 60 μl of 0.1 mM Tris and diluting the obtained material (1:10). For the initial genotyping step, FLA analysis was used. 2 μl of DNA (50–200 ng) was added to Platinum Taq DNA Polymerase. The PCR mix was incubated at 94°C for 12 min followed by 35 cycles of 94°C for 30 sec, 57°C for 30 sec, 72°C for 30 sec, and final extension at 72°C for 10 min. Size determination was carried out on a 3130XL ABI Genetic Analyzer (Applied Biosystems, Waltham, MA) and the data were analysed using the Peak Scanner Software (Thermo Fisher Scientific, Waltham, MA). For the fish that screened positive for the variant, the FLA results were confirmed by Sanger sequencing.

Founder screening and identification of heterozygous adult fish

sgRNA for both targets were co-injected and we have genotyped the fish for potential mutations at both target sites. One strain with an allele containing a frameshift deletion resulting in a premature stop codon (nkx3.2^uu2803) was selected for further experiments. The identified F1 founders were crossed with wild-type zebrafish (AB strain), and their adult offspring (F2) were genotyped. Heterozygous F2 fish of mutant line nkx3.2^uu2803 were incrossed and the offspring was observed with bright-field and fluorescence microscopy. Embryos showing phenotypes were euthanised with an overdose of MS-222 (300mg/L) and genotyped by FLA and/or Sanger sequencing (forward primer: 5’-TGTAAAAC-GACGGCCAGTGAGGAGTCTCGCCATCTGAA-3’; reverse primer: 5’-GTGTCTTACAGATGAA-GCTTTGAGTGGT-3’).

In vivo microscopy

Fluorescent images were obtained with an inverted Leica TCS SP5 confocal microscope using LAS-AF software (Leica Microsystems). Embryos were sedated with 0.16% MS-222 and embedded in 0.8% low melting agarose onto the glass bottom of the 35mm dishes. To prevent drying, embedded embryos were covered with system water containing 0.16% MS-222. Screening for GFP in zebrafish larvae was performed using a Leica M205FCA fluorescence microscope with the appropriate filter.

Histological analysis

Zebrafish juveniles and adults at 14, 30 and 90 dpf were euthanised with an overdose of MS-222 (300mg/L), fixed in 4% PFA and washed in PBST buffer. 30 and 90 dpf fish were decalcified in 0.5 M EDTA for one week with EDTA exchange every third day. Fish were transferred into 99.5% ethanol, followed by Xylene and embedded into paraffin. Sagittal head sections of 6μm were prepared with Leica RM2155 Rotary Microtome. Tissue sections were deparaffinised with xylene and re-hydrated through 99.5% to 70% ethanol series and transferred to water. Sections were then stained with Nuclear Fast Red (Vector Laboratories, Burlingame, CA) for 30 sec followed by a brief water rinse and dehydration in 95% and 99.9% ethanol. Prior to mounting with VectaMount (Vector Laboratories, Burlingame, CA), slides were washed in Clear-Rite 3 (Richard-Allan Scientific, Kalamazoo, MI) three times for three minutes each. Sections were imaged with 40x objective on Hamamatsu NanoZoomer S60 Digital Slide Scanner.

Skeletal staining

Staining of cartilage and bone was done based on the previously published protocol by Walker and Kimmel [34]. Zebrafish wild-type and mutant fish at 5, 9, 14 and 30 dpf, were euthanised with an overdose of MS-222 (300mg/L), fixed in 4% PFA and transferred to 50% ethanol. For cartilage staining, specimens were immersed in alcian blue solution (0.02% Alcian Blue 8 GX, 50mM MgCL2, 70% ethanol), and for bone staining, specimens were immersed in alizarin red solution (0.5% Alizarin Red S). For double staining of cartilage and bone, specimens were immersed in double staining solution (99% alcian blue solution, 1% alizarin red solution). After staining overnight, specimens were washed twice with 50% ethanol and then immersed in water for 2 hours before being bleached in a solution of 1.5% H202 and 1% KOH until pigmentation was removed. 30 dpf specimens were then immersed in trypsin solution (1% trypsin, 35% sodium tetraborate) for 30 minutes followed by incubation in a solution of 10% glycerol and 0.5% KOH for 1 hour. All specimens were imaged with a Leica M205FCA microscope in a solution of 50% glycerol and 0.25% KOH, followed by storage in 50% glycerol and 0.1% KOH.

