This article is mentioned in:

Introduction

The neural crest (NC) is a multipotent cell population and is one of the key features of craniofacial development (1). Neural crest cells (NCCs) originate from the dorsal neural tube and generate numerous cell types, including cartilage and pigment cells (2). Defective development of NC has been demonstrated to result in a number of syndromes, such as Treacher Collins and DiGeorge syndrome, which are characterized by developmental defects in the craniofacial region (3-5). In humans, craniofacial malformations have been reported to be associated with 75% of all congenital birth malformations (6) and are accompanied by abnormal brain development and poor prognosis.

MicroRNAs (miRNAs/miRs) have been demonstrated to regulate ~30% of genes in Caenorhabditis elegans, mouse and human and serve an important role in embryonic development (7). The Homo sapiens (hsa)-miR-1 family includes two miRNAs, hsa-miR-1-1 and hsa-miR-1-2, which exhibit the same mature sequence, although they are encoded by different genes (8). Previous studies have indicated that miR-1 participates in regulating the development of the heart and skeletal muscle (9,10). The outflow track of the heart, as well as the tendons, muscles and craniofacial structures, have all been reported to be derivatives of the NC (1). miR-1 has been revealed to be conserved and important for embryonic development in zebrafish, especially NC development (11). However, the specific role of miR-1 in NCCs is currently unknown.

Zebrafish exhibit rapid embryonic development, a short life cycle and large spawning capacity. In addition, the embryos are transparent, so the process of embryonic skeletal development can be observed directly (12). Therefore, zebrafish are a suitable animal model to study the function of genes in craniofacial development. In the present study, the miR-1-knockout zebrafish model was a homozygous mutant, which was used to examine the role of miR-1 in NCCs.

The current study aimed to evaluate the role of miR-1 in the development of NC using a miR-1 deficient zebrafish model.

Materials and methods

Animals

The wild-type (WT) zebrafish embryos were of the Tuebingen strain. WT and homozygous miR-1 mutant zebrafish (n=60 per group) were purchased from the Institute of Model Animals of Nanjing University. The primers used for genotyping of the miR-1 mutant zebrafish, the sequence of the mutant allele and the identification of the genotype are presented in Tables SI and SII and Fig. S1. The embryos were grown in Holtfreter's solution (0.05 g/l KCL, 0.1 g/l CaCl2, 0.2 g/l NaHCO3 and 3.5 g/l Nacl) with a 14 h light/10 h dark cycle at 28.5˚C in the zebrafish facility of the Model Animal Research Center, Nanjing University (Nanjing, China). The zebrafish were fed three times a day with pellet food, and their health and behavior was monitored daily. All experiments were approved by the Ethics Committee of the School of Stomatology of Nanjing Medical University, and all procedures were performed according to the guidelines of the Animal Care Committee of Nanjing Medical University.

Whole mount in situ hybridization

Whole mount in situ hybridization was performed using standard procedures with slight modifications as previously described (13). Probes were synthesized using a DIG RNA labeling kit (Roche Diagnostics). The probes of forkhead box D3 (foxd3), muscle segment homeobox-b (msxb), snail homolog 1b (snai1b), transcription factor AP2a (tfap2a), distal-less 3 (dlx3b), neurog1 (ngn1) (Table SIII) were designed using Primer 5.0 software (Premier Biotech, Inc.). Embryos (n=5 per group) at 24 h post-fertilization (hpf) were collected, fixed in 4% paraformaldehyde overnight at 4˚C and subjected to gradient dehydration with methanol (25, 50, 75 and 100%). The embryos were maintained in 100% methanol at -20˚C for subsequent use. Following rehydration at gradient dilutions using methanol/PBS, the embryos were treated with proteinase K (10 µg/ml; Gibco; Thermo Fisher Scientific, Inc.) at room temperature for 10 min. Subsequently, embryos were prehybridized with the hybridization solution at 65˚C for 2 h. In situ hybridization signals were detected by sheep anti-digoxigenin-AP Fab-fragments (Roche Diagnostics) and the color reaction was carried out by staining with BCIP/NBT (Roche Diagnostics) as previously described (14).

Alcian blue staining

Zebrafish embryos at 96 hpf (n=5 per group) were fixed in 4% paraformaldehyde overnight at 4˚C, dehydrated with ethanol (50 and 100%) and stained with Alcian blue (0.15 mg/ml; Sigma-Aldrich; Merck KGaA) overnight at room temperature (15). After the embryonic cartilage was stained, the embryos were incubated in saturated boron overnight at room temperature. Subsequently, bleaching was performed in a mixture of 3% H2O2 and 1% KOH at a ratio of 15:85 v/v, and the specimens were digested in 1% trypsin at 37˚C until all embryonic tissues were degraded. The digestion mixture was replaced with 1% KOH solution, and subsequently the specimens were incubated in a glycerol gradient (25, 50, 75 and 100%). The embryos were observed using a stereomicroscope (magnification, x20).

Histology and immunostaining

For whole mount immunostaining, the embryos (n=5 per group) were fixed in 4% paraformaldehyde overnight at 4˚C pretreated with proteinase K (10 µg/ml) at 37˚C for 30 min and blocked in 2% BSA solution (BioFroxx) at room temperature for 1 h. Subsequently, the embryos were stained with an anti-phospho-histone H3 (PHH3; 1:100; cat. no. 3377; Cell Signaling Technology, Inc.) antibody overnight at 4˚C and then with an anti-rabbit Alexa Fluor@594 (1:400; Jackson ImmunoResearch Laboratories, Inc.) at 4˚C for 2 h. A TUNEL assay was performed using an In Situ Cell Death Detection kit (Roche Diagnostics) as previously described (16). The staining was subsequently observed using laser scanning confocal microscopy (magnification, x20) (17).

Culture of NCCs

Mouse embryos were harvested at gestational day E9.5 according to a previously published protocol (18). The pregnant adult mice (n=3; Model Animal Research Center, Yangzhou University) were sacrificed by cervical dislocation following anesthesia with sodium pentobarbital (50 mg/kg). The first branchial arch of the embryos was separated, washed in PBS and dissociated in 0.25% trypsin-EDTA (Gibco; Thermo Fisher Scientific, Inc.) for 30 min at 37˚C. The separated cells were plated into a dish in Nutrient Mixture F-12 (1:1)media (Gibco; Thermo Fisher Scientific, Inc.) with 15% FBS (ScienCell Research Laboratories, Inc.), supplemented with 100 U/ml penicillin-streptomycin (PS), 100 mg/ml L-glutamate, 0.1 mM minimum essential medium non-essential amino acids (Gibco; Thermo Fisher Scientific, Inc.) and 1,000 U/ml leukemia inhibitory factor (PeproTech, Inc.). The inhibitor (5'-AUA CAUACUUCUUUACAUUCCA-3'; 100 nM) and mimics (forward, 5'-UGGAAUGUAAAGAAGUAUGUAU-3'; and reverse, 5'-AUACAUACUUCUUUACAUUCCA-3'; 50 nM) of miR-1 and their negative controls (NCs; inhibitor NC, 5'-CAG UACUUUUGUGUAGUACAAA-3'; mimics NC, 5'-UUU GUACUACACAAAAGUACUG-3'; 5'-CAGUACUUUUG UGUAGUACAAA-3'; all Guangzhou RiboBio Co., Ltd.) were transfected into NCCs when cells grew to 50% density using a riboFECT™ CP Transfection kit (Guangzhou RiboBio Co., Ltd.) according to the manufacturer's protocol.

Cell Counting Kit-8 (CCK-8) assay

CCK-8 assay (Dojindo Molecular Technologies, Inc.) was used to evaluate cell proliferation according to the manufacturer's protocol. NCCs were seeded at 2x103 cells/well in 96-well plates. The cells were incubated at 37˚C for 12 h (day 0) and transfected with the miR-1 inhibitor. CCK-8 reagent in Nutrient Mixture F-12 media was added to each well at the specified time points (days 0, 1, 2, 3 and 4). Following 1-h incubation at 37˚C, the optical density was measured at 450 nm using a microplate reader with three blank controls.