Optical projection tomography

A custom-built Optical Projection Tomography (OPT) system was used for imaging of the zebrafish embryos at 5 dpf that were euthanised with an overdose of MS-222 (300mg/L), fixed and stained with alcian blue [35,36]. The OPT system, reconstruction algorithms, and alignment workflow were based on the previously described method [37]. All embryos were kept in 99% glycerol before they were loaded into the system for imaging. The rotational images were acquired using a 3X telecentric objective with a pixel resolution of 1.15 μm/pixel. The tomographic 3D reconstruction was done using a filtered back projection (FBP) algorithm in MATLAB (Release R2015b; MathWorks, Natick, MA) together with the ASTRA Toolbox [38]. For the data alignment, the registration toolbox elastix [39,40] was used. To reduce the computational time all 3D volumes in the registration were down-sampled to half the resolution.

The registration workflow was similar to the methods described in ref. [37] where the wildtype fish were initially aligned and used to create an average reference fish using an Iterative Shape Averaging (ISA) algorithm [41]. All wild-type (n = 10) and nkx3.2^{uu2803/uu2803} (n = 11) zebrafish were then aligned to the reference. After the alignment, a voxel-wise method was used to detect voxels that are significantly different between the groups. The Mann-Whitney U test was used to compare corresponding voxels in wild-type and mutant. The p-value threshold is set using a false discovery rate (FDR) [42] and a permutation test [43]. The FDR was set so that those random groupings showed only a small number of significant voxel differences (p<2.5x10^-4; FDR = 0.045). All registration and analysis were done on the green channel of the RGB images.

Micro-computed tomography and segmentation

Five wild-type and five nkx3.2^{uu2803/uu2803} zebrafish at 90 dpf were euthanised with an overdose of MS-222 (300mg/L), fixed, and analysed with micro-computed tomography (μCT, SkyScan 1172, Bruker microCT, Belgium) at a voltage of 60 kV, a current of 167 μA, and an isotropic voxel size of 5.43 μm. Cross-sections were reconstructed using software package NRecon (NRecon 1.6.10, Bruker microCT, Belgium). The specimens were placed in 2mL Eppendorf tubes filled with 1% agarose and the 10ul pipette tip was used to keep the mouths of some wild-type fish in the open position. BMP image stacks obtained with μCT were imported into, segmented, and imaged using VGStudio MAX version 3.2.5 (Volume Graphics, Germany).

Zebrafish nkx3.2^{uu2803/uu2803} line generated with CRISPR/Cas9 survives to adulthood and has open mouth phenotype

Clustered Regulatory Interspaced Short Palindromic (CRISPR)/CRISPR-associated protein 9 (Cas9) was used to generate nkx3.2 mutant line to analyse both embryonic and adult mutant phenotypes. Both of the designed sgRNAs showed similar activity and during the genotyping step we selected founder fish with a frameshift mutation caused by one of the sgRNAs (Fig 1A). We generated a mutant allele with a 7 bp deletion in nkx3.2 exon 1 (c.286_292 del, p.Lys95*), causing a frameshift, which resulted in a premature stop codon that shortens the peptide sequence to 95 amino acids, compared to 245 amino acids in the wild-type (Fig 1A). The last eight amino acids encoded by the first exon and all amino acids encoded by the second exon including the entire DNA-binding homeodomain of Nkx3.2 are predicted to be absent in this shortened protein and these changes generate a severe loss-of-function allele.

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Fig 1. Zebrafish nkx3.2 null allele generated with CRISPR/Cas9.