Wound healing assay

NCCs were seeded at 1.5x105 cells/well in a 6-well plate. Following transfection with the miR-1 inhibitor, a wound was created with a 10 µl pipette tip, and the cells were incubated in serum-free medium at 37˚C for 24 h. Cell migration was observed at 0, 12 and 24 h using a Leica DMIRE2 microscope (magnification, x100; Leica Microsystems GmbH) in phase contrast mode (19).

Cell cycle and apoptosis analysis

NCCs transfected with the miR-1 inhibitor were cultured for 24 h without PS. Following an additional 48-h culture, cell cycle and apoptosis analyses were performed as previously described (20) using flow cytometry (FACSCalibur; BD Biosciences). For cell cycle analysis, the cells (~1x104) were fixed overnight with 75% ethanol at 4˚C in the dark, and then stained with propidium iodide (PI; MultiSciences Biotech Co., Ltd.) containing RNase. Apoptosis was detected by harvesting cells using a centrifuge at room temperature (5 min, 210 x g) and staining the cells (~1x104) with an Annexin V-FITC Apoptosis Detection kit (BD Pharmingen; BD Biosciences).

PI-/Annexin V-FITC+ cells (early apoptotic population) and PI+/Annexin V-FITC+ cells (late apoptotic population) were sorted. Analysis was performed using FACScan cytometer (BD Biosciences) and FlowJo V7 software (Tree Star).

Apoptotic cell staining

NCCs were seeded at 6x104 cells/well in a 24-well plate, transfected with the miR-1 inhibitor and then cultured for 3 days at 37˚C. Acridine orange/ethidium bromide (AO/EB) staining was performed using an AO/EB Detection kit (Biotechgate) according to the manufacturer's instructions. After fixing in 4% paraformaldehyde at 4˚C for 15 min, TUNEL staining of NCCs was performed using an In Situ Cell Death Detection kit (Roche Diagnostics) according to the manufacturer's instructions. The cells were observed using fluorescence microscopy (magnification, x400; Leica Microsystems, Inc.).

Reverse transcription-quantitative PCR (RT-qPCR)

Total NCCs RNA was isolated using TRIzol® reagent (Invitrogen; Thermo Fisher Scientific, Inc.), as previously described (21). The miRNA expression levels of miR-1 relative to the internal control U6 were evaluated after reverse transcription (37˚C for 60 min and 85˚C for 5 min) using an All-in-One™ miRNA RT-qPCR Detection kit (GeneCopoeia, Inc.), the thermocycling conditions were set according to the manufacturer's instructions. The miR-1 (5'-CGCGTGGAATGTAAAGAA GTATGTA-3') and U6 (5'-AAAACAGCAATATGGAGC GC-3') forward primers and universal reverse primers (cat. no. QP015) were purchased from GeneCopoeia, Inc. Relative expression levels were calculated using the 2-ΔΔCq method (22).

Western blot analysis

Proteins of cells were extracted using RIPA buffer (Beyotime Institute of Biotechnology), boiled for 5 min and analyzed its protein concentrations using a Bicinchoninic Acid Protein Assay Kit (Beyotime Institute of Biotechnology). The proteins were separated using 10% or 12% SDS-PAGE as previously described (19), transferred onto PVDF membranes and then blocked with 5% BSA at room temperature for 2 h. Primary antibodies against BCL-2, poly ADP-ribose polymerase (PARP), pro- and cleaved-caspase-9 (1:500; cat. nos. 12789, 13371 and 10380, respectively; Wuhan Sanying Biotechnology), pro- and cleaved-caspase-3 (1:1,000; cat. no. 9661S; Cell Signaling Technology, Inc.) and β-actin (1:1,000; cat. no. sc-47778; Santa Cruz Biotechnology, Inc.) were used overnight at 4˚C. Next, the membranes were incubated with HRP-labeled Goat Anti-Rabbit or HRP-labeled Goat Anti-mouse secondary antibody (1:8,000, cat. nos. 474-1506 and 474-1806, respectively; SeraCare KPL) at room temperature for 1 h and visualized by enhanced chemiluminescence (Thermo Fisher Scientific, Inc.). Semi-quantitative analysis was performed using ImageJ 1.52 software (National Institutes of Health).

Statistical analysis

All data are presented as the mean ± standard deviation. Differences between two groups were analyzed using GraphPad Prism v6 statistical software (GraphPad Software Inc.). Unpaired Student's t-test was used for the statistical analysis. P<0.05 was considered to indicate a statistically significant difference.

Results

Loss of miR-1 in zebrafish results in craniofacial, pigment cell and cardiac defects

In the present study, a miR-1 mutant zebrafish model was generated to evaluate the role of miR-1 in the development of NC. The expression of miR-1 was primarily localized in the head, pharyngeal arch and pectoral fin of zebrafish (Fig. 1A). In the miR-1 mutant zebrafish, the expression of miR-1 was suppressed in the aforementioned areas (Fig. 1B).

Figure 1 Loss of miR-1 in zebrafish results in craniofacial, pigment cell and cardiac defects. In situ hybridization of miR-1 in (A) wild-type and (B) mutant zebrafish at 24 hpf. (C) Zebrafish larvae at 72 hpf with mutants exhibiting mandibular retrognathia (left arrow) and edema around the heart (right arrow). (D) Alcian blue staining at 96 hpf. (E) Quantitative analysis of b/a. (F) Quantitative analysis of c/a. a, width between ch; b, distance between mc and ch; and c, the distance between pq and ch. (G) Quantitative analysis of head size. (H) Distribution and number of iridophores at 72 hpf. Data are presented as the mean ± standard deviation (n=5). *P<0.05 and **P<0.01 vs. WT. WT, wild-type; hpf, hours post-fertilization; miR-1, microRNA-1; mc, Meckel's cartilage; pq, palatoquadrate; ch, ceratohyal cartilage.

A number of abnormal phenotypes were observed in the mutant compared with the WT zebrafish, including a smaller head, a decreased mandibular arch length and mandibular retrognathia (Fig. 1C-G). In the mutant zebrafish, the number of melanocytes and iridophores (NCC-derived pigment cells) was decreased (Fig. 1C and H). In addition, the mutant zebrafish exhibited abnormal cardiac development with enlarged hearts compared with those of the WT zebrafish (Fig. 1C). A quantitative analysis of the association among the palatal square joint, McGuire cartilage and hyoid small horn cartilage has been previously performed (23). In the present study, the head size and mandibular arch length were decreased in the mutant group compared with those in the WT group (Fig. 1D-G).

These results suggested that the lack of miR-1 affected the development of NCCs in zebrafish, which resulted in severe phenotypic defects.

miR-1 affects NCC apoptosis during the migration and differentiation periods in embryonic development

To determine which stage of NC development was affected by miR-1, the expression of marker genes of specific stages (foxd3, msxb, snai1b, tfap2a, dlx3b and ngn1) was detected via in situ hybridization. These genes have been demonstrated to be associated with the induction, specialization, migration and differentiation of NCCs (24,25).

Foxd3 has been indicated to serve an important role in NC derived tissues, such as the pharyngeal arch (24). In the present study, two regions of NCCs were labelled with the foxd3 probe in WT and mutant zebrafish; however, the mutant group appeared to exhibit lower expression levels of foxd3 compared with that in the WT group (Fig. 2A). This indicated that the migration of NCCs may be suppressed in the miR-1 mutant group. Msxb in zebrafish has also been reported to be required for NCC survival (25). However, msxb expression exhibited no difference between the two groups (Fig. 2B). Therefore, it was speculated that the lack of miR-1 affected the migration, but not the survival of NCCs.

Figure 2 MicroRNA-1 knockout affects neural crest cell apoptosis during the migration and differentiation periods of embryonic development. In situ hybridization of zebrafish at 24 hpf with (A) foxd3, (B) msxb, (C) snai1b, (D) tfap2a, (E) dlx3b and (F) ngn1 probes. (G) Whole mount immunostaining of zebrafish at 24 hpf with PHH3 (red) and DAPI (blue). (H) Quantitative analysis of immunostaining. (I) TUNEL staining at 24 hpf with TUNEL (red) and DAPI (blue). (J) Quantitative analysis of TUNEL staining. Data are presented as the mean ± standard deviation (n=5). *P<0.05 vs. WT. WT, wild-type; foxd3, forkhead box D3; msxb, muscle segment homeobox-B; snai1b, snail homolog 1b; tfap2a, transcription factor AP2A; dlx3b, distal-less 3; ngn1, neurog1; PHH3, phospho-histone H3; hpf, hours post fertilization; ns, not significant.