(A) To-scale schematic of the two-exon zebrafish nkx3.2 gene locus on chromosome 14. Grey boxes represent the coding sequence, the black box labelled “HD” marks the location of the homeodomain-coding sequence in the second exon, and the white box marks the 1,226bp 3’ UTR, truncated for illustrative purposes. gRNA target site is marked by turquoise background. The sequence of the zoomed in section of exon 1 shows the alignment between the wild-type and mutant (uu2803) alleles. The mutant allele has a 7 base-pair deletion that results in a frameshift and a premature stop codon (highlighted in red) immediately after the deletion site. (B) Wild-type, heterozygous nkx3.2^uu2803/+ and homozygous nkx3.2^{uu2803/uu2803} fish were genotyped by fragment length analysis. The wild-type displayed one peak (297,15), nkx3.2^uu2803/+ fish one wild-type (297,12) and one mutant (290,86) peak and nkx3.2^{uu2803/uu2803} fish one mutant peak (290,53). (C) nkx3.2^{uu2803/uu2803} fish at 90 dpf display a prominent fixed open mouth phenotype compared to wild-type at 90 dpf. Scale bars: 1 cm.

https://doi.org/10.1371/journal.pone.0255953.g001

Heterozygous embryos (nkx3.2^uu2803/+) displayed no gross morphological differences compared to wild-type (nkx3.2^+/+) embryos as long as observed. Homozygous mutant embryos displaying morphological differences were generated by crossing two heterozygous nkx3.2^uu2803/+ adult zebrafish. Homozygous mutants (nkx3.2^{uu2803/uu2803}) were able to survive to adulthood and displayed a prominent fixed open mouth phenotype (n = 6/6, Fig 1C). Adult homozygous mutants (further in the text referred to as mutants) at 90 dpf displayed no significant differences in standard length (SL) relative to adult wild-types, although they showed a rather wide distribution of SL (S1 Fig).

The differences in SL between wild-type and mutant groups were apparent at 30 dpf and 60 dpf. Possible explanations are that bigger SL in mutants at 30 dpf was due to overeating caused by constant swimming through food with an open mouth, whereas smaller SL at 60 dpf was due to some competition for food before separation of the mutant group at 60 dpf (S1 Fig). However, other explanations such as differences in tank densities or cryptic genetic variation are also plausible, and we cannot rule them out.

Confocal live imaging displays fusion of Meckel’s cartilage with palatoquadrate in nkx3.2^{uu2803/uu2803} zebrafish embryo and larvae

In order to follow the first mandibular arch development, confocal live imaging was performed using a transgenic sox10:egfp line labelling neural crest-derived cells of the pharyngeal arches. Clear phenotypic differences in the jaw joint-forming region were detectable from 3 dpf onwards (Fig 2, S1 Movie). The jaw joint was lost in nkx3.2^{uu2803/uu2803} fish as Meckel’s cartilage and the palatoquadrate were fused from 3 dpf onwards (n = 4/4, Fig 2E–2H”) and the retroarticular process (RAP) was missing at the posteroventral tip of Meckel’s cartilage (n = 4/4, Fig 2E–2H”, S1 Movie). nkx3.2 mutants furthermore showed an unorganised cell-mass of small rounded chondrocytes posterior to the jaw joint fusion site, in the anterior part of the palatoquadrate at 3–5 dpf (n = 4/4, Fig 2E–2F”). By 7dpf chondrocyte of the anterior palatoquadrate were elongated and displayed typical stacking (n = 4/4, Fig 2G–2G”, S1 Movie). At 14 dpf, all chondrocytes were elongated and displayed alignment throughout the entire Meckel’s–palatoquadrate fused element (n = 3/3, Fig 2H–2H”, S1 Movie).

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Fig 2. Larval development of wild-type and nkx3.2^{uu2803/uu2803} jaw joints.