To examine migration, the expression of snai1b and tfap2a was also examined in the NC area of the head. In situ hybridization indicated that snai1b and tfap2a expression appeared to be decreased in the mutant group at 24 hpf compared with that in the WT group (Fig. 2C and D). These results suggested that migration was inhibited in the mutant zebrafish.

During embryonic development in zebrafish, the first and second pharyngeal arch develop into the mandible and hyoid arches (1). The results of the present study indicated that the expression of dlx3b in miR-1 mutant embryos appeared to be decreased compared with that in the WT group (Fig. 2E). This was consistent with the mutant zebrafish phenotype, namely lower mandibular arch length and mandibular retrognathia, which was observed in the current study. Knockdown of miR-1 appeared to decrease in ngn1 expression compared with that in the WT group (Fig. 2F). This evidence collectively suggested that miR-1 may serve an important role during the migration and differentiation periods of NCCs.

Cell proliferation, which was assessed by immunofluorescence staining of PHH3, exhibited no differences between the two groups (Fig. 2G and H). An assessment of apoptosis in zebrafish embryos at 24 hpf was performed using a whole-mount TUNEL assay. The results demonstrated that miR-1-knockout embryos exhibited an increased level apoptosis compared with that in the WT embryos (Fig. 2I and J). These results demonstrated the importance of miR-1 in regulating apoptosis during NC development.

miR-1 regulates NCC functions in vitro

To examine the function of miR-1 in vitro, miR-1 was knocked down in NCCs isolated from mice using a miR-1 inhibitor (Fig. 3A). Following inhibition of miR-1, the proliferative ability of NCCs was slightly reduced compared with that of the control group at day 3, whereas no significant differences were observed at day 1, 2 and 4 (Fig. 3B). In addition, a wound healing assay indicated that the migratory rate of the miR-1 inhibitor group was decreased compared with the control group (Fig. 3C and D).

Figure 3 miR-1 regulates the function of NCCs in vitro. (A) miR-1 expression assessed via reverse transcription-quantitative PCR following transfection with the miR-1 inhibitor. (B) Cell proliferation of NCCs analyzed by Cell Counting Kit-8 assay. (C) Wound healing assay of NCCs. Scale bar, 100 µm. (D) Quantitative analysis of wound healing assay. (E) Cell cycle analysis of NCCs performed using flow cytometry. (F) Quantitative analysis of cell cycle analysis. (G) Apoptosis analysis of NCCs evaluated via flow cytometry. (H) Quantitative analysis of apoptosis analysis. Data are presented as the mean ± standard deviation (n=3). *P<0.05, **P<0.01 and ***P<0.001 vs. Control. NCCs, neural crest cells; miR-1, microRNA-1; PI, propidium iodide.

Subsequently, cell cycle and apoptosis analysis was performed via flow cytometry. The results of the cell cycle analysis suggested that knockdown of miR-1 increased the number of cells at the G2/M phase compared with that in the control group, indicating that cell cycle was arrested at G2/M phase. By contrast, the numbers of NCCs at the G0/G1 and S phases were reduced compared with those in the control group (Fig. 3E and F). In addition, following transfection with the miR-1 inhibitor, the number of early and late apoptotic cells was increased compared with that in the control group (Fig. 3G and H). These in vitro results demonstrated that miR-1 may be crucial for apoptosis in NCCs and were consistent with the in vivo observations.

miR-1 affects NCC apoptosis via the mitochondrial apoptosis pathway and caspase-3

Early and late apoptosis in mouse NCCs was detected via AO/EB staining. The results demonstrated that the number of early and late apoptotic cells was increased in the miR-1 inhibitor group compared with that in the control group (Fig. 4A and B). In addition, TUNEL staining demonstrated that compared with the control group, an increased number of apoptotic cells was observed three days after transfection with miR-1 inhibitor (Fig. 4C and D).

Figure 4 miR-1 affects the apoptosis of neural crest cells via the mitochondrial apoptosis pathway and caspase-3. (A) Detection of apoptosis on day 3 following miR-1 inhibitor transfection using AO/EB staining. Scale bar, 100 µm. (B) Quantitative analysis of AO/EB staining. (C) TUNEL staining of NCCs. Scale bar, 100 µm. (D) Quantitative analysis of TUNEL staining. (E) Protein levels of the mitochondrial apoptosis pathway markers BCL-2, pro- and cleaved-caspase-3, pro- and cleaved-caspase-9 and PARP detected via western blotting. (F-K) Quantification of the protein levels of (F) BCL-2, (G) pro-caspase-9, (H) cleaved-caspase-9, (I) pro-caspase-3, (J) cleaved-caspase-3 and (K) PARP, which are presented in (E). (L) miR-1 expression assessed by reverse transcription-quantitative PCR. (M) Expression levels of pro- and cleaved-caspase-3 after transfection with the miR-1 mimic. (N and O) Quantification of the protein levels. Data are presented as the mean ± standard deviation (n=3). *P<0.05, **P<0.01 and ***P<0.001 vs. Control. miR-1, microRNA-1; PARP, poly ADP-ribose polymerase, AO, acridine orange; EB, ethidium bromide.

Alterations in the apoptotic pathway were examined in mouse NCCs by detecting the expression levels of apoptosis-associated proteins using western blotting (Fig. 4E-K). The expression levels of cleaved-caspase-3 and -9 were increased in the miR-1 inhibitor group compared with those in the control group (Fig. 4H and J), whereas the expression levels of pro-caspase-3 were decreased in the miR-1 inhibitor group compared with those in the control group (Fig. 4I). The expression levels of BCL-2, PARP and pro-caspase-9 were also examined. In the miR-1 inhibitor group, the expression levels of PARP were upregulated, while those of BCL-2 and pro-caspase-9 were decreased compared with the control group (Fig. 4F, H and I). Therefore, these results suggested that the mitochondrial apoptosis pathway was activated following miR-1 inhibition. The increased protein expression levels of cleaved caspase-3 in the miR-1 inhibitor group compared with the control group was consistent with the increased apoptosis that was observed in the mutant zebrafish compared with that in the WT group. To further analyze the relationship between miR-1 and caspase-3, NCCs were transfected with a miR-1 mimic (Fig. 4L). The results indicated that the expression level of cleaved-caspase-3 was downregulated in cells transfected with miR-1 mimics compared with that in the control group (Fig. 4M-O), suggesting that miR-1 inhibited the activation of caspase-3 and affected the apoptosis of NCCs.

Collectively, these results suggested that miR-1 regulated the apoptosis of NCCs via the mitochondrial apoptosis pathway.

Discussion

In a previous study, a latitudinal expression of miR-1 was observed in WT zebrafish via in situ hybridization, and it was demonstrated that the expression of miR-1 increased during embryonic development at 24 hpf (11).

In the present study, in situ hybridization was used to detect the expression of miR-1 in WT zebrafish embryos. The expression of miR-1 was demonstrated to be primarily localized in the head, pharyngeal region and pectoral fin of zebrafish at 24 hpf. Previous findings have indicated that the injection of miR-1 morpholino oligomers in zebrafish resulted in impaired craniofacial chondrogenesis, severe maxillofacial malformations and an abnormal function of NCCs within 4 days post-fertilization. However, it was not clear whether these defects were also affected by the injection of the morpholino (11). Therefore, a miR-1 knockout zebrafish model was generated in the present study to elucidate the role of miR-1 in NC development.