(A-H’) Maximum projections of confocal live imaging Z-stacks acquired from ventral side of wild-type zebrafish head at 3 dpf (A, A’), 5 dpf (B, B’), 7dpf (C, C’), 14 dpf (D, D’) and nkx3.2^{uu2803/uu2803} zebrafish head at 3 dpf (E, E’), 5 dpf (F, F’), 7 dpf (G, G’), 14 dpf (H, H’) in Tg(sox10:egfp) background. Corresponding single Z-planes are shown in A”–H”. (A-D´´) The jaw joint in wild-type zebrafish (dashed box) is magnified in A’-D”. The retroarticular process is visible from 3 dpf onwards, marked by white arrowhead. Chondrocytes of the anterior Meckel’s cartilage and posterior palatoquadrate align in stacks. Posterior Meckel’s cartilage and anterior palatoquadrate articulate the jaw joint. (E-H”) Fusion of jaw joint articulating elements in nkx3.2 mutants. The fusion site is magnified in (E’-H”) and indicated by asterisks. (E’-F’ and E”-F”) At 3 dpf and 5 dpf nkx3.2 mutants display unorganised and rounded cells in the anterior palatoquadrate, posterior to the jaw joint fusion site (G’, G”) At 7 dpf nkx3.2 mutants display elongated chondrocytes that start to align in stacks. (H’, H”) At 14 dpf nkx3.2 mutants display elongated chondrocytes that are aligned throughout the fused Meckel’s-palatoquadrate element. m—Meckel’s cartilage, pq—palatoquadrate. Scale bars: 100 μm (A-H), 25 μm in (A’-H’ and A”-H”).

https://doi.org/10.1371/journal.pone.0255953.g002

Histological analysis of nkx3.2^{uu2803/uu2803} zebrafish shows chondrocyte alignment and hypertrophy at the jaw joint fusion site

Histological sections were prepared to further analyse the chondrocyte arrangement within the first mandibular arch in nkx3.2 mutant larval, juvenile, and adult zebrafish. A joint gap between Meckel´s cartilage and the palatoquadrate is visible in wild-type zebrafish at 14 dpf (Fig 3A and 3A’). In wild-types at 14 dpf, articular chondrocytes line the joint cavity and hypertrophic chondrocytes form the articulating elements (Fig 3A and 3A’). In nkx3.2 mutant larvae at 14 dpf, Meckel´s cartilage and the palatoquadrate were not separated but fused–the jaw joint was absent (Fig 3B and 3B’). Chondrocytes within the fused element were hypertrophic and aligned. At the presumptive fusion site, the element appeared to be increased in width caused by piled-up rows of aligned chondrocytes (Fig 3B and 3B’). The exact fusion point was difficult to determine in both 30 dpf and 90 dpf mutants as we did not see any clear border between articular and quadrate (n = 7/7; Fig 3D, 3D’, 3F and 3F’), but apart from the fusion and its phenotypic consequences, ossification seemed not to be affected. These results are different from Miyashita et al. [9] who showed a clear boundary between articular and quadrate bones. However, we cannot exclude that a very faint border could exist in our mutants, but was not contrasted with Nuclear Red staining. Articular chondrocytes lining the articulating tips of the articular and quadrate were consequently absent in nkx3.2 mutant juvenile fish at 30 dpf (Fig 3D and 3D’). Chondrocytes within the fused element displayed hypertrophic morphology similar to wild-type (Fig 3C–3D’). By 90 dpf, ossification of the fused articular and quadrate was completed, visible by the presence of adipose tissue inside the bones in both wild-type and mutant adult fish (Fig 3E–3F’).

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Fig 3. Histological analysis reveals loss of jaw joint without affected chondrocyte hypertrophy in the first pharyngeal arch of nkx3.2^{uu2803/uu2803} zebrafish.

Sagittal sections of wild-type 14 dpf (A, A’), 30 dpf (C, C’) and 90 dpf (E, E’) and nkx3.2 mutant 14 dpf (B, B’) 30 dpf (D, D’) and 90 dpf (F, F’) zebrafish stained with Nuclear Red. (A, A’, C, C’, E, E’) Stained histology sections displaying a normal jaw joint development between Meckel’s cartilage (m) and palatoquadrate (pq), respectively articular (art) and quadrate (q) in wild-type fish. (B, B’, D, D’, F, F’) nkx3.2 mutants do not display a jaw joint. Chondrocyte maturation and ossification seem not to be affected in nkx3.2^{uu2803/uu2803} besides the absence of joint-typical cells lining the articulating elements. Dashed box in (A-F) marks the magnified region in (A’-F’). Asterisk marks the fused element. m–Meckel’s cartilage, pq–palatoquadrate, art–articular, q–quadrate. Scale bars: 200μm (A-F), 100μm (A’-F’).

https://doi.org/10.1371/journal.pone.0255953.g003

Optical projection tomography reveals morphological changes in the head of nkx3.2^{uu2803/uu2803} larvae

In order to characterise the larval nkx3.2 mutant phenotype in greater detail, we used optical projection tomography (OPT) on 5 dpf cartilage-stained wild-type (n = 10) and mutant (n = 11) larvae to reconstruct 3D models of cartilage morphology. Multiple wild-type (n = 10) and mutant (n = 11) 3D models were overlaid and combined to produce an average wild-type morphology (Fig 4A, 4D and 4G) and an average nkx3.2 mutant morphology (Fig 4B, 4E and 4H), respectively. Overlaying these grouped 3D reconstructions allowed the calculation of voxels that were statistically more intense or less intense in the mutant group (Fig 4C, 4F and 4I), corresponding to locations with more or less cartilage present, respectively. This analysis was robust to false positives (S2 Fig).

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Fig 4. Optical projection tomography of cartilage-stained wild-type and nkx3.2^{uu2803/uu2803} larvae reveals subtle phenotypic changes.

(A, D, G) Maximum projection of 5 dpf wild-type group (n = 10). (B, E, H) Maximum projection of 5 dpf nkx3.2 mutant (n = 11) group. (C, F, I) Maximum projection of both groups with coloured voxels representing voxels with statistically significant (p<2.5x10^-4) differences in intensity. Magenta shows voxels with higher intensity and cyan shows voxels with lower intensity in nkx3.2 mutant group. Asterisk marks the fusion site. ch–ceratohyal, m–Meckel’s cartilage, ac–auditory capsule, pq–palatoquadrate, rap–retroarticular process. Scale bars: 150μm.

https://doi.org/10.1371/journal.pone.0255953.g004

The nkx3.2 mutant group showed significantly higher intensity of the cartilage labelling at the jaw joint region (magenta in Fig 4F and 4I) consistent with the presence of fused cartilage in mutants compared to the jaw joint gap in wild-type larvae. Cyan voxels ventral to the jaw joint in Fig 4F indicate the absence of the RAP. The palatoquadrate displayed significantly higher intensity in nkx3.2 mutant group consistent with the increased thickness of this element (magenta in Fig 4C, 4F and 4I). The anterior part of Meckel’s cartilage displayed an increased posterior intensity and decreased anterior intensity in the nkx3.2 mutant group indicating a change in the shape and increased thickness of Meckel’s cartilage (Fig 4I). Interestingly, this analysis also revealed significantly reduced cartilage staining signal in the posterior part of the head, around the auditory capsule (Fig 4F and 4I). The mutant group clearly showed the shorter parts of the posterior auditory capsule changing the shape of the notochord insertion area, compared to the wild-type group (Fig 4F and 4I).

Cartilage and bone staining analysis of wild-type and nkx3.2^{uu2803/uu2803} larval, juvenile and adult zebrafish

Skeletal staining of larval, juvenile and adult nkx3.2 mutant zebrafish was performed to analyse both cartilage and bone abnormalities in comparison to wild type at a greater range of developmental stages (Fig 5). The loss of the jaw joint caused by the fusion of Meckel´s cartilage and palatoquadrate was clearly visible from 5 dpf onwards and was most recognizable by the absence of the RAP (Fig 5B, 5D and 5F). The resulting open mouth phenotype could be observed from 9 dpf onwards (Fig 5D, 5F and 5H). No other cartilage phenotypes within the pharyngeal arches could be detected at any developmental stage, consistent with the OPT results at 5 dpf in Fig 4. Ossification of the dentary, articular, and quadrate appeared to take place normally in nkx3.2 mutants at 30 dpf (n = 2/2, Fig 5H and 5H’) when compared to wild-type fish (Fig 5G and 5G’).