NCCs have been reported to contribute to the development of bones, cartilage, cranial ganglia and connective tissues in the face and neck (1). Previous studies have demonstrated that the abnormal expression of genes associated with NC may result in craniofacial, bone and cartilage dysplasia due to abnormal cell migration, proliferation and differentiation (26-29). For example, dlx3b has been associated with the differentiation of the pharyngeal arch in zebrafish (30). Also, Ngn1 has been reported to regulate the differentiation of the dorsal root ganglion in zebrafish development (31). In the current study, in situ hybridization evidence collectively suggested that miR-1 serve an important role during the migration and differentiation periods of NCCs.

miRNAs are a class of highly conserved, noncoding RNA molecules with a length of 18-25 nucleotides. The mature sequence of miR-1 is highly conserved among different vertebrates, including zebrafish, mice and humans, which suggests that miR-1 may exhibit similar functions among vertebrates (32). The role of miR-1 in regulating the development of heart, cartilage and liver, among other organs, has been reported (33-35). In the present study, the following abnormal phenotypes were observed in the miR-1-knockout model: Mandibular retraction, shortening of the inferior dental arch and lingual arch, and atypical neural crest-derived tissues, such as abnormal pericardial edema and pigment cells.

Li et al (36) reported that miR-1 decreased cardiomyocyte apoptosis via mediating the expression of apoptosis-related genes in the heart. In the current study, TUNEL staining revealed an increase in apoptosis of pharyngeal NCCs in miR-1 mutant compared with WT zebrafish, which was consistent with previous reports (36). By contrast, PHH3 immunofluorescence staining did not demonstrate any differences between the WT and mutant groups; additional time points may be required to determine whether alterations in NCC proliferation were caused by miR-1 knockout.

Caspase-3 has been reported to be a key protein in the mitochondrial apoptosis pathway, and RARP is one of the principal targets of caspase-3(37). Also, pro-caspase-9 requires to be activated to subsequently cleave and activate caspase-3(38). Western blotting results suggested that miR-1 may regulate the mitochondrial apoptosis pathway.

The biogenesis of miRNAs is a complex process. After several processing steps (39), mature single-stranded miRNAs have been indicated to enter the miRNA-induced silencing complex, which silences the expression of target genes primarily at the post-transcriptional level (40). The target genes to be silenced are selected via base-pairing interactions between the miRNA and the target mRNA that contains a partial or complete complementary sequence, generally localized in 3'-untranslated region (41). The results of the current study suggested that caspase-3 may be targeted by miR-1. However, the regulatory pathways and mechanisms of this interaction require additional studies.

In conclusion, the present study demonstrated an important role of miR-1 in regulating the apoptosis of NCCs during embryonic development via modulating the mitochondrial apoptosis pathway. A normal expression of miR-1 was identified to be essential for the development of NC derivatives, including the pharyngeal cartilage, mandible and hyoid bone. A thorough understanding of molecular pathway by which miR-1 regulates its target genes may aid the prevention and treatment of miR-1-associated developmental malformations.

Supplementary Material

Genotypic identification of mutant zebrafish via PCR amplification and subsequent electrophoresis. Each pair of two adjacent lanes (from left to right) represents the PCR identification results of one specimen. The first lane indicates the amplification product of the primer pair Mir1-1-11bp wild-type F1/Mir1-1 R, and the second lane represents the amplification product of the primer pair Mir1-1-11bp mutant F1/Mir1-1 R. The results indicated that all specimens were homozygous mutants. F, forward; R, reverse; Mir1, microRNA-1.

Genotype of microRNA-1 knockout mutant in zebrafish.

Primers used in the identification of genotype by PCR.

The sense and antisense probes used in whole mount in situ hybridization.

Acknowledgements

Not applicable.

Funding

Funding: The current study was funded by the National Natural Science Foundation of China (grant no. 81771029), the Natural Science Fund for Colleges and Universities in Jiangsu Province, China (grant no. 18KJA320004), the Southeast University-Nanjing Medical University Cooperative Research Project (grant no. JX218GSP20180705) and the Priority Academic Program Development of Jiangsu Higher Education Institutions (grant no. 2014-037).

Availability of data and materials

All data generated or analyzed during this study are included in this published article.

Authors' contributions

NZ, WQ, DW, AGR and LY performed the experiments and analyzed the data. NZ, YM, CM, ZX and JM designed the study. NZ, AGR, ZX and JM wrote the manuscript. All authors read and approved the final manuscript.

Ethics approval and consent to participate

The present study was approved by the Ethics Committee of the School of Stomatology of Nanjing Medical University (approval no. 1403049), and all procedures were performed according to the guidelines of the Animal Care Committee of Nanjing Medical University.

Patient consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

References