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Fig 5. Skeletal staining of wild-type and nkx3.2^{uu2803/uu2803} zebrafish.

(A-F) lateral views of cartilage-stained wild-types and nkx3.2 mutants at 5, 9 and 14 dpf. (G-H) lateral views of cartilage- and bone-stained wild-types and nkx3.2 mutants at 30 dpf. Boxes in (A-H) indicate the zoomed-in regions in the insets or zoomed-in panels (G’, G”, H’, H”). The measurements in mm refer to standard length (SL). Asterisks indicate the fusion between Meckel’s cartilage and palatoquadrate in the jaw joint. jj–jaw joint, O–occipital region, bvc–basiventral cartilage, WA–Weberian apparatus. Scale bars: 2mm (A-H), 200μm (A’-H’ and G”-H”).

https://doi.org/10.1371/journal.pone.0255953.g005

μCT reveals craniofacial phenotypes in adult nkx3.2^{uu2803/uu2803} zebrafish

In order to examine the skeleton of adult zebrafish we performed μCT followed by 3D reconstruction of wild-type and nkx3.2 mutant zebrafish at 90 dpf (Fig 6). The resting position of the mouth is closed (CM) in wild-type individuals (Fig 6A, 6C and 6F), so to be better able to compare the phenotype of the wild-type to the fixed open mouth phenotype of the mutant, μCT scans were also performed on 90 dpf zebrafish with their mouths held open (AOM in Fig 6D and 6G) with a pipette tip which produces an unnatural but reasonable approximation of the appearance of the zebrafish head skeleton with an open mouth.

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Fig 6. μCT reveals craniofacial phenotypes in adult wild-type and nkx3.2^{uu2803/uu2803} zebrafish.

(A, B) lateral view of wild-type and nkx3.2 mutant zebrafish at 90 dpf. (C, D, E) lateral view of the head of wild-type with closed mouth (CM), wild-type with artificially open mouth (AOM), and nkx3.2 mutant. (F, G, H) Anterior view of wild-type with CM, wild-type with AOM, and nkx3.2 mutant. (I, J, K) ventral view of isolated wild-type dentary (CM), wild-type dentary (AOM), and nkx3.2 mutant dentary. (L, M, N) Virtual thin sections (60μm thick, averaged) through the symphysis of the dentaries displayed in I, J, K, respectively. The measurements in mm refer to standard length (SL). Asterisks in (B, E) indicate the fixed open mouth phenotype caused by fused jaw joint. The arrowhead in (E) indicates the downturned ceratohyal phenotype relative to (D). Angles in (I, J) show the flexibility of the symphysis in wild-types, the arrow in K and N indicates the thickened and deformed symphysis phenotype in nkx3.2 mutants. Colour scheme: Red–lower jaw; dark blue–infraorbitals; dark green–ceratohyal and anal fin; blue–basihyal; lilac–branchial arches; dark purple–supraoccipital; lime green–exoccipital and basioccipital; yellow–all remaining craniofacial bones; cyan–cleithrum and pectoral fins, arctic blue–Weberian apparatus; blue violet–vertebrae; green–pelvic fins; bronze–dorsal fin; orange–caudal fin vertebrae and hypurals; magenta–caudal fin rays. AA–anguloarticular, AF–anal fin, BH–basihyal, CF–caudal fin, CH–ceratohyal, DF–dorsal fin, DT–dentary, io3 –infraorbital 3, jj–jaw joint, le–lateral ethmoid, mx–maxilla, O–occipital region, PCF–pectoral fin, pmx–premaxilla, PVF–pelvic fin, QT–quadrate, WA–Weberian apparatus. Scale bars: 5mm (A, B), 1mm (C-N).

https://doi.org/10.1371/journal.pone.0255953.g006

The fusion of the Meckel’s and palatoquadrate cartilages early in the development of nkx3.2 mutants effectively contributed to a fusion of the articular and quadrate that ossify from these cartilage precursors. This made it extremely challenging to demarcate the articular and quadrate in the nkx3.2 mutants at 90 dpf, which is why the lower jaw was not segmented in red in Fig 6B, 6E, 6H and 6K. As in younger nkx3.2 mutants (Fig 5D, 5F and 5H), the mouth of 90 dpf mutants was fixed in an open position, and this likely resulted in forces being exerted on the basihyal and ceratohyal by the interconnecting muscles during juvenile development. The outcome of these forces varied between different individuals. Fig 6E shows an individual where the ventral position of the lower jaw appeared to have pushed the basihyal and ceratohyal posteroventrally, resulting in a sharp angle between the anteroventrally-pointing anterior end of the ceratohyal and the posteroventrally-pointing posterior end of the basihyal. Other individuals (S3A Fig) had a relatively normal position of the basihyal and ceratohyal in the mouth compared to the ventrally-positioned lower jaw, resulting in the basihyal partially obstructing the open mouth, its anterior end positioned dorsally to the entire lower jaw. In some mutants (n = 2/5) the basihyal was bent half-way and ossified in an L-shape (S3B Fig).

Viewed anteriorly (Fig 6F–6H), the face of the mutant appeared “pinched” at the position where the jaw joint would have formed, resulting in a reduced area of the mouth opening (n = 5/5). There were also impacts on the bones of the upper jaw, the premaxilla and maxilla, which also appeared to be posteriorly compressed into the cranium and compressed laterally in line with the “pinched” jaw apparatus. As recently described mutant line was reported to lack kinethmoid [9], we segmented the corresponding region and found that the kinethmoid is present in our mutants at both 60 and 90 dpf, and relatively morphologically unchanged as compared to wild-type fish (n = 9/9, S4 Fig).

The cartilaginous symphysis joint between the paired bones of the dentary flexes during feeding in wild-type zebrafish as the jaw is opened and the width of the mouth opening is increased by the lateral flaring of the suspensorium [44,45], illustrated in Fig 6I, 6J, 6L and 6M. In contrast, the symphysis of nkx3.2 mutants tended to be thickened and deformed (n = 5/5, Fig 6K and 6N) possibly leading to reduced flexibility.

Juvenile and adult nkx3.2^{uu2803/uu2803} zebrafish display loss of basiventral cartilage and parapophyses

Next, we assessed the effect of nkx3.2 loss-of-function on the axial skeleton of juveniles and adults. Mutants at 8.7 mm SL displayed a loss of basiventral cartilage in the precaudal rib-bearing vertebrae (Fig 7B) compared to wild-types of 8.8 mm SL (Fig 7A). Wild-type fish at 10.5 mm SL also lacked these cartilages (Fig 7C), as they have been entirely replaced through endochondral ossification by the parapophyses, small articulating bones that connect the ribs to the vertebrae. However, in 30–35 dpf nkx3.2 mutants (8.7–9.4 mm SL), it was clear that this endochondral ossification of basiventral cartilage had not taken place, as the parapophyses were absent (n = 4/4, Fig 7B and 7D). Instead, the ribs were fused directly to the vertebrae without any articulating process, while some other (more posterior) ribs were entirely disconnected from the vertebrae. This phenotype persisted in 90 dpf nkx3.2 mutants, as seen in μCT segmented models and virtual histological sections of the vertebrae (Fig 7E–7H). In contrast, the neural arches, zygapophyses, and haemal arches appeared to develop normally (n = 4/4).

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Fig 7. Parapophyses are absent in the rib-bearing vertebrae of nkx3.2^{uu2803/uu2803} zebrafish.

(A-F) Dorsolateral views of rib-bearing vertebrae in 30–35 dpf, and 90 dpf wild-type and nkx3.2 mutant zebrafish. (A-D) Cartilage- and bone- stained juvenile zebrafish, (E, F) μCT models. (G, H) μCT virtual transverse cross-sections of wild-type and nkx3.2 mutant rib-bearing vertebrae at 90 dpf. The measurements in mm refer to standard length (SL). Arrowheads in (B, D, F) indicate the absence of parapophyses on the rib-bearing vertebrae. Parapophyses (pop) are highlighted in green in (E). c–centrum, r–rib. Scale bars: 200μm(A-D), 500μm (E-H).

https://doi.org/10.1371/journal.pone.0255953.g007

OPT, skeletal staining, and μCT reveal changes in the occiput and Weberian apparatus of nkx3.2^{uu2803/uu2803} zebrafish

Comparison of OPT reconstructions of 5 dpf larvae (Fig 4) suggested the reduction of the cartilage of the auditory capsule changing the shape of the notochord insertion area in nkx3.2 mutants. At 30 dpf cartilage and bone staining revealed changes in the occipital region and cervical vertebrae in nkx3.2 mutant compared to wild-type zebrafish. More specifically, nkx3.2 mutants displayed smaller lateral occipital fenestrae, due to changes in the shape of the exoccipital and supraoccipital as well as reduced basiventral cartilages and cartilages associated with forming Weberian apparatus (Fig 5G” and 5H”).

To investigate these phenotypes in greater detail, these bones were segmented from μCT scans of 90 dpf wild-types and nkx3.2 mutants. In the occipital region at the back of the skull, mutants displayed a dramatic fusion between the basioccipital and exoccipital (n = 5/5, wild-type is shown in Fig 8A, 8C, 8E and 8G), making it impossible to clearly demarcate these two elements, hence they are the same colour in mutants displayed in Fig 8B, 8D, 8F and 8H. The medial gap between the paired exoccipital bones (Fig 8E) was also partially or completely fused in mutants (Fig 8F). As a result of this fusion and the fusion between the exoccipital and basioccipital, the posterodorsal opening of the cavum sinus impar (csi) was lost or greatly deformed (Fig 8G and 8H). The anteroventral surface of the basioccipital that contacts the parasphenoid and prootic was highly convex in mutants, compared to only slightly convex in wild-types (Figs 8C, 8D and S5). The posterodorsal exoccipital struts were impacted by the cervical vertebrae in nkx3.2 mutants (n = 5/5, Fig 8B), causing the lateral occipital fenestrae to be reduced in area and rotated posteromedially.

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Fig 8. μCT reveals phenotypes in the occiput and Weberian apparatus of nkx3.2^{uu2803/uu2803} zebrafish.

Lateral, ventral, anterior, and posterior views of wild-type (A, C, E, G) and homozygous mutant (B, D, F, H) occiput and Weberian apparatus. Cervical vertebrae (CV) 2–4 have been removed in (G, H) for clarity. Asterisk in B, D indicates the absence or severe reduction of lateral process 2 from CV2. Arrowhead in B, D indicates the absence of anterior ramus of tripus on CV3. Arrow in D indicates the V-shaped anteroventral edge of basioccipital. Asterisk in F indicates the posterior fusion of the medial gap between the paired exoccipital bones. Dotted line in G highlights the cavum sinus impar (csi), while the asterisk in H indicates its absence. The measurements in mm refer to standard length (SL). AO–asteriscus otolith, BO–basioccipital, csi—cavum sinus impar, cl–claustrum, EO–exoccipital, lof–lateral occipital fenestrae, lp–lateral process, na–neural arch, os–os suspensorium, r4 –rib 4, sc–scaphium, sn2 –supraneural 2, SO–supraoccipital, tr–tripus. Scale bars: 500μm.

https://doi.org/10.1371/journal.pone.0255953.g008

Wild-type zebrafish possess four distinct cervical vertebrae (CV), while most mutants (n = 4/5) lacked the first and most anterior CV (Fig 8A and 8B), with one mutant also missing CV2. Dorsal to CV1 in wild-type zebrafish are the scaphium and claustrum, which were entirely lost or highly reduced in all mutants lacking CV1 (Fig 8A, 8B, 8G and 8H). The lateral process (lp2) on CV2 was significantly reduced in mutants, varying between individuals from a ~50% reduction in size to almost complete loss (Fig 8C and 8D). The tripus of CV3 was deformed in mutants—the anterior ramus and articulating process were absent (Fig 8A–8D). The articulation of rib 4 and the os suspensorium to CV4 was also altered in mutants, consistent with the absence of parapophyses observed in the rib-bearing vertebrae of vertebrae 5–11 (Fig 7E and 7F